1

UGWU, ONYEDIKACHI DANIEL

PG/M.Sc./2010/58003

HYDROLYSIS OF CASSAVA STARCH BY MALTED ACHA (DIGITARIA

EXILIS) FOR BIOETHANOL PRODUCTION

FACULTY OF BIOLOGICAL SCIENCES

DEPARTMENT OF BIOCHEMISTRY

Ameh Joseph Jnr

Digitally Signed by: Content manager’s Name

DN : CN = Webmaster’s name

O= University of Nigeria, Nsukka

OU = Innovation Centre

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TITLE

HYDROLYSIS OF CASSAVA STARCH BY MALTED ACHA

(DIGITARIA EXILIS) FOR BIOETHANOL PRODUCTION

A PROJECT REPORT SUBMITTED IN PARTIAL

FULFILMENT OF THE REQUIREMENTS FOR AWARD OF

DEGREE OF MASTER OF SCIENCE (M. Sc) IN INDUSTRIAL

BIOCHEMISTRY & BIOTECHNOLOGY,

UNIVERSITY OF NIGERIA,

NSUKKA

BY

UGWU, ONYEDIKACHI DANIEL

(PG/M.Sc/2010/58003)

DEPARTMENT OF BIOCHEMISTRY

UNIVERSITY OF NIGERIA

NSUKKA

SUPERVISOR: DR. S.O.O EZE

FEBRUARY, 2013

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CHAPTER ONE

INTRODUCTION

Energy and environmental issues are the major concerns facing the global community today (Hu

et al., 2008). Renewable fuels (biofuels) such as bioethanol are becoming increasingly important

due to heightened concern for the green house effect, depleting oil reserves and rising oil prices

(Ohgren et al., 2007). Ethanol, chemically known as ethyl alcohol, is a clear, colourless liquid,

with an agreeable odour (Bugaje, 2008). Bioethanol can be utilized as oxygenator of gasoline,

elevating its oxygen content, allowing a best oxidation of hydrocarbons and reducing the amount

of aromatic compounds and carbon monoxide released into the atmosphere (Cardona and

Sanchez, 2007). Bioethanol is obtained from bioenergy crops and biomass which distinguishes it

from that which is produced synthetically from petroleum (Ranola et al., 2009). Different

countries use different bioenergy crops such as corn, cassava and sugarcane for bioethanol

production. Cassava and sugarcane are used mainly in Nigeria and Brazil (Naylor et al., 2007).

Experts have pointed out that cassava is the best energy crop for bioethanol production due to its

comparative advantages over all known energy crops (Wang, 2002). The ethanol yield of cassava

per unit land area is the highest among all known energy crops (Wang, 2002). High fermentable

sugar content, stable shelf live, complete and easier hydrolysis, low cost of raw materials and

simpler ethanol processing technology are the advantages of cassava flour over other flours

(Grace, 1977; Ocloo, 2002 and Vijayagopal et al., 1980). Malted cereals have been used as

sources of starch hydrolyzing enzymes, due to the fact that germination induces the synthesis of

hydrolytic enzymes (Obatolu, 2002). These malted cereals are employed in the enzymatic

saccharification of starch in most starch-based industries in Nigeria (Egwim and Oloyede, 2006).

Barley and wheat malts give high yield of amylase and fermentable sugar but are quite expensive

to import. However local sources can be used effectively for malt and enzyme production.

Sorghum and acha (Digitaria exilis) have been shown to have higher germination capacity than

other cereals such as maize and rice (Egwim and Oloyede, 2006). Sorghum alpha amylase have

been shown to be the closest alternative to imported alpha amylase for industrial purposes and

acha alpha amylase can also be as good as sorghum amylase and indeed could be a better source

(Egwim and Oloyede, 2006). Development of hydrolytic enzymes was significantly higher in

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Digitaria exilis than in sorghum (Nzelibe and Uwasike, 1995). Acha has been shown to has

higher alpha amylase yield than sorghum, acha may be a better source of alpha amylase and can

substitute for sorghum alpha amylase in industrial processing (Egwim and Oloyede, 2006). In the

present study, malted acha (Digitaria exilis) was employed as local enzyme source in the

hydrolysis of locally sourced cassava starch for bioethanol production.

1.1 Ethanol

Ethanol (CH3CH2OH) is a chemical compound which contains hydrogen, carbon and oxygen in

its chemical structure. It is also known as ethyl alcohol or grain alcohol (U.S. EPA, 2008). It is a

clear, colorless liquid with an agreeable odour (Bugaje, 2008). It is also referred to as the type of

alcohol found in alcoholic beverages. Ethanol has a somewhat sweet flavor when diluted with

water; a more pungent, burning taste when concentrated, it is more volatile than water,

flammable, burns with a light blue flame, and has excellent fuel properties for spark ignition

internal combustion engines (Wyman, 2004).

Figure 1: chemical structure of ethanol (Muhd, 2008)

Ethanol ranks second only to water as the most widely used solvent in chemical industry. It acts

as solvent for an immense range of industrial products, including paints, lacquers, dyes and oils

(Ocloo and Ayernor, 2010). It is used in medicine and motor fuels. It is also used in antifreeze

compounds and rocket fuels, pharmaceuticals, printing and cosmetics.

1.2 Historical development of bioethanol as fuel

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The use of ethanol as fuel dates back to 1826, when Samuel Morey developed an engine that ran

on ethanol and turpentine called camphene. Bioethanol was used in Germany and France by the

then incipient industry of internal combustion (IC) engines as early as 1894 (Demirbas and

Karslioglu, 2007). Bioethanol as fuel gained more prominence in 1908, when the Ford Motors in

the USA developed the Henry Ford‟s model T vehicle which was designed to use gasoline,

ethanol (from corn) or a combination of both. The use of bioethanol for fuel was widespread in

Europe and the United States during this period. Brazil has utilized bioethanol as transportation

fuel since 1925. The potential of bioethanol was ignored, especially after the World War II,

because it became more expensive than petroleum-based fuel. The energy crisis of the 1970‟s

then renewed interest in ethanol production for fuel and chemicals in both the USA and Brazil,

where mass production of bioethanol grown from corn and sugar cane started, respectively (Balat

and Balat, 2009; Balat, 2009).

The United States is the world‟s largest producer of bioethanol fuel, accounting for nearly 47%

of global bioethanol production. Brazil is the world‟s largest exporter of bioethanol and second

largest producer after the United States (Balat and Balat, 2009). Brazil produces her bioethanol

from sugarcane and cassava while the USA produces hers from corn (Naylor et al., 2007). China

is also a leading contender in bioethanol production, producing over I billion litres per year from

wheat and corn, while France which is leading other European countries, produces over 200

million gallons of ethanol from sugar beets and wheat (Sperling and Cannon, 2004).

Table 1: World bioethanol production during 2005 and 2006 (billion liters), (Balat and

Balat, 2009).

Country 2005 2006 share total in 2006 (%)

USA 15.0 18.3 46.9

Brazil 15.0 17.5 44.9

China 1.0 1.0 2.6

India 0.3 0.3 0.8

France 0.15 0.25 0.6

Others 1.55 1.65 4.2

Total 33.0 39.0

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Nigeria in 2004 joined the league of biofuel users, with a policy thrust according to the Nigeria

National Petroleum Cooperation (NNPC), to generate fuel ethanol from cassava and sugar cane.

This policy thrust was designed with the aim of generating wealth and reducing environmental

pollution (Kupolokun, 2006; Umar, 2006). At present, the Federal Government of Nigeria has

agreed to the blending of 5 percent ethanol (E5) by composition with premium motor spirit

(PMS) (Ezeobi 2008). The government adopted E5 because it believes that the level will not

damage vehicles in Nigeria, although this proportion is expected to increase to 10 per cent (E10)

in the nearest future (Ugwuanyi, 2008). The core focus of the Nigeria biofuel programme is to

ensure the production of fuel ethanol domestically.

1.3 The need for bioethanol

Bioethanol being a biofuel is produced from biological sources and has a lot of benefits which

makes it a better energy source than fossil-based fuels.

1.3.1 Environmental benefit of bioethanol

Carbon dioxide emission due to combustion of fossil fuels has become a major environmental

concern. Carbon dioxide emission contributes greatly to green house effect, climate change and

global warming. Bioethanol, is primarily seen as a good fuel alternative because the source crops

can be grown renewably and in most climates around the world. In addition, the use of

bioethanol is generally CO2 neutral. This is achieved because, in the growing phase of the source

crop, CO2 is absorbed by the plant and oxygen is released in the same volume that CO2 is

produced in the combustion of the fuel. This creates an obvious advantage over fossil fuels,

which emit CO2 as well as other poisonous emissions that have great negative impact on the

environment (Cardona and Sanchez, 2007; Hu et al., 2008). Bioethanol is bio-degradable, more

environmentally friendly and less toxic than fossil fuel.

Also, bioconversion processes in general do not produce hazardous compounds, and if toxic

solvents and chemicals are avoided in the processing stages, then fewer environmental pollutants

are produced. In addition, biomass production and microbial conversion processes can be

developed and used in a more distributed manner, avoiding the need for transport of fuels via

cargo ships or pipeline for long distances (Drapcho et al., 2008).

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Also, CO2 from ethanol fermentation can be used to extract oils and nutraceutical compounds

from biomass instead of using toxic organic solvents such as hexane (Walker et al., 1999).

Bioethanol, can be used in biodiesel production from biological oils in place of toxic petroleum-

based methanol traditionally used (Drapcho et al., 2008).

1.3.2 Respite to energy issues, depleting oil reserves and rising oil prices

Bioethanol has also become increasingly important as alternative energy source, due to depleting

oil reserves and rising oil prices (Ohgren et al., 2007). Currently, available fossil fuel sources are

estimated to become nearly depleted within the next century, with petroleum fuel reserves

depleted within 40 years (Energy Information Agency, 2007). For instance, the United States

imports 10 million barrels of oil per day of the existing world reserves (1.3 trillion barrels) as

shown in table (1). Crude oil prices have risen from less than $20/barrel in the 1990‟s to nearly

$100/barrel in 2007. The true cost of oil has been estimated as greater than $100/barrel since

2004 (Drapcho et al., 2008).

Bioethanol can create energy security and vary energy portfolio (Azmi et al., 2011). The U.S

Energy Information Administration determined that total world energy consumption in 2005 was

488EJ (exajoule, 1018

J). World consumption is expected to surpass 650EJ by 2025 (Energy

information Agency, 2007).

Bioethanol, has emerged as the most suitable renewable alternative to fossil fuel, as their quality

constituents match diesel and petrol respectively (Nugesha, 2009). Bioethanol is the most

employed liquid biofuel either as fuel or as a gasoline enhancer.

Table 2: World oil reserves and U.S import based on leading producers (Energy Information

Agency, 2007).

Country Oil reserves (billion barrels) US oil imports (million

barrels)

Saudi Arabia 267 1.50

Canada 179 1.62

Iran 132 -

Iraq 115 0.66

Kuwait 104 0.24

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United Arab Emirate 98 -

Venezuela 80 1.30

Russia 60 -

Libya 39 -

Nigeria 36 1.08

United States 21 -

China 18 -

Qatar 15 -

Mexico 13 1.60

Algeria 11 0.22

Brazil 11 -

Others 91 1.84

Total 1290 10.06 (60%)

1.3.3 Socio-economic benefit of bioethanol

The utilization of biofuels has important economic and social effects. Sheeman and Himmel

(1999) pointed out that the diversification of fuel portfolio would bring money and job back to

the USA economy. Moreover, the development of energy crops dedicated to the biofuel

production would imply a boost to agricultural sector. This analysis is also valid for developing

countries (Cardona and Sanchez, 2007), of which Nigeria is one. Fuel ethanol production has

increased remarkably, because many countries look for means of reducing oil import, boosting

rural economies and improving air quality. Global biofuel demand is projected to grow 133% by

2020 (Kosmala, 2010) and this will be of great benefit to those economies involved in bioethanol

production for export.

1.3.4 Advantages of bioethanol as engine fuel

Adding bioethanol to gasoline increases the oxygen content of the fuel, which implies a less

amount of required additive. The increased percentage of oxygen allows a better oxidation of the

gasoline hydrocarbons with consequent reduction in the emission of CO and aromatic

compounds (Malca and Freire, 2006). Wang et al (1999) corroborated this by writing that “using

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bioethanol blended fuel for automobiles can significantly reduce petroleum use and exhaust

greenhouse gas emission”.

Bioethanol has a higher octane number, broader flammability limits, higher flame speed and

higher heat of vaporization (Yoosin and Sorapipatana, 2007). These properties allow for higher

compression ratio and shorter burn time, which lead to theoretical efficiency advantages over

gasoline in an internal combustion engine (Balat, 2009). Octane number is a measure of the

gasoline quality for prevention of early ignition, which leads to cylinder knocking. An

oxygenated fuel such as bioethanol, with high octane number, provides a reasonable antiknock

value (Balat and Balat, 2009).

It is believed that a given volume of ethanol could provide energy enough to drive about 75-80%

of the distance as the same amount of gasoline, although it has only about two-third of the energy

content (Galbe and Zachi, 2002).

Bioethanol when related to MTBE (methyl tert butyl ether), which is also an oxygenator of

gasoline, is not toxic and does not pollute ground water. Bioethanol is most commonly blended

with gasoline in concentrations of 10% bioethanol to 90% gasoline, known as E10 and

nicknamed “gasohol” (Oliveria et al., 2005). Bioethanol can be used as a 5% blend with petrol

under the EU quality standard EN228. Bioethanol can be used at higher levels, for example, E85

(85% bioethanol) (Demirbas and Karslioglu, 2007)

1.3.5 Disadvantages of bioethanol as engine fuel

Bioethanol has lower energy density than gasoline (bioethanol has 66% of the energy that

gasoline has). Also, the high oxygen content of ethanol and its ability to oxidize into acetic acid

induce compatibility issues with some materials used in the engine, such as metals or polymers.

In addition, ethanol leads to azeotropes with light hydrocarbon fractions and can lead to volatility

issues, low flame luminosity, lower vapour pressure and high latent heat of vapourization

(making cold starts difficult), miscibility with water (which can cause demixing issues when

blended with hydrocarbons), toxicity to ecosystem (since its combustion in engines induces

aldehyde emissions, which has negative impact on health) (Jeuland et al., 2004). Though the use

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of bioethanol as engine fuel has some disadvantages, its advantages as engine fuel far outweigh

its disadvantages.

1.4 Feed stocks for bioethanol production

Bioethanol feedstock can be divided into three major groups: (1) starchy materials (2) sugar or

sucrose-containing feed stocks and (3) lignocellulosic biomass

1.4.1 Starch feed stocks

The starch feed stocks are mainly; cereals, tubers and roots. Cereal grains are used mostly as

food and feed. However because of their high starch content, they are also good feed stocks for

conversion to biofuels and other biobased products. Ethanol is the only biofuel that has been

produced commercially from these feed stocks in large quantities (Drapcho et al., 2008). Cereals

used as bioethanol feed stocks include: wheat, sorghum, rice and oats. The United States and

Canada are predominantly producers of bioethanol derived from corn. Tubers and roots are

potential feed stocks for ethanol production because of their high starch content. The two crops

that have been given much attention are cassava and potato. Cassava is mainly used as feed stock

in Brazil, Nigeria and China (Drapcho et al., 2008).

1.4.2 Sugar feed stocks

Main feedstock for ethanol production is sugar cane in form of either cane juice or molasses (by

-product of sugar mills). About 79% of ethanol in Brazil is produced from fresh sugar cane juice

and the remaining percentage from cane molasses (Wilkie et al., 2000). Sugar cane molasses is

the main feedstock for ethanol production in India, cane juice is not presently used for this

purpose (Ghosh and Ghose, 2003). Beet molasses are other source of fermentable sugars for

ethanologenic fermentation. Nigeria and other tropical countries use sugar cane as main feed

stock for ethanol production (Cardona and Sanchez, 2004). Sugar beet is the main feedstock for

the production of bioethanol in European Union member states (Cardona and Sanchez, 2007).

1.4.3 Lignocellulosic feedstocks

The three main components of lignocellulosic biomass are cellulose, hemicelluloses and lignin.

Cellulose, which is an abundant component in plants and wood, come in various forms and a

large fraction, comes from domestic and industrial wastes (Abella et al., 2007). Cellulose and

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hemicelluloses can be hydrolyzed with chemicals and or enzymes to monomeric sugars, which

can subsequently become converted biologically to biofuels (Drapcho et al., 2008).

Table 3: Comparison of the production and bioethanol yield from different energy crops (Wang, 2002)

Type Annual yield (ton/ha/yr) Conversion rate to sugar (%) Conversion rate to ethanol (L/ton) Annual ethanol

yield (kg/ha/yr)

Sugar cane 70 12.5 70 4900

Cassava 40 25 150 6000

Sweet sorghum 35 14 80 2800

Corn 5 69 410 2050

Wheat 4 66 390 1560

1.4.4 Cassava as a preferable feedstock for bioethanol production in Nigeria

Cassava (Manihot esculenta), also known as manioc, sagu, yucca and tapioca, is one of the most

important tropical root crops (Drapcho et al., 2008). It is a perennial woody shrub, with up to 32

per cent (fresh) starch content (Dai et al., 2006), and has 70 percent moisture. Thus, on a dry

basis it contains about 73 percent starch, which gives it a theoretical ethanol yield of 0.45 L/kg

(Drapcho et al., 2008). Cassava is the highest producer of carbohydrate with perhaps the

exception of sugar cane. The yield in major producing regions varies between 6.4 and 17 tons/ha

of fresh root, although under good conditions, yields could reach 90 tons/ha of fresh root

(Bokanga, 1996).

Cassava has a comparative advantage over other feed stocks in ethanol production in that, it is a

cheap substrate that is easily available in tropical countries (Amutha and Gunasekaran, 2001). It

can be planted on marginal lands where other agricultural crops such as sugarcane, rice, wheat

and corn do not grow well. It has a high tolerance to drought because it can survive even during

the dry season when soil moisture is low, but humidity is high (Nadir et al., 2009). Also it

requires lower soil quality compared to sugar cane as it thrives better in poor soils than any other

major plant (Nadir et al., 2009). Also, large area of little used land can be utilized for cultivation

and fertilization is rarely necessary. Moreover, a cassava ethanol plant requires less complex

processing equipment resulting in lower investments. This is due to the unique characteristics of

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cassava starch (Wang, 2002) and the low amounts of impurities which make the extraction of

starch from the root relatively easy.

Cassava crop is highly efficient in producing starch, and its starch is one of the best fermentable

substances for the production of ethanol. Cassava starch has several characteristics, which favour

its industrial use in general and in particular as a raw material in ethanol production. Some

characteristics of cassava starch are: its high purity, neutral flavour, solubility, development of

high viscosity and low tendency to retrograde compared with other starches such as those from

potato, rice and corn. According to Grace (1977) and Ocloo (2002), cassava flour promises to be

a good substrate for alcohol production due to its high content of fermentable sugars and stable

shelf life. Besides these, cassava flour has advantages such as complete and easier hydrolysis

compared to other flours (Vijayagopal et al., 1980). The ethanol yield of cassava per unit land

area is the highest among all known energy crops (Wang, 2002).

Nigeria is the world‟s largest cassava producer, producing about 36 million tons in 2004, 41.6

million metric tons in 2005(Drapcho et al., 2008), while in 2007, it was estimated at about 46

million metric tons (FAO, 2008), these figures represent about 18% of the world production.

With the current emphasis by the federal government on cassava production, future projections,

under a middle road scenario, targets 150 million tons by 2020 (Phillips et al., 2004). Adeoti

(2008) in his research found that fuel ethanol production from cassava root poses no threat to the

cassava food resource of Nigeria under various growth assumptions made.

1.5 Starch

Starch is a major energy storage component in cereal grains such as corn, wheat, sorghum, rice,

tubers (potato, yam, coco yam etc), and roots (cassava and sweet potato), (Van Beynum and

Roels, 1985).

1.5.1 Chemical composition of starch

Starch, chemical formula (C6H10O5)n, is a polymer of glucose and consist of two types of

polysaccharides; amylose and amylopectin, which represent approximately 98-99 % of the dry

weight. The ratio of the two polysaccharides varies according to the botanical origin of the

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starch; normal starches contain 70-80 % amylopectin and 20-30 % amylose (Jane, 2009). In

mutant lines of diploid species originating from crops such as maize, starches can be obtained

with amylose contents in the range of 0% (waxy maize) to 84% (amylomaize) (Matveev et al.,

2001).

Amylose and amylopectin have different structures and properties; however, both molecules are

composed of a number of monosaccharides (glucose) linked together with alpha-1-4 and/or

alpha-1-6 linkages.

Figure 2: Structure of the starch granule (Jane, 2009).

1.5.2 Amylose

Amylose is a mainly linear polymer consisting of long chains of alpha-1,4-linked glucose units.

Its molecular weight is approximately 1×105

- 1×106, it has a degree of polymerisation (DP) by

number (DPn) of 324-4920 with around 9-20 branch points equivalent to 3-11 chains per

molecule (Smith, 2001). The chains can easily form single or double helices. On the basis of X-

ray diffraction studies, the presence of A-type and B-type amylose is indicated. The structural

elements of B-type are double helices, which are packed in an anti parallel, hexagonal mode. The

central channel surrounded by 6 double helices is filled with water (36 H2O/unit cell). A-Type is

very similar to B-type, except that the central channel is occupied by another double helix,

making the packing closer. In this type, only 8 molecules of water per unit cell are inserted

between the double helices. (Sajilata et al., 2006).

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Figure 3: Chemical structure of amylose (Reis et al., 2002).

1.5.3 Amylopectin

Amylopectin is a much larger molecule than amylose with a molecular weight of 1×107

– 1× 109

and a heavily branched structure built from about 95 % alpha-1,4 and 5 % alpha-1,6 linkages.

The DPn is typically within the range 9600-15900. Chain lengths of 20 to 25 glucose units

between branch points are typical.

Figure 4: chemical structure of amylopectin (Reis et al., 2002).

Amylopectin structure is often described by a cluster model (Figure 2). Short chains (A) of DP

12-16 that can form double helices are arranged in clusters. The clusters comprise 80 % to 90 %

of the chains and are linked by longer chains (B) that form the other 10 % to 20 % of the chains.

Most B-chains extend into 2 (DP about 40) or 3 clusters (DP about 70), but some extend into

more clusters (DP about 110). B chains bind to other B chains or to a C chain which has a

reducing end and of which there is one per molecule (Sajilata et al., 2006).

On the basis of X-ray diffraction experiments, about 70 % of the mass of starch granule is

regarded as amorphous and about 30 % as crystalline (semicrystalline character). The amorphous

regions contain the main amount of amylose but also a considerable part of the amylopectin. The

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crystalline region consists primarily of the amylopectin (Sajilata et al., 2006). The schematic

organisation of amylose and amylopectin molecules in starch granule can be seen in Figure 3.

Classification based on diffractometric spectra does not follow the morphological classification,

but is also able to group most starches conveniently according to their physical properties.

Generally, most cereal starches give the so-called A-type pattern; some tuber starches (e.g.

potato) and cereal starches rich in amylose yield the B-type pattern, while legume starches

generally give a C-type pattern while V-type occurs in swollen granules (Pérez et al., 2009).

These types depend partly on the chain length making up the amylopectin lattice, the density of

packing within the granules, and the presence of water (Sajilata et al., 2006).

1.5.4 Sources and industrial application of starch

The most important sources for starch are potatoes, maize, wheat and cassava (Ellis et al., 1998).

The properties of the starches from these species differ considerably, not only in the relative

proportions of amylose and amylopectin and the characteristics of these molecules, but also in

the amount of non-starch components of starch granules, such as lipids, proteins and phosphate

groups.

Numerous industrial applications of starch exist, either in its native form or after modification.

Starch is, for example, applied in adhesive, agrochemical, cosmetic, detergent, food, medical, oil,

drilling, paper and board, pharmaceutical, plastic, purification, and textile industries (Ellis et al.,

1998).

Starch can be modified by thermal treatment, by chemical treatment, or by enzymatic treatment.

The most important enzymatic treatment method used to modify starch consists of treatment with

hydrolytic enzymes during which glycosidic bonds are broken (Guzman-Maldonado and

Paredes-Lopez, 1995).

1.5.5 Starch swelling and gelatinization

Starch granules are partially insoluble in water. They must typically undergo a multi-stage

process called gelatinization in order to solubilize the particles. At low temperatures, starch is

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insoluble because of hydrogen bonding forces between starch molecules. Though the hydrogen

bonding forces are weak, the large number of bonds involved keeps the granules from dissolving

in cold water. When starch granules in water are heated from 20oC - 60

oC, they absorb water that

disrupts the weak hydrogen bonds, and allows them to swell (Xie et al., 2006). At this phase, the

process is reversible. With further heating, the granules begin to rupture and collapse, releasing

starch chains into solution, leaving a viscous dispersion of swollen granule fragments and

dissolved starch molecules. The point at which the process becomes irreversible is called the

initial or onset gelatinization temperature. For cornstarch granules, this generally occurs between

65-80oC (Van Beynum and Roels, 1985). Further heating above the gelatinization temperature

causes further hydrogen bonding disruption, swelling and crystalline structure melting (Parker

and Ring, 2001) In addition, differential scanning calorimetry (DSC) measurements indicate that

heat is taken up during this temperature increase (Donovan, 1979). Together with all these

phenomena occurring during gelatinization, an irreversible order-disorder transition takes place

(Jenkins and Donald, 1998).

Gelatinization of starch is affected by the starch-water ratio (Parker and Ring, 2001). When the

moisture content of the starch-water mixture is low, complete swelling and disruption of the

starch granules is not possible and only part of the crystallinity of the starch granules is lost. The

remaining crystallinity only disappears after heating to higher temperatures. This could be

quantitatively described by a model based on the Flory equation that was developed for the

description of the degree of gelatinization as a function of temperature and starch concentration

(Baks et al., 2007). The article of Jenkins and Donald (1998) provides an overview of the various

theories that provide explanations for the phenomena that take place during thermal

gelatinization of starch.

Starch can also be gelatinized by application of high pressures. In certain aspects, high pressure

gelatinization differs from thermal gelatinization. During high pressure gelatinization,

disintegration of the granules is less (or the granules even remain intact), amylose solubilization

is less, and swelling of the granules is limited (Knorr et al., 2006). On the other hand, the effect

of the starch-water ratio on high pressure gelatinisation is similar. In case of thermal

gelatinisation, a higher temperature is required to achieve complete gelatinisation and in case of

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high pressure gelatinisation a higher pressure is required to achieve complete gelatinisation

(Kawai et al., 2007).

Figure 5: Gelatinization of starch granule (Kendra, 2012)

1.6 Enzymatic hydrolysis of starch

Starch is a high yield feedstock for ethanol production, but its hydrolysis is required to produce

ethanol by fermentation. Starch degradation process catalysed by enzyme is known as enzymatic

hydrolysis. One of the most important enzymatic reactions that are carried out at an industrial

scale is the enzymatic hydrolysis of starch. Starch can be hydrolysed enzymatically to yield

several commercially relevant hydrolysates (Marchal, 1999). Starch can be converted to simple

sugars by acid or enzymatic hydrolysis, but commercially, most hydrolysis today has been

replaced by enzymatic hydrolysis, as enzymatic hydrolysis has many advantages over acid

hydrolysis. Enzymes have higher specificity, so products can be developed with better control.

Secondly, enzymatic hydrolysis is performed under milder conditions, which results in fewer

side reactions and lower operating costs (Nigam and Singh, 2011). Starch must be hydrolyzed to

single glucose units before being fermented into ethanol. The enzymatic hydrolysis of starch

consists of three steps: gelatinization, liquefaction and saccharification.

Gelatinization of starch is required to increase the accessibility of the substrate and to

enhance the hydrolysis rate (Van der Maarel et al., 2002). After gelatinization, starch can be

hydrolyzed. The degree of gelatinisation is an important parameter during the enzymatic

hydrolysis of starch. The degree of gelatinisation is affected by temperature, pressure, starch

concentration, and treatment time (Baks et al., 2007). Besides the degree of gelatinisation, the

activity and stability of the enzyme are also very important.

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During liquefaction which is the first stage of hydrolysis, the viscosity of the gelatinized

starch mixture is reduced due to partial hydrolysis of the carbohydrate polymers. During the

second hydrolysis stage called saccharification, these partially hydrolyzed starch chains are

broken down into glucose, maltose, maltotriose, and some higher oligomers. The dextrose

equivalent varies between 40 and 98 depending on the enzyme that was used.

The properties of the product formed during enzymatic hydrolysis of starch can be varied by

using different enzymes (Guzman-Maldonado and Paredes-Lopez, 1995; Van der Maarel et al.,

2002). Hydrolysate products are usually characterized by their dextrose equivalent (D.E.), which

is a measure of the amount of glycosidic linkages that have been hydrolysed (Chaplin and Bucke,

1990). Three major product groups can be distinguished (Kennedy et al., 1988). Firstly, high

maltose syrups (dextrose equivalent between 40 and 55) are used in the confectionery industry

for the manufacture of hard sweets, in industries producing frozen desserts to control the crystal

formation, and in the brewing industry due to their high non-glucose fermentability (Fullbrook,

1984). Secondly, high dextrose equivalent syrups (dextrose equivalent between 56 and 68) are

used in the confectionery industry; in brewing and fermentation industries, and in general food

production (for soft drinks, jams, conserves, and sauces). Thirdly, high glucose syrups (dextrose

equivalent between 96 and 98) are used in brewing and fermentation industries, in beverage

industry, for the manufacture of crystalline D-glucose, and as starting material for the production

of fructose syrup (Fullbrook, 1984).

Alpha amylase (1,4-α-D-glucanohydrolase, E.C. 3.2.1.1) is often used for the enzymatic

hydrolysis of starch at an industrial scale, where it plays an essential role in the liquefaction and

subsequent saccharification of starch to give smaller carbohydrates (Chaplin and Bucke, 1990;

Van der Maarel et al., 2002).

1.7 Alpha amylase

The starch-degrading enzyme, alpha amylase (1,4-α-D-glucanohydrolase, E.C. 3.2.1.1) is widely

distributed in nature. This extracellular enzyme hydrolyses α-1,4 glycosidic linkages randomly

throughout the starch molecule in an endo-fashion, producing oligosaccharides and

monosaccharides including maltose, glucose and alpha limit dextrin (Nigan and Singh, 2011).

19

1.7.1 Structural and functional characteristics of alpha amylase

Alpha amylase is a monomeric, calcium binding glycoprotein. Its single polypeptide chain has

496 amino acid residues with four disulfide bridges. The alpha amylases belong to glycosyl

hydrolase family 13, which also include pollulanase, iso-amylase, and cyclodextrin glycosyl

transferase. These enzymes are highly homologous in structure consisting of two distinct

domains. Domain A contains the catalytic site surrounded by a (β/α)8-barrel that was first

discovered in triosephosphate isomerase (Janecek et al., 1 997). Domain B is composed of a

complex loop of varying length inserted between β strand 3 and α helix 3 of the (β/α)8-barrel.

The functional diversity and stability of different enzymes may be attributed to domain B

(Janecek et al., 1997).

Al1 alpha amylases share eight conserved residues and seven of these residues are located at the

active site of the catalytic (β/α)8-barrel.

Figure 6: Schematic representation of the(β/α)8- barrel (A) and 3D structure of the α-amylase of

Aspergillus oryzae or Taka amylase(B), obtained from the Protein Database (Janecek et al.,

1997).

There are always two invariant residues with carboxylic acid side chains (Glu, Asp) located at

about 4.5 to 5.5 A from each other, a characteristic of retaining enzymes which conserve the

anomeric configuration in the product.

The enzyme-catalyzed hydrolysis of the glycosidic bond takes place via general acid catalysis,

and requires a proton donor and a nucleophile/base. The hydroxyl group at the anomeric carbon

of the newly formed product can be the same (retaining) or inverted (inverting) compared to the

configuration in the glycosidic linkage. The proton donor is located at the same position in both

retaining and inverting enzyme. The nucleophilic base in the retaining enzyme is in close

20

proximity to the sugar anomeric carbon. However, this base is more distant in inverting enzymes,

in order to accommodate a water molecule between the two catalytic residues. Hence, the

commonly observed distance between the two carboxylic acid residues in the retaining enzymes

is about 4.5 to 5.5 A as compared to 10 A in the inverting enzymes (McCaxter and Withers,

1994). Two X-ray structures have been determined for cyclodextrin glycosyltransferase in

complex with an intact substrate, and the other with a covalently bound intermediate. The

study provides more definite proof for the α-retaining mechanism used by all the enzymes in

the α-amylase family.

1.7.2 Factors affecting the activity and stability of alpha amylase

The activity and stability of α-amylase are affected by temperature, pressure, pH, substrate

concentration, and additives (Fitter et al., 2001). According to Wiseman 1987, alpha amylase has

an isoelectric point of 5.4, excellent pH and temperature of enzymatic activity at 4.7 and 55oC

respectively. Optimal temperature for alpha amylase has been reported from 40oC – 60

oC; while

in the presence of calcium ion, it was increased to 75oC. The optimal requirements for the

activity and stability of alpha amylase vary with the enzyme source.

1.7.3 Sources of alpha amylase

Alpha amylases are found in most organisms that require the conversion of stored or ingested

carbohydrate. They are widespread among higher plants, animals and microorganisms (Kumar et

al., 2009). Alpha amylase has been derived from several fungi, yeasts and bacteria. However,

enzymes from fungal and bacterial sources have dominated applications in industrial sectors

(Gupta et al., 2003). In animals, the main sources of the enzyme are: the salivary gland and the

pancreas, whereas in plants, alpha amylase is most often associated with seed germination.

Table 4: Optimal pH and temperature requirements for various alpha amylases (Maureen, 1999)

Enzyme Source pH range Optimal pH Optimal

temperature(oC)

Inactivation

temperature(oC)

Pancreatic Porcine

pancrease

5.5-8.5 6.0-7.0 40-45 75

Bacterial Bacillus 4.5-9.0 6.5-7.5 70-85 95

21

subtilis

Thermo

stable

Bacterial

Bacillus

licheniforms

5.8-8.0 7.0 90-105 120

Fungal Aspergillus

oryzae

4.0-7.0 5.0-6.0 55-85 80

Malt Barley 3.5-7.5 4.5-7.0 60-70 85

1.7.4 Industrial application of alpha amylase enzyme

Alpha amylases are one of the most important and widely used enzymes whose spectrum of

application has widened in many sectors such as clinical, medical and analytical chemistry. The

most widespread applications of α-amylases are in the starch industry, where they are used in the

hydrolysis of starch into fructose and glucose syrups (Nielsen and Borchert, 2000). Beside their

use in starch saccharification, they also find application in food, baking, detergent, textile, paper,

brewing and distilling industries (Aktinson and Morituna, 1991). For instance, alpha amylase is

employed in the milling and baking industry (Anderson and Watson, 1982) to hydrolyse starch to

smaller carbohydrate, so as to reduce the dough viscosity and increase sugar levels, prolong

freshness, improve softness and crust quality. Similarly, in the brewing and beverage industries

(Shallenberger,1990; Yeshajaha, 1991), alpha amylase is employed in mash thinning, improving

runoff of wort and the general quality of the end product. The sweetener and confectionery

industries (Johnson, 1976; Wardrip, 1971; Alvin et al., 2002) have used alpha amylase to control

the ratios of different saccharrides to achieve specific product qualities. Also it has been reported

that, the use of enzymes in detergent formulations enhances the detergent ability to remove tough

stains and make the detergent environmentally safe. Amylases are the second type of enzymes

used in the formulation of enzymatic detergents, and 90% of all liquid detergents contain these

enzymes (Gupta et al., 2003). These enzymes are used in detergents for laundry and automatic

dishwashing to degrade the residues of starchy foods such as potatoes, gravies, custard,

chocolate, etc. to dextrin and other smaller oligosaccharides (Mukherjee et al., 2009). Moraes et

al (1999), Oner (2006) and Ocloo and Ayernor (2010), separately reported the use of amylase in

hydrolysis of starch for bioethanol production. Increasing utility and consumption of alpha

22

amylase in different industries has placed a greater stress on increasing indigenous production

and search of more rapid process (Gupta et al., 2008).

1.7.5 Malted cereals as sources of alpha amylase

Malted cereals have been used as sources of starch-hydrolyzing enzymes, due to the fact that

germination induces the synthesis of hydrolytic enzymes (Obatolu, 2002). Malting forms a

critical stage in the production of cereal-based beverages in which amylase and proteases

inherently embedded in the cereal grain are activated for the purpose of hydrolysis of starch and

protein into sugars and amino acids respectively (Okafor, 1987). Evans et al (2003), reported that

alpha amylase is synthesized during cereal development and stored in matured endosperms.

Alpha-amylase, as other amylases, increase markedly during germination. It has been shown that

alpha amylase yield will peak within 3 - 4 days of cereal germination (Egwim and Oloyede,

2006). George-Kraemer et al (2001), have shown that amylase activity is a good predictor of

Diastatic Power (DP) which is required in brewing processes and an important characteristic for

estimating the quality of malt for beer production (Evans, et al., 1995).

Sprouting cereals appear to be one of the popular sources of industrial amylase for some

developing economies. Amylase obtained from cereals during malting is the main enzyme

employed in enzymatic saccharification of starch in most starch-based industries in Nigeria.

Such industries include breweries, pharmaceuticals, distilleries etc. The major cereals employed

in Nigerian industries are maize, sorghum and millet (Egwim and Oloyede, 2006). Solomon et al

(1987) developed standard malt and amylase for the brewing industry from sorghum. Okafor et

al (1998) have shown however that maize, sorghum and millet have poor malting quality. This

results in low yield of amylase and consequently a low yield of fermentable sugars. Barley and

wheat are most popular sources of cereal amylase employed in the food industries. It is estimated

that about 10.4% of barley produced worldwide is used in malt and enzyme production for the

brewery and confectionery industry (Palmer, 1989). The cultivation of barley in Nigeria has

however met with little success. As such, huge amount of foreign exchange was spent annually

in importing barley malt before the ban in 1988. Industrialists have therefore resorted to

importation of microbial amylases for starch hydrolysis. However, present realities show that

local resources can be used effectively for malt and enzyme production. The common cereals

include maize, rice, sorghum, acha and millet. Earlier studies have shown sorghum to be the

23

most suitable (Shambe et al., 1989). Other local and unpopular cereals may also be a close

alternative (Egwim and Oloyede, 2006). Sorghum and acha (Digitaria exilis) were shown to

have higher germination capacity than other cereals such as maize and rice (Egwim and Oloyede,

2006). Before now, sorghum alpha amylase was shown to be the closest alternative to imported

alpha amylase for industrial purposes and acha alpha amylase can also be as good as sorghum

amylase and indeed could be a better source (Egwim and Oloyede, 2006). Nzelibe and Uwasike

(1995), reported that development of hydrolytic enzymes was significantly higher in Digitaria

exilis ("acha") than in sorghum. Though recently, Egwim and Oloyede (2006), estabilished that

alpha amylase from acha (Digitaria exilis) is a superior alternative to sorghum alpha amylase for

industrial processes. Acha has been shown to have higher alpha amylase yield than sorghum.

Acha may be a better source of amylase production and can substitute for sorghum alpha

amylase in industrial processing (Egwim and Oloyede, 2006).

1.8 Acha (Digitaria exilis)

Acha (Digitaria exilis) is probably the oldest African cereal which has survived for several

centuries mostly as weed (Jideani, 1999; Jideani, 1990). Acha, also known as finidi, fonio,

hungry rice, fonio blanc and petit mil, is the dry seed of Digitaria exilis (Jideani, 1999). For

thousands of years, West Africans have cultivated acha across the dry savannas. Indeed, it was

once their major crop. Even though few other people once heard of it, this crop still remains

important in areas scattered from Cape Verde to Lake Chad. In certain parts of Nigeria, for

instance, it is either the staple or major part of their diet (TNAP, 1996). It is one of the crops in

the West African sub region neglected by researchers and thus described as one of the orphan

crops or lost crops of Africa (National Research Council, 1996). Acha grows to a height of 30-80

cm (Gyang and Wuyep, 2005).

There are over 300 Digitaria species, which are sometimes grown as fodder; only three or four

are sometimes grown as cereals (CIRAD, 2004). The crop is perhaps one of the world‟s fastest

maturing cereals, producing grains just 6 to 8 weeks (42 to 56 days) after they are planted

(Ibrahim, 2001). The late varieties take up to 150 days to grow (CIRAD, 2004). In West Africa,

the two common species cultivated are Digitaria exilis or fonio, white fonio, fundi or findi, acha,

24

hungry rice and Digitaria iburua or black fonio, iburu etc. Of the two species, white fonio

(Digitaria exilis) is the more widely grown and used.

1.8.1 Acha cultivation, production and processing

Fonio thrives in the sandy, rocky soils of the Sahel both in drought and flood, and grows so fast

that two or three crops can be harvested each year; it is capable of growing where other crops

fail. It grows mostly in the Northern part of Nigeria and has a common Hausa name as Acha.

Philip and Itodo (2006) reported that Acha is not a demanding crop and will tolerate a wide

range of soil fertility. The potential for the production of acha grain in Nigeria is high, as it is

superior to other cereals in performance under moisture stress and low soil fertility. It also has

good capacity to respond to improved inputs.

In Nigeria, it is widely grown in commercial quantities in the cool region of Plateau State (which

is the highest producing state with an estimated production of 20,000 tons per annum), part of

Bauchi, Kebbi, Taraba, Kaduna, Gombe and Niger States (Gyang and Wuyep, 2005). Each year,

West African farmers devote approximately 300,000 hectare to acha cultivation, and yields of

600-700 kg per ha are recorded which translate to 180,000- 210,000 tons of grain annually

(National Research Council, 1996). Acha production and processing have been at a zero

mechanization level and no proper research effort has been carried out towards its mechanization

in Nigeria (Philip, 2011). There are no machines for planting, harvesting, threshing and dehulling

of acha crop on Nigerian farms. In order to bring acha to prominence, its competitiveness has to

be addressed by providing appropriate information and technologies that can support its mass

production (Philip and Issac, 2012).

Figure 7: Paddy Acha (Philip and Itodo, 2006)

25

1.8.2 Nutritional Importance of acha

Acha is one of the most nutritious of the cereals known to man (NAS, 1996). Ruskin et al (1996)

reported that acha is rich in methionine and cysteine, the limiting amino acids of most cereals.

Acha contains about 7% crude protein that is high in leucine (9.8%), methionine (5.6%) and

valine (5.8%). It is believed that its methionine content is twice as high as those of egg proteins

(Temple and Bassa, 1991).The low carbohydrate content of acha has made it to be a complement

in diabetes‟ diets. It gives attractive flavour to porridges, is a good substitute for semolina for

Europeans and has been used for brewing beer. According to Philip and Itodo (2006), acha has

the potential of providing enough food for the increasing population of poor people in West

Africa and in the Continent. Its increased utilization has prompted present study on the

possibility of modifying the grain into „acha malt‟ for making quality malt drinks and for

industrial application in hydrolyzing starch for the production of sugar syrup and bioethanol.

1.9 The Malting process

Malting is the germination of cereal grain in moist air under controlled conditions, the primary

objective being to promote the development of hydrolytic enzymes which are not present in the

ungerminated or raw grain. The malting process can be divided into three physically distinct

operations i.e. steeping, germination and drying (kilning) (Dewar et al., 1997).

1.9.1 Steeping

Steeping, which is the soaking of the cereal grain in water, is widely acknowledged as the most

critical stage of the malting process. It is performed to increase the moisture content of the grain,

to ensure rapid and uniform germination while minimizing embryo growth and respiration

(French and McRuer, 1990; Bamforth and Barclay, 1993). During imbibitions, the grain swells,

increasing its volume by approximately one quarter, the width and dept of the grain increase but

the length does not (Briggs, 1998). The acrospires, or sprout, and rootlet begin to grow, and these

are traditionally used as a guide to the malting process. The growing acrospires presses a visible

groove into the outer tissues and its length is noted as a fraction of the grain‟s (Briggs, 1998).

When imbibitions begin, the embryo and husk absorb water more rapidly than the endosperm,

and it has been thought that the embryo regulates water uptake (Bamforth and Barclay, 1993).

26

More recently, it has been suggested that structural differences in starchy endosperm also affect

water uptake during germination (Chandra et al., 1999).

1.9.2 Germination

During the malting process, germination is targeted to maximize the production of extractable

material for malt, through the process of endosperm modification. Modification describes the

physical and chemical changes that occur during malting, including the degradation of the

starchy endosperm and the associated biochemical changes (Briggs, 1998). The rate of

modification depends on several factors; the rate of moisture distribution through the endosperm,

the rate of enzyme synthesis, the extent of enzyme release and the structure of the endosperm

which will determine its digestibility (Bamforth and Barclay, 1993). Factors that are important

for the successful initiation of germination are: adequate moisture, temperature and presence of

oxygen (French and McRuer, 1990). For barley grain, germination normally occurs between

16oC to 20

oC, higher temperatures can increase growth rate, however, this cannot be sustained

over time as the rate of enzyme formation decreases.

The hormone activity of giberellic acid (GA) and abscisic acid (ABA) plays a large part in

endogenous regulation of germination by inducing and repressing the synthesis and secretion of

hydrolytic enzymes respectively (Fincher and Stone, 1993). GA stimulates the production of

hydrolytic enzymes, including alpha – amylase and beta – glucanase (Briggs, 1998; Fincher and

Stone, 1993). Alpha-amylase acts to break down starch to generate soluble sugars and is the most

extensively studied aleurone enzyme (Gibson, 2004).

1.9.3 Kilning

Kilning gradually dries the grain, reducing the high moisture levels required for germination

down to less than 5%. This must be regulated carefully to ensure the survival of the enzymes that

are crucial to brewing and distilling processes (Bamforth and Barclay, 1993). On completion of

kilning the malt grain is milled, ready to be used.

27

1.10 Fermentation

Fermentation is the internally balanced oxidation/reduction of organic compounds that takes

place in the absence of external electron acceptors (O2, NO3−, SO4

2−, etc.). In fermentation, the

oxidation of an intermediate organic compound is coupled to the reduction of another organic

intermediate. Fermentation, typically results in the formation of a carbon compound that is more

reduced than the original organic electron donor, and a carbon compound that is more oxidized.

Therefore, only a partial amount of the potential energy in the compound is captured by the

microorganism (Drapcho et al., 2008).

1.10.1 Production of ethanol from hydrolyzed starch by Saccharomyces cerevisiae

The yeast Saccharomyces cerevisiae is the universal organism for fuel ethanol production using

starch and sugar feed stocks. The sugars that are metabolizable by this organism include glucose,

fructose, mannose, galactose, sucrose, maltose, and maltotriose. Ethanol production by S.

cerevisiae is carried out via the glycolytic pathway (also known as the Embden-Myerhof-Parnas

or EMP pathway). In the fermentation of glucose by yeast such as Saccharomyces cerevisiae,

glycolysis proceeds with the electron (e-), transferred to NAD

+. Pyruvate is split into

acetaldehyde and CO2. To regenerate NAD+, acetaldehyde is reduced to ethanol by accepting the

2 e-. Thus, the net energy yield for the yeast cell is 2 ATP/ mol glucose, formed by substrate-

level phosphorylation. The oxidation states of C in glucose, ethanol, and carbon dioxide are 0,

−2, and +4, respectively, demonstrating the internal oxidation/reduction that is characteristic of

fermentation processes.

In the simplest form, production of ethanol from glucose can be expressed by the following

equation:

C6H12O6 + 2 Pi + 2 ADP →2 C2H5OH + 2 CO2 + 2 ATP + 2 H2O ……….. (1)

Glucose → 2 ethanol + 2 carbon dioxide + energy

From the above equation, it can be calculated that the theoretical yield is 0.511 g ethanol

produced per gram glucose consumed. This yield can never be realized in practice since not all

of the glucose consumed is converted to ethanol but part of it is used for cell mass synthesis, cell

maintenance, and production of by-products such as glycerol, acetic acid, lactic acid, and

28

succinic acid. Under ideal conditions, however, 90 to 95 percent of the theoretical yield can be

achieved (Thomas et al., 1996).

29

Glucose

Glucose-6-phosphate

Figure 8: Fermentation of glucose to ethanol by yeast. (Adapted from Drapcho et al., 2008)

Acetaldehyde + CO2

ATP

ADP

Fructose -6- Phosphate

ATP

ADP

Fructose -1- Phosphate

Acetaldehyde + CO2

Glyceraldehyde -3- phosphate

NAD+

NADH

P

1,3- biphosphoglycerate

ADP

ATP

3- Phosphoglycerate

Phosphoenolpyrurate

ADP

ATP

Pyrurate

NADH

NAD+

Ethanol

Glyceraldehyde -3- phosphate

NAD+

NADH

P

1,3- biphosphoglycerate

ADP

ATP

3- Phosphoglycerate

Phosphoenolpyrurate

ADP

ATP

Pyrurate

NADH

NAD+

Ethanol

30

Sugar transport is the first and also most important step in ethanol production. It has been

suggested to be the rate-limiting step of glycolysis in yeast (Becker and Betz, 1972).

Saccharomyces cerevisiae has a complex and highly effective system for sugar transport. There

are 20 genes for hexose transporters on the genome. The rate of transport can exceed 107 glucose

molecules per cell per second in fast fermenting cells (Kruckeberg, 1996).The hexose

transporters in S. cerevisiae can be divided into three classes: low-affinity transporters (Km (glucose)

=50–100 mM), moderate-affinity transporters (Km (glucose) about 10 mM), and high-affinity

transporters (Km (glucose) =1–2 mM) (Reifenberger et al., 1997).

Despite the complexity of the system, all the hexose transporters in S. cerevisiae use facilitated

diffusion for glucose transport. This mode of transport requires only a concentration gradient

across the plasma membrane. Sucrose, the major sugar in sugarcane and sugar beet, is

hydrolyzed to glucose and fructose by the enzyme invertase, which is located between the cell

membrane and the cell wall. The two monosaccharide products then are taken up by the cell.

Fructose enters the glycolytic pathway via its conversion to fructose-6-phosphate by hexokinase.

The two sugars obtained in starch hydrolysis, maltose and maltotriose, pass intact through the

cell membrane and are hydrolyzed intracellularly to glucose by the enzyme α-glucosidase.

Maltotetraose and higher polysaccharides (dextrin) are not metabolized by S. cerevisiae (Russel,

2003).

Glucose and sucrose are the two most preferred substrates among the sugars metabolized by S.

cerevisiae. In industrial applications, where many sugars are present, glucose and sucrose are

always consumed first. In fact, the presence of these two sugars causes repression in uptake and

metabolism of other sugars (Verstrepen, 2004).This catabolite repression is the result of both

competition for sugar transporters and down-regulation of the genes involved in the uptake and

metabolism of the other sugars. For example, the uptake of fructose is slowed down by glucose

since both sugars are transported into the cell by the same carriers, which have higher affinity for

glucose. In addition, glucose also can repress the expression of specific fructose transporters

(Verstrepen, 2004). Glucose represses the expression of the maltose-utilisation genes even when

maltose is present (Federoff et al., 1983).Glucose also causes repression of the galactose

utilisation genes even in the presence of that sugar (Adams, 1972). Catabolite repression is not

31

immediately relieved after glucose and sucrose are depleted. In fact the effect may last for

several hours. The repression of utilization of other sugars by glucose and/ or sucrose may have a

negative effect on fermentation rates in industrial ethanol production (Drapcho et al., 2008).

Saccharomyces cerevisiae requires certain minerals (e.g., Ca, Mg, Mn, Co, Fe, Cu, K, Na, Zn)

for growth and ethanol fermentation. A number of organic compounds also have been observed

to improve ethanol production. Most of the required nutrients normally are already available in

industrial feed stocks for ethanol production (Jones and Greenfield, 1984).

Saccharomyces cerevisiae is inhibited by its own product, that is, ethanol. Ethanol increases

fluidity and permeability of cell membrane and causes leakages of ions and small metabolites

(Salgueiro et al., 1988). Ethanol inhibition in S. cerevisiae can be reversed by addition of

calcium. However, addition of calcium to the fermentor must be controlled tightly since high Ca:

Mg ratios may cause antagonism of essential biochemical functions of magnesium and

negatively affect growth and ethanol production (Russel, 2003).

Acetic or lactic acid can also inhibit S. cerevisiae. In ethanol fermentation these acids are minor

by-products and accumulate only to relatively low concentrations, which are well below the

inhibitory levels. However, in industrial ethanol production completely sterile conditions are

never maintained and contamination by lactic acid and acetic acid bacteria can raise the

concentrations of these acids to harmful levels. The inhibitory effects of lactic acid and acetic

acid on growth and ethanol production are synergistic (Narendranath et al., 2001) and pH

dependent . Oxygen plays an important role in S. cerevisiae metabolism (Russel, 2003). Ethanol

inhibition of growth is reduced under micro-aerobic conditions compared to anaerobic conditions

(Hoppe and Hansford, 1984). Aeration conditions also impact the synthesis of by-products. At a

respiratory quotient (RQ) controlled at 17 or below, glycerol synthesis was almost eliminated

(Franzen, 2003). However, excess oxygen will result in significantly higher cell mass production

and lower ethanol yield (Russel, 2003).Thus, in practice optimal aeration conditions must be

carefully determined and maintained to achieve optimal ethanol production (Drapcho et al.,

2008).

32

1.11 AIM AND OBJECTIVES

The aim of this study was to investigate the potential of acha (Digitaria exilis) as local source of

hydrolytic enzymes in the production of bioethanol from cassava starch. The specific objectives

included:

Production of hydrolytic enzymes by malting of acha grains.

Hydrolysis of cassava starch by the malted acha and determination of the simple/reducing

sugar yields.

Determination of the optimal conditions for the hydrolysis of cassava starch by the

malted acha.

Fermentation of the cassava starch hydrolysate by yeast and the determination of the

ethanol yield.

Distillation of the alcohol produced from the fermented cassava starch hydrolysate.

33

CHAPTER TWO

MATERIALS AND METHODS

2.1 Materials

The cassava starch and acha grains used for this study were obtained from the Department of

Biochemistry, University of Nigeria Nsukka. The dry cassava starch flour was washed with

water and sun dried to remove dirt from it. The acha grains were carefully sorted to remove

damaged kernels and debris.

2.1.1 Instruments/equipment

Equipment Manufacturer

Autoclave Arnold and Sons, Nigeria

Beakers Pyrex, England

Blender Landers Medellin, USA.

Burette Pyrex, England

Centrifuge PAC, Pacific, England

Condenser Pyrex, England

Digital Weighing Balance G and GR

, China

Filter Paper Whatman, England

Glass Rod Pyrex, England

Hot Air Oven (0-200oC) Gallenkamp, England

Hot Plate Gallenkamp, England

Measuring Cylinder Pyrex, England

pH-Meter Hanna Instruments, India

34

Reagent Bottle Pyrex, England

Refrigerator Haier Thermocool, Nigeria.

Retort Stand Pyrex, England

Round Bottom Flask Pyrex, England

Spatula Pyrex, England

Spectrophotometer Spectrumlab 22PC, England

Syringe Dana-Jet, Nigeria

Test Tube Pyrex, England

Thermometer Pyrex, England

Water Bath Waterbath, Model DK, England

Wrist Watch Corum, Switzerland

2.1.2 Chemicals

The chemicals used for this study were of analytical grade and were procured from scientific

shops in Nsukka, Enugu State of Nigeria as well as from the Department of Biochemistry,

University of Nigeria, Nsukka. The chemicals and their manufacturers include;

Chemical Manufacturer

Acetic Acid M&B, England

Ammonium Molybdate Hopkin & Williams, England

Anhydrous Sodium Carbonate M&B, England

Baker‟s Yeast Keramel, Germany

Copper Sulfate Riedel-de Haan, Germany

Crystalline Phenol M&B, England

Dinitrosalicylic Acid BDH, England

Dipotassium Hydrogen Phosphate Lab Tech, India

35

Distilled Water STC, UNN

Glucose BDH, England

Potassium Dichromate Riedel-de Haan, Germany

Potassium Dihydrogen phosphate Merck, Germany

Potassium Sodium Tartrate Sigma, USA

Sodium Acetate Sigma, USA

Sodium Hydrogen Arsenate M&B, England

Sodium Hydroxide Riedel-de Haan, Germany

Sodium Sulfate Riedel-de Haan, Germany

Sodium sulfite M&B, England

Sodium Thiosulfate Penta Hydrate M&B, England

Starch Hopkins & Williams, England

Sulfuric Acid M&B, England

2.2 Methods

2.2.1 Preparation of reagents

Reagents for reducing sugar determination and preparation of glucose standard curve

Alkaline copper tartrate reagent

A) A quantity, 2.5 g anhydrous sodium carbonate, 2 g sodium bicarbonate, 2.5 g potassium

sodium tartrate and 20 g anhydrous sodium sulfate were dissolved in 80 ml distilled water

and made up to 100 ml.

B) A quantity, 15 g copper sulfate was dissolved in a small volume of distilled water. One

drop of sulfuric acid was added and the volume made up to 100 ml with distilled water.

Then, 4 ml of solution B was mixed with 96 ml of solution A before use.

36

Arsenomolybdate reagent

A quantity, 2.5 g ammonium molybdate was dissolved in 45 ml of distilled water and 2.5 ml of

sulfuric acid was added and mixed well. Then, 0.3 g of disodium hydrogen arsenate was

dissolved in 25 ml of water. The two solutions were mixed thoroughly and incubated at 37oC for

24 h.

Stock Standard glucose solution

A quantity, 100 mg of glucose, was dissolved in 100 ml of distilled water.

Working standard glucose solution

A quantity, 10 ml of stock, was diluted to 100 ml with distilled water.

Reagents for the determination of ethanol yield during fermentation

Acid dichromate solution (0.01 molL-1

)

A quantity, 125 ml of water was measured into 500 ml conical flask. Then, 70 ml of concentrated

sulfuric acid was carefully added with constant swirling. The flask was cooled under cold water

tap and 0.75 g of potassium dichromate added. The solution was diluted to 250 ml with distilled

water.

Starch indicator solution (1.0 % solution)

A quantity, 1.0 g of soluble starch, was dissolved in 100 ml of hot distilled water. The solution

was stirred until the starch dissolved.

Sodium thiosulfate solution (0.03 molL-1

)

A quantity, 7.44 g of Na2S3O3.5H2O, was dissolved with distilled water in a one litre volumetric

flask. The solution was subsequently made up to the one litre mark.

37

Reagents for assay of α-amylase activity during malting

Dinitrosalicylic Acid reagent (DNS Reagent)

The following, 1 g dinitrosalicylic acid, 200 mg crystalline phenol and 50 mg sodium sulphite

were dissolved by stirring in 100 ml of 1 % NaOH and stored at 4oC.

2 % soluble starch

A quantity, 2 g of starch dissolved in 100 ml of hot water and stirred thoroughly.

Preparation of buffers

0.05 M Citrate buffer

A quantity, 10.51g of sodium citrate, was dissolved in 500ml of distilled water, then the pH was

adjusted to 6.0 with citric acid and the volume made up to 1000ml with distilled water.

0.2 M Acetate buffer

A quantity, 5.45 g of sodium acetate, was dissolved in 200 ml of distilled water to give 0.2 M

sodium acetate buffer.

A quantity, 3 ml of acetic acid, was dissolved in 252 ml of distilled water to give 0.2 M acetic

acid.

0.2 M Phosphate buffer

A quantity, 2.72 g of potassium dihydrogen phosphate, was dissolved in 100 ml of distilled water

to give 0.2 M potassium dihydrogen phosphate buffer.

A quantity, 3.48 g of dipotassium hydrogen phosphate, was dissolved in 100 ml of distilled water

to give 0.2 M dipotassium hydrogen phosphate buffer.

2.2.2 Glucose standard curve by the Nelson-Somogy method.

A quantity, 0.1, 0.2, 0.3, 0.4, 0.5, 0.6, 0.7, 0.8, 0.9 and 1 ml, of the working standard glucose

solutions was each pipetted into ten test tubes. The sample and standard test tubes were made up

to 2 ml with distilled water. Then, 2 ml of distilled water was added into a separate test tube to

38

set as blank. After wards, 1ml alkaline copper tartrate reagent was added to each test tube. The

test tubes were placed in a boiling water bath for ten minutes. The tubes were cooled and 1ml

arsenomolybdate reagent was added to each test tube. The volume of each test tube was made up

to 10ml with distilled water. The absorbance of the blue colour was read at 620 nm after ten

minutes. The graph of the absorbance against glucose concentration was plotted.

2.2.3 Alpha amylase yield in sprouting acha cereal (Ezeogu et al., 1997)

A quantity, 500 g of cereal, was cleaned and sprouted using a modified method described by

Ezeogu et al (1997). Samples were withdrawn from the sprouting cereal at 24 h intervals up to

144 h (six days). A quantity, 1 g of sprouting cereal, was milled and added with 5 ml of pre-

chilled 0.05 M citrate buffer, pH 6.0; the resulting homogenate was centrifuged at 10,000 xg for

10 minutes. Enzyme activity was assayed in the supernatant as yield of crude enzyme.

2.2.4 Assay of Enzyme Activity (Kang et al., 1997)

Alpha amylase activity was assayed using a modified method described by Kang et al (1997). A

quantity, 0.1 ml of supernatant (crude enzyme), was pipetted into separate test tubes and 0.9 ml

of 2 % soluble starch was added. The mixtures were incubated under constant shaking in a water

bath at 50oC for 30 minutes. The reaction was subsequently stopped by adding dinitrosalicylic

acid (DNSA) reagent. The mixture was boiled for at least 3 minutes for colour development.

Absorbance of the mixture was read at 550 nm against blank. The blank contained all the assay

reagents without the enzyme.

2.2.5 Malting of acha grains (modification of Ezeogu et al., 1997)

Acha grains (1 kg) were steeped in 3 litres of distilled water after prior sterilization with sodium

hypochlorite solution (containing 1 % available chlorine for 20 min). Steeping lasted for 96 h.

Grains were thereafter germinated for six days in germination trays in an atmosphere of near

water saturation. Grains were turned every 12 h and watered by spraying with 60 ml distilled

water At 24 h intervals, germinating grains were withdrawn and immediately kiln dried. Kilning

was for 24 h at 50°C in a forced draught oven. The malt was ground using a blender.

39

2.2.6 Hydrolysis of cassava starch using malted acha (Ocloo and Ayernor, 2010)

The method described by Ocloo and Ayernor (2010) was used. A quantity, 1 kg of cassava flour,

was mixed with 5000 ml of water to form slurry. The slurry was heated to gelatinization at 70°C

and allowed to cool at 50oC. About 250 g of malted Acha was added to the gelatinized mash at

70oC, stirred, and the mixture allowed to cool gradually to 50°C for the amylase in the malt to

convert the gelatinized starch to sugars. Thinned liquour was then heated to 70°C and the last

batch of 250 g malted Acha added to further convert the unhydrolysed starch to sugars. The

mixture was boiled briefly and immediately filtered using sieving cloth. The sweetwort (which is

the liguid extracted from the hydrolysed cassava starch which contains the sugars that will be

fermented by the yeast) produced was boiled again to arrest further enzyme action and then

cooled. Cassava flour hydrolysate having specific reducing sugar content was obtained.

2.2.7 Determination of reducing sugar content by the Nelson-Somogy method.

A quantity, 0.2 ml of the sweetwort, was made up to 2 ml with distilled water and 2 ml of

distilled water was added into a separate test tube to set as blank. After wards, 1 ml alkaline

copper tartrate reagent was added to each test tube. The test tubes were placed in a boiling water

bath for ten minutes. The tubes were cooled and 1 ml arsenomolybdate reagent was added to

each. The volume of each test tube was made up to 10 ml with distilled water. The absorbance of

the blue colour that developed was read at 620 nm after ten minutes. The reducing sugar content

was deduced from the graph of the glucose standard curve.

The reducing sugar concentration was calculated from the standard glucose curve using the

equation: y = 0.017 x + 0.126

Where y = absorbance and x = concentration of reducing sugars (µg/ml).

The total reducing sugar concentration = x (µg/ml) X dilution factor (200)

The dextrose equivalent (DE) or percentage reducing sugar is calculated as:

g reducing sugar expressed as glucose

% Reducing sugar = X 100

g dry solid weight

40

2.2.8 Determination of optimum quantity of enzyme (malted acha) to substrate (cassava

starch) during the hydrolysis of cassava starch (modification of Ocloo and Ayernor, 2010)

A) A quantity, 1 g of cassava starch, was weighed into five test tubes and 10 ml of water was

added to give 10 % w/v slurry. The slurry mixtures in the test tubes were shaken to mix

thoroughly. Then, 0.1 g, 0.2 g, 0.3 g, 0.4 g, and 0.5 g of the malted acha were weighed

out respectively. The test tubes were labeled as follows: 0.1 g, 0.2 g, 0.3 g, 0.4 g and 0.5

g respectively. The test tubes containing the starch slurries were put in a water bath and

heated to 70oC for 20 min until the starch slurries gelatinized. The test tubes were

subsequently allowed to cool to 50oC. The weighed out malted acha were added to the

labeled test tubes accordingly. The mixtures were stirred thoroughly using a glass rod and

the temperature raised gradually to 70oC and then incubated for 2 h in a water bath. The

mixture was sieved using a sieving cloth. The sweetwort was boiled further to arrest

further enzyme action and its reducing sugar content determined using the Nelson-

Somogy method.

The experiment above was repeated using 2 g of cassava starch.

2.2.9 Determination of optimum pH for cassava starch hydrolysis using malted acha

(modification of Ocloo and Ayernor, 2010)

A quantity, 1 g of cassava starch, was weighed into eight test tubes labeled, pH 3.0, pH 3.5, pH

4.0, pH 4.5, pH 5.0, pH 5.5, pH 6.0, pH 6.5, pH 7.0 and pH 7.5 respectively. Buffer solutions of

pH range 3.0 - 7.5 were prepared using both 0.2 M acetate (3.0 - 5.0) and 0.2 M phosphate (5.5 -

7.5) buffers. Then, 10 ml of the corresponding buffers were added to each of the labeled test

tubes accordingly to form slurry. The cassava starch slurries in the test tubes were shaken to mix

thoroughly. The test tubes containing the starch slurries were put in a water bath and heated to

70oC for 20 min until the starch slurries gelatinized. The test tubes were allowed to cool to 50

oC.

A quantity, 0.5 g of malted acha was added to each test tube. The mixture was stirred thoroughly

using a glass rod and the temperature raised gradually to 70oC and then incubated for 2 h in a

water bath. The mixture was sieved using a sieving cloth. The sweetwort was boiled further to

arrest further enzyme action and its reducing sugar content determined by the Nelson-Somogy

method.

41

2.2.10 Determination of optimum temperature for cassava starch hydrolysis using malted

acha (modification of Ocloo and Ayernor, 2010)

A quantity, 1 g of cassava starch, was weighed into four test tubes labeled, 50oC, 60

oC, 70

oC and

80oC respectively. Then, 10 ml of 0.2 M acetate buffer of pH 4.5 was added to each of the

labeled test tubes to form slurry. The slurry mixtures in the test tubes were shaken to mix

thoroughly. The test tubes containing the starch slurries were put in a water bath and heated to

70oC for 20 min until the starch slurries gelatinized. The test tubes were allowed to cool to 50

oC.

A quantity, 0.5 g of malted acha was added to each test tube. The mixtures were stirred

thoroughly using a glass rod. Each of the labeled test tube was incubated at its particular

temperature (i.e, 50oC, 60

oC, 70

oC and 80

oC) for 2 h in a water bath respectively. The mixture

was sieved using a sieving cloth. The sweetwort was boiled further to arrest further enzyme

action and its reducing sugar content determined by the Nelson-Somogy method.

2.2.11 Determination of optimum time for cassava starch hydrolysis using malted acha

(modification of Ocloo and Ayernor, 2010)

A quantity, 1 g of cassava starch, was weighed into seven test tubes labeled, 1 h, 2 h, 3 h, 4 h, 5

h, 6 h, and 7 h respectively. Then, 10ml of 0.2 M acetate buffer of pH 4.5 was added to each of

the labeled test tubes to form slurry. The slurry mixtures in the test tubes were shaken to mix

thoroughly. The test tubes containing the cassava starch slurries were put in a water bath and

heated to 70oC for 20 min until the starch slurries gelatinized. The test tubes were then allowed

to cool to 60oC. A quantity, 0.5 g of malted acha was added into each of the test tubes. The

mixture was stirred thoroughly using a glass rod, the temperature maintained at 60oC and then

incubated for 1 h, 2 h, 3 h, 4 h, 5 h, 6 h, and 7 h in a water bath accordingly. The mixture was

sieved using a sieving cloth after each time interval. The sweetwort was boiled further to arrest

further enzyme action and its reducing sugar content determined by the Nelson-Somogy method.

Note: it is important to state that the sweetworths produced after the hydrolysis of the cassava

starch by the malted acha was diluted by a factor of 1:200 dilutions (i.e 1 ml of the sweetworth in

200 ml of water) before the determination of its reducing sugar content by the Nelson-Somogy

method.

42

2.2.12 Production of ethanol from cassava starch hydrolysate (modification of Ocloo and

Ayernor, 2010)

A) A quantity, 20 g of cassava starch was weighed into 200 ml of 0.2 M acetate buffer of pH 4.5

to form slurry. The slurry mixture was stirred thoroughly and heated in a water bath to 70oC for

20 min, until the cassava starch slurry gelatinized. The slurry was allowed to cool to 50oC. A

quantity, 10 g of malted acha was added to the gelatinized cassava starch and the mixture

incubated at 60oC for 5 h. The mixture was sieved using a sieving cloth after the hydrolysis. The

sweetwort was boiled further to arrest further enzyme action and its reducing sugar content

determined by the Nelson-Somogy method. The cassava starch hydrolysate produced after the

hydrolysis of the cassava starch by the malted acha, was fermented in fermentation bottles

(previously sterilized to exclude any microorganis) containing about 30 ml cassava starch

hydrolysate and 1 ml of yeast inocula, of varying concentrations of 5 % (5 g of baker‟s yeast

dissolved in 100 ml of distilled water), 10 %, 15 %, 20 % and 25 % respectively, for each

fermentation bottle. The bottles were topped with tubes to allow carbon dioxide (CO2) to escape.

Fermentation was allowed for 9 days at 28oC - 30°C. The ethanol yield of each fermentation

medium was determined after each day.

2.2.13 Determination of ethanol yield (Acid dichromate redox titration method)

The fermented cassava starch hydrolysate sample was diluted to 1:50 (1 ml of sample made up to

50 ml) with distilled water. Then, 10 ml of acid dichromate solution was transferred to a 250 ml

conical flask with matching stoppers. Also, 1 ml of the diluted sample was added into a sample

holder (5 ml beaker). The sample holder was subsequently suspended over the dichromate

solution and held in place with a rubber stopper. The flask was stored overnight at 25 oC -30

oC,

after which the stopper was carefully removed and the sample holder discarded. The wall of the

flask was subsequently rinsed with 100 ml of distilled water and 1 ml of potassium iodide

solution was also added. The flask was swirled to mix its content. Blank was prepared by adding

10 ml of acid dichromate solution into another conical flask, after which 100 ml of water and 1

ml of potassium iodide solution were added and the flasks swirled to mix. A burette was filled

with sodium thiosulfate and both the sample and blank flask titrated with the sodium thiosulfate

solution. 1 ml of starch indicator solution was added when the brown iodine colour faded to

yellow and the titration was continued until the blue colour caused by the indicator disappeared.

43

The samples were titrated twice to get concordant results. The ethanol concentration was

calculated using the equations for the reaction shown below.

Cr2O72-

+ 14 H+ + 6I

- 2 Cr

3+ + 3I2 + 7H2O ………….. (1)

2 S2O32-

+ I2 S4O62-

+ 2I- ……………………………… (2)

The average volume of sodium thiosulfate used for both blank and sample titrations were

determined. The volume of sodium thiosulfate used for the sample was subtracted from the

volume used for the blank titration. This volume of sodium thiosulfate was now used to

determine the ethanol concentration. The number of moles of sodium thiosulfate in this volume

was calculated.

Using the equations above, the relationship between the moles of sodium thiosulfate and the

moles of ethanol was determined as follows

- 6 mol of S2O32-

is equivalent to 1 mol Cr2O72-

- And 2 mol of Cr2O72-

is equivalent to 3 mol of C2H5OH

- Then 1 mol of S2O32

is equivalent to 0.25 mol of C2H5OH

The equation for the ethanol yield in mol/dm3 is given as

Ethanol yield (mol/dm3) = 0.0075 M of ethanol × difference in volume of sodium thiosulfate

used in blank and sample × dilution factor.

Ethanol yield (g/dm3) = Molarity × Molar mass

Ethanol yield (g/100 ml) = Concentration (g/dm3)/ 10

44

2.2.14 Distillation process (Ocloo and Ayernor, 2010)

About 4000 ml of the fermented cassava starch hydrolysate was measured into a volumetric flask

at 20 °C and was washed into the distillation flask with about 20ml of distilled water. The

sample was distilled slowly into a receiving flask at temperatures between 78.3oC – 80°C, using

a simple distillation set up. About 150 ml of the distillate was collected and stored in a bottle.

2.2.15 Qualitative determination of ethanol (Chromic Acid test)

The chromic acid test may be used to distinguish an aldehyde, primary and secondary alcohols

from ketones and tertiary alcohols. A quantity, 5 drops of test sample was added to a test tube. A

quantity, 10 drops reagent grade acetone was also added to the test tube. Also, 2 drops of the

Jones reagent (chromic acid reagent) were added. The test tube was stirred to mix its contents.

The solution was allowed to stand for 1 minute. The result was recorded. The appearance of a

blue-green color or precipitate was a positive result.

2.2.16 Statistical analyses

The comparison of the significant effect of yeast concentration and time was made using one –

way Analysis of Variance (ANOVA). The variability of ethanol concentration, with differences

in means was compared using Post – Hoc multi comparison test. Their differences were

considered significant at p< 0.05 and our decisions made using Least Significant Difference

(LSD) method at p< 0.05. Also a regressional model was proposed to account for how yeast

concentration relate with time in determining ethanol yield. Statistical Package for Social

Sciences (SPSS) version 16 was used for this analysis.

45

CHAPTER THREE

RESULTS

3.1 Result of reducing sugar determination

Figure 9: Glucose standard curve

y = 0.0176x + 0.1263 R² = 0.9961

0

0.2

0.4

0.6

0.8

1

1.2

0 10 20 30 40 50 60

Ab

sorb

an

ce

Concentration (µg/ml)

46

3.2 Result of the malting process

The alpha amylase activity (mg glucose/ml/min) of the malted acha was determined for 6 days of

malting. Maximum alpha amylase activity was observed on the 5th

day of malting (Fig 10).

Figure 10: alpha amylase activity during malting of acha

0.04

0.042

0.044

0.046

0.048

0.05

0.052

0 1 2 3 4 5 6 7

am

yla

se a

ctiv

ity (

mg/m

l/m

in)

Malting time in days

47

3.3 Optimal conditions for cassava starch hydrolysis using the malted acha

Malted acha (0.1 g, 0.2 g, 0.3 g, 0.4 g and 0.5 g) was used to hydrolyse 10 % and 20 % w/v

cassava starch slurries. As shown in figure 11, 0.5 g malted acha to 10 % starch slurry gave the

maximum reducing sugar yield as was observed.

Figure 11: Optimum quantity of enzyme (malted acha) to substrate (cassava starch) during

cassava starch hydrolysis.

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0 0.1 0.2 0.3 0.4 0.5 0.6

Ab

sorb

an

ce (n

m)

Quantity of malted acha (g)

10% starch slurry

20% starch slurry

48

The optimum pH for the hydrolysis of cassava starch with malted acha was determined over a

pH range of 3.0 to 7.5. As shown in figure 12, optimum pH of 4.5 was observed.

Figure 12: Effect of pH on cassava starch hydrolysis.

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0.9

1

0 1 2 3 4 5 6 7 8

Ab

sorb

ance

(n

m)

pH

49

The optimum temperature for the hydrolysis of cassava starch with malted acha was determined

over a temperature range of 50oC to 80

oC. Optimum temperature of 60

oC was observed (Fig 13).

Figure 13: Effect of temperature on cassava starch hydrolysis.

0

0.1

0.2

0.3

0.4

0.5

0.6

0 10 20 30 40 50 60 70 80 90

Ab

sorb

an

ce (

nm

)

Temperature (oC)

50

The optimum time for the hydrolysis of cassava starch with malted acha was determined over a

time range of 1 to 7 h. Optimum time of 5 h was observed (Fig14).

Figure 14: Effect of time (in hours) on cassava starch hydrolysis.

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0 1 2 3 4 5 6 7 8

Ab

sorb

ance

(n

m)

Time (h)

51

3.3 Results of the alcohol production process/ fermentation

The effect of yeast concentration and time on ethanol yield was determined over a period of 9

days; yeast concentrations of 5 % to 25 % were used. As shown in figure 14, least ethanol yield

of 1.38 g/100ml which was produced by the 5 % fermentation medium after one day and

maximum ethanol yield of 8.11 g/100ml, which was produced by the 25 % yeast fermentation

medium after 9 days were observed.

Figure 15: Effect of yeast concentration (percent) and time (days) on ethanol yield (g/100ml)

during fermentation.

0

1

2

3

4

5

6

7

8

9

0 2 4 6 8 10

eth

an

ol

yie

ld (

g/1

00m

l)

fermentation time in days

five percent

ten percent

fifteen percent

twenty percent

twenty-five percent

52

The graph of the mean ethanol concentration (yield) against time as shown in figure 16 was a

hyperbolic curve of which the curve started on the 3rd

day of fermentation indicating that

inhibition has set in.

Figure 16: Graph of mean ethanol concentration (yield) (g/100ml) against fermentation time

(days).

53

CHAPTER FOUR

DISCUSSION AND CONCLUSION

4.1 Discussion

The goal of the study was to employ locally sourced raw materials for the production of

bioethanol. Locally sourced cereal (Digitaria exilis) was used as the source of hydrolytic

enzymes for the hydrolysis of locally sourced starch (cassava) feed stock.

Bioethanol has emerged as the most suitable renewable alternative to fossil fuel, since, it is

environmentally friendly, biodegradable, less toxic and do not contribute to green house gas

effect and global warming. In addition, large scale bioethanol production has positive economic

and social effect by providing more money and job in those countries involved in its large scale

production. Malted cereals have been used as sources of starch hydrolysing enzymes, due to the

fact that germination induces the synthesis of hydrolytic enzymes such as alpha amylase

(Obatolu, 2002). Alpha amylase plays important role in the liquefaction and subsequent

saccharification of starch to give smaller carbohydrate (Chaplin and Bucke, 1990). The major

cereals previously used in Nigeria as sources of alpha amylase are maize, sorghum, millet and

barley. Due to the ban on the importation of barley in 1988 by the Federal Government of

Nigeria, sorghum became the closest alternative to barley in terms of synthesis of hydrolytic

enzymes until Nzelibe and Uwasike (1995) reported that “development of hydrolytic enzymes

was significantly higher in Digitaria exilis (Acha) than in sorghum”. Acha is referred to as the

lost crop of Africa, since little is known about this cereal and its potentials. In the present study,

the potential of acha (Digitaria exilis) as a local source of hydrolytic enzymes for cassava starch

hydrolysis was studied and the optimum conditions for this process investigated.

The development of alpha amylase during the germination of acha grain was studied for six days,

after the grain was initially steeped for four days. The enzyme activity was given as the amount

of enzyme required to liberate a unit of glucose per minute (mg glucose/ml/min) under the

reaction conditions. The result represented by figure 9, showed a gradual increase in the alpha

amylase activity with time from day 0 to day 5 of sprouting, after which there was a sharp

decline in the amylase activity on the 6th

day. The least activity of the alpha amylase (0.0416 mg

glucose/ml/min) was observed on day 0 of the acha sprouting while the maximum activity

54

(0.0510) was observed on the 5th

day of the sprouting. The result showed that time was

significant to alpha amylase yield during germination i.e., as time increased, alpha amylase yield

also increased. This observation agrees with the finding of Egwim and Adenomom (2009) which

showed that time was significant to alpha amylase yield in the cereals they studied of which acha

grain was among them. Nzelibe and Uwasike (1995) reported that alpha amylase activity

increased with germination time. Egwim and Oloyede (2006) also observed a similar trend when

they sprouted different cereals including acha for 200 h and obtained the maximum activity of

the alpha amylase at 96 h. The present study showed that acha must be sprouted for at least five

days for maximum alpha amylase production. According to Egwim and Adenomon (2009), the

yield of alpha amylase which is a function of time (when other conditions are constant), is a good

indicator of malting quality. This fact also agrees with the findings of Jin – Xin et al (2006), who

did show a strong relationship between amylase activity and malting quality.

Optimum quantity of enzyme (malted acha) to substrate (cassava starch) during cassava starch

hydrolysis with malted acha was studied. The experiment was carried out in two batches, in the

first batch, the substrate concentration was kept constant at 10 % w/v (1 g of cassava starch in 10

ml of distilled water), while the enzyme (malted acha) quantity was varied from 0.1 g – 0.5 g.

In the second batch, the substrate concentration was kept at 20 % w/v (2 g of cassava starch in

10ml of water), while the enzyme (malted acha) quantity was varied also from 0.1 g – 0.5 g. At

the end of the hydrolysis, the result (see figure 11) showed that 0.5 g enzyme (malted acha) to 10

% w/v cassava starch slurry gave the maximum concentration of reducing sugar released after

the hydrolysis. The 0.5 g enzyme (malted acha) to 20 % w/v starch slurry gave a lower reducing

sugar concentration when compared to the 0.5 g enzyme (malted acha) to 10 % w/v starch slurry.

From the result, it can be observed that lower substrate concentrations are more suitable in the

enzymatic hydrolysis of starch. Mojovic et al (2006), made a similar observation when he

reported that a 16% corn starch suspension gave far much higher glucose yield when hydrolyzed,

than a 40% corn starch suspension that was also hydrolyzed by the same quantity of enzyme.

Monica et al (2011) reported that increased substrate concentrations produced a decreased

enzymatic activity. A similar observation has also been reported by Baks et al (2008). These

observations could be due to diffusional constraints during enzymatic hydrolysis of starch caused

by increasing viscosity by gelatinized starch and also due to substrate inhibition experienced

when higher substrate concentrations were used.

The optimum pH for the hydrolysis of cassava starch by malted acha was determined over the

pH range of 3.0 to 7.5, the result represented by figure 12, showed that the amount of reducing

sugar produced during the hydrolysis of the cassava starch using the malted acha increased

gradually as the pH increased from 3.0 to 4.5, after which there was a gradual decrease in the

55

amount of reducing sugar produced as the pH increased further from 4.5 to 7.5. Higher reducing

sugar concentrations were observed between the pH 4.5 - 6.0, with the maximum concentration

of reducing sugar obtained at the pH 4.5.This means that the optimum pH for the hydrolysis of

cassava starch using the malted acha was 4.5. Lower reducing sugar concentration was obtained

at pH values less than 4.5 and higher than 6.0, with the least reducing sugar concentration

obtained at pH 3.0 and 7.5 respectively. The lower reducing sugar concentration observed at

these pH ranges (3.0 - 4.0 and 6.5 - 7.5) may be due to denaturation effect on the hydrolytic

enzymes of the malted acha at these pH values. The effect of pH observed in the present study,

indicated that change in pH had a significant effect on the rate of hydrolysis of cassava starch by

the amylolytic enzymes present in the malted acha. It has been reported that pH affects the

stability and activity of enzymes (Chaplin and Bucke, 1990). Similarly, Betiku and Ajala (2010)

reported that pH had a significant effect on the rate of hydrolysis of bread fruit starch by alpha

amylase.

The optimum temperature for the hydrolysis of cassava starch using malted acha was determined

over a temperature range of 50oC – 80

oC. The result represented by figure 13, showed that the

reducing sugar concentration during the hydrolysis of the cassava starch using malted acha

increased between 50oC – 60

oC and then decreased as the temperature increased above 60

oC.

The optimum temperature for the hydrolysis process was 60oC, since maximum amount of

reducing sugar was released at this temperature as indicated by the peak shown in figure (13).

The least amount of reducing sugar was released at higher temperature of 80oC. These

observations showed that, hydrolytic enzymes in malted acha worked best at milder temperatures

and became inactivated, with a reduction in their activity, at higher temperatures during the

enzymatic hydrolysis. This observation was possibly due to the denaturation of the hydrolytic

enzymes present in the malted acha at high temperatures. Ayernor et al (2006) and Carter et al

(1989) reported that, above the optimum temperature of hydrolysis, the higher the temperature of

hydrolysis, the lower the amount of glucose obtained and vice versa. Also, Ayoola et al (2012),

observed a similar trend, where the optimum temperature for the hydrolysis of both cassava and

sorghum starch by alpha amylase was 60oC. They reported that there was a decline in glucose

production as the temperature was increased from 60oC to 80

oC, then to100

oC. Our findings in

the present study agree with these previously published works. The relatively mild optimum

56

temperature of 60oC for the hydrolysis of cassava starch with malted Acha is economically

favourable for its large scale production, since it will help to save and reduce energy cost.

The effect of time on the hydrolysis of cassava starch with the malted acha was also studied over

a time range of 1 – 7 h. The result, figure 14, showed a gradual increase in the concentration of

reducing sugars during the hydrolysis of the cassava starch with the malted acha as time

increased from 1 – 5 h after which there was a sharp decrease in the reducing sugar concentration

as the time increased to 7 h. High concentration of reducing sugar was recorded between 3 – 5 h,

with the maximum concentration of reducing sugars obtained at the 5 h interval. This

observation showed that time is an important factor in the hydrolysis of cassava starch by malted

acha. Akin-Osanaiye et al (2005) reported that duration of hydrolysis significantly affected the

amount of reducing sugar released during the saccharification of Carica papaya by Aspergillus

niger. In their report the amount of reducing sugar released with time increased till after 48h

after which there was a decrease in the amount of reducing sugar released as time increased.

Afiukwa et al (2009) also did observe that time had effect on the hydrolysis of starch by crude

extract from partially germinated mango seeds. The optimum time (5 h) for the hydrolysis of the

cassava starch by the malted acha as observed from the present study, is relatively short and this

can be advantageous in large scale industrial starch hydrolysis.

The optimal conditions for the hydrolysis of starch by hydrolytic enzymes depends on the

sources of these enzymes, substrate concentration, amount of enzyme present and other

molecules present in the solution. Hammond and Ayernor (2000) studied the optimum pH and

temperature conditions for the hydrolysis of cassava starch by malted cereals prepared from rice,

maize, sorghum and millet. They reported that high amount of sugar was produced by malted

maize, sorghum and rice at pH 5.0, while that of malted millet was at pH 5.5. They also reported

an optimum temperature of 60oC for malted millet, maize and sorghum, while that of rice was

55oC. Ocloo and Ayernor 2010 obtained 50

oC as the optimum temperature for the hydrolysis of

cassava flour by malted rice. For malted barley, optimal pH for its amylase activity has been

reported as from 4.5 – 7.0, and its optimum temperature reported as between 60-70oC. Egwim

and Oloyede (2006), reported an optimum temperature of 70oC for the amylase enzyme in the

malted acha they studied. Afiukwa et al (2009) reported an optimum temperature of 60oC and pH

6.0 for the hydrolysis of starch by amylase crude extract from partially germinated mango seed,

57

while Adebiyi et al (2005) reported an optimum temperature of 55oC and pH 5.0 for the

hydrolysis of sorghum bicolor starch with amylase from Rhizopus spp. Optimal temperature for

alpha amylase has been reported from 40 – 70oC (Haq et al., 2010). In the present study,

optimum temperature of 60oC and pH of 4.5 were observed for the hydrolysis of cassava starch

by malted acha. Subsequently, optimal conditions of starch to malted acha concentration,

temperature (oC), pH, and time (h) were employed in the hydrolysis of cassava starch by malted

acha and a cassava starch hydrolysate (CSH) having a reducing sugar content of 5.4% was

obtained.

The effect of different concentrations of yeast (5 %, 10 %, 15 %, 20 % and 25%) and time on

ethanol yield during the fermentation of cassava starch hydrolysate (CSH) was determined over a

period of 9 days, at a temperature between 28oC – 30

oC. Saccharomyces cerevisae (Baker‟s

yeast) was used and the ethanol concentration (yield) of each fermentation media was

determined after each day. The result obtained (figure 15) indicated that nearly equal amounts of

ethanol were produced by the various fermentation media with different concentrations of yeast,

at the end of the 9-day fermentation period. This observation supports the fact that yeast

concentration did not affect the maximum ethanol yield significantly (p< 0.05). A similar

observation was made by Ocloo and Ayernor (2010), who used yeast concentrations ranging

between 5- 20% and obtained nearly equal amounts of ethanol from the different fermentation

media after five days of fermentation. The yeast concentration, however, significantly (p< 0.05)

affected the time for fermentation to be completed, i.e. to achieve maximum ethanol yield.

Similar observation was also made by Ocloo and Ayernor (2010) in their work. Also, this

observation agrees with the findings of Akin-Osanaiye et al (2005), Morris and Sarad, (1990)

who reported that amount of yeast influenced ethanol production by reducing the time of

maximum ethanol production.

However, the result showed that time had a significant (p< 0.05) effect on the ethanol yield by

the various fermentation media. The graph of the mean ethanol concentration against time as

shown in figure 16 gave a hyperbolic curve. The hyperbolic nature of the curve indicates that

there are some inhibitory actions occurring after the 3rd

day of fermentation and this could be as

a result of the inhibition of the yeast by its product (ethanol) and other fermentation by products.

Gutcho (1973) reported that high alcohol content in a fermenting medium slowed down or

58

stopped the growth of yeast. A positive correlation (r = 0.875, R2 = 0.765) was obtained between

yeast concentration and time on the amount of ethanol produced. The maximum ethanol

concentration obtained after the 9-day fermentation was 8.11 (g/100ml), while over 95% of this

maximum ethanol concentration was obtained by nearly all the fermentation media at the end of

the fermentation period.

Having established the relationship between yeast concentration and time on ethanol yield, a

yeast concentration of 20% was used to ferment cassava starch hydrolysate with 5.4% reducing

sugar content for six days, to obtain appreciable ethanol yield of 7.42 (g/100ml). The implication

of this is that time and money were saved and cost of ethanol production reduced. At the end of

the 6-day fermentation period, the ethanol produced in the fermentation medium was distilled

using simple distillation setup at temperatures of between 78oC – 80

oC. Qualitative test on the

distillate was positive for ethanol and the ethanol was stored in a plastic container.

4.2 Conclusion

From this research work, it was evident that malted acha (Digitaria exilis), though being a

relatively unknown crop, has hydrolytic potentials just like other malted cereals, which can be

employed by Nigeria in its different sugar-based industries, breweries and distilleries. It was also

observed that malted acha has mild optimal conditions of temperature, pH and time at which it

works best and this could be of great economic benefit in its application in large scale starch

hydrolysis for bioethanol production and other purposes. Since very little is known about acha,

further research needs to be carried out on the development of new breeds of this plant with

better amylolytic potentials.

59

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