1 i

TITLE PAGE

PRODUCTION AND CHARACTERIZATION OF PECTINASES OBTAINED FROM

ASPERGILLUSNIGER UNDER SUBMERGED FERMENTATION SYSTEM USING PECTIN

EXTRACTED FROM ORANGE PEELS AS CARBON SOURCE.

A PROJECT WORKSUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS

FOR THE AWARD OF DEGREE OF MASTER OF SCIENCE (M.Sc) IN BIOCHEMISTRY

(INDUSTRIAL BIOCHEMISTRY AND BIOTECHNOLOGY), UNIVERSITY OF NIGERIA,

NSUKKA

BY

EZIKE, TOBECHUKWU CHRISTIAN

(PG/M.Sc/10/52393)

DEPARTMENT OF BIOCHEMISTRY

UNIVERSITY OF NIGERIA

NSUKKA

SUPERVISORS: PROF. F.C. CHILAKA AND DR. S.O.O EZE

JUNE, 2012

CHAPTER ONE

2

1.0 INTRODUCTION

Orange fruit (Citrussinensis) is one of the most important citrus crops grown all over the

world especially in the United States, Brazil and China. These countries represent more than

two third of global citrus fruit production (UNCTAD, 2005). Orange fruit is highly relished

as a fresh fruit owing to its rich organoleptic and thirst quenching properties and has a high

appeal. It is extensively used in food industries for fruit juice production. Processing and

utilization of orange fruits into various products eventually leads to generation of waste in

form of peels, pulp and seeds. The waste has about 66 million tons annual production and a

huge amount of it is discarded to nature causing serious environmental problems (Pourbafrani

et al., 2007; Tripodo et al., 2004). Orange waste is conventionally bio-transformed

anaerobically into humus, although many valuable by-products can be produced from the rich

waste. In other words, wealth can be derived from this waste by value addition and products

such as pectin, peel oil, dietary fibres and predominantly pectinases can be easily harnessed

(Bali, 2003).Of these products, pectin and pectinases have a wide global market.

Pectins are high molecular weight acid polysaccharide primarily made up of α (1→4) linked

D-galacturonic acid residues. They occur as structural polysaccharides in the middle lamella

and primary cell walls of young plant cells (Kashyap et al., 2001), where they contribute to

the firmness and structure of plant tissues (Sathyanarayana and Panda, 2003). Pectinases are

responsible for the degradation of pectins. These enzymes are classified based on their

preferred substrate (pectin, pectic acid or oligo-D-galacturonate), the degradation mechanism

(transelimination or hydrolysis) and the type of cleavage (random (endo) or terminal (exo)

(Kashyap etal., 2001).Pectinase has wide industrial application in extraction, clarification,

filtration and depectinization of fruit juices and wines in food and wine industries. It is also

used in textile industries for treatment of natural fibres and degumming of texture fibres

(Molina et al., 2008). They have also been reported to work on purification of viruses

(Salazar and Jayasinghe, 1999) and in making of paper (Beg et al. 2003).

Microbial pectinases account for 25% of the global food enzymes sales (Singh etal., 1999),

and are widely accepted as the best sources for the production of enzymes from agro-wastes.

Some bacteria (Bacilluslicheniformis, Aeromonascavi, Lactobacillus etc.)are known to

produce industrial enzymes but filamentous fungi are desired for the production of enzymes

because their nature is generally regarded as safe (GRAS)by United States Food and Drugs

Administration (USFDA) and are employed in food industry (Sumantha etal., 2005).

3

Recently, the production of pectinases from agro-wastes by fungi has been described as more

attractive (Sebastian etal., 1996; Acuna-Arguelles etal., 1995).

Fungi canproduce both intracellularas well as extracellular enzymes. All fungi are

heterotrophic, and rely on carbon compounds synthesized by other living organisms. Small

molecules like mono, disaccharides, fatty acids and amino acids can easily pass through but

for breaking down of larger complex compounds like pectin, fungi secrete extracellular

enzymes. The extracellular enzymes are easier to be extracted than intracellular enzymes

which require more time and costly chemicals for extraction(Hankin and Anagnostakis,

1975). The advantage of using microorganisms for the production of enzymes is that, they are

not influenced by climatic and seasonal factors, and can be subjected to genetic and

environmental manipulations to increase the yield.

Pectinases can be produced by both submerged and solid-state fermentation (SSF) as well as

many other enzymes (Murad and Foda, 1992). Submerged fermentation(SmF) is the

cultivation of microorganisms on liquid broth. It requires high volumes of water, continuous

agitation and generates lot of effluents. SSF incorporates microbial growth and product

formation on or within particles of a solid substrate under aerobic conditions, in the absence

or near absence of free water, and does not generally require aseptic conditions for enzyme

production. Pectinolytic enzyme synthesis is highly influenced by carbon and nitrogen

sources (De Gregorio et al., 2002), presence of pectin (Solis-Pereira et al., 1993), pH

(Yogesh, et al., 2009) and temperature (Bailey, 1990). Therefore, the advantage of SmF is

that physicochemical properties such as pH, temperature and oxygen tension are easier to

control than in solid state fermentation (Canel and Moo-Young, 1980; Costa etal., 1998;

Castilho etal., 2000).

Orange peels also hold a promising substrate for pectinase production because they contain

appreciable amount of pectin (Table 1) which could serve as the carbon source for the

production of pectinase through microbial system (Dhillon, et al., 2004).As pectin is the ideal

substrate for production of pectinases, it was thought that attempts should be made to extract

pectin from orange peels and also isolate potential pectinolyticfungi from natural sources and

employ them in pectinase production.

Table 1: Typical levels of pectin in plants

Fruits % of Pectin

4

(Wikipedia, 2012a)

1.1 Description of Orange Fruit

Orange (Citrussinensis) belongs to citrus fruits and is believed to have originated from Asia

(Beaven et al., 1972). It is the most commonly grown tree fruit in the world and is widely

cultivated in tropical and subtropical climates. The fruit is commonly peeled and eaten fresh,

or squeezed for its juice. It has a thick bitter rind that is usually discarded. A cross-section of

orange fruit shows 3 different layers:

1. A rough, robust and bright color (from yellow to orange) skin or rind, known as

epicarp or flavedo, which covers the fruit and protects it from damage. Its glands

contain the essential oils that give the fruit its typical citrus fragrance.

2. A white, thick and spongy mesocarp or albedo, which together with the epicarp forms

the pericarp or peel of the fruit.

3. The internal part that makes the pulp. It is divided into individual segments or juice

sacs (with or without seeds, according to varieties) by a thick radial film or endocarp.

This part is rich in soluble sugars, significant amounts of vitamin C, pectin, fibres,

different organic acids and potassium salt, which give the fruit its characteristic citrine

flavor.

1.1.1 Scientific Classification of Orange Fruits

(fresh weight)

Apple

Apricot

Cherries

Oranges

Carrot

Citrus peels

1 – 5.5

1

0.4

0.5 – 3.5

1.4

30

5

Kingdom Plantae

Division Magnoliophyta

Class Magnoliopsida

Order Sapindales

Family Rutaceae

Genus Citrus

Specie sinensis

Botanical name Citrus sinensis

1.1.2 Production of Orange Fruits

Citrus fruits are produced all around the world. According to FAO (2004) data, 140 countries

produced citrus fruits. However, most production is concentrated in certain areas. Main citrus

fruit producing countries are Brazil, the Mediterranean countries, the United States (where

citrus fruits for consumption as fresh fruit are mainly grown in California, Arizona and

Texas, while most orange juice is produced in Florida) and China. These countries represent

more than two thirds of global citrus fruit production (UNCTAD, 2005). Almost 99% of the

fruit from Brazil is processed for export; it is the overwhelming giant in worldwide orange

juice production(Wikipedia, 2012b).

Production of orange juice between Brazil and Florida make up roughly 85% of the world

market. Brazil exports 99% of its production, while 90% of Florida's production is consumed

in the US. Orange juice is traded internationally in the form of frozen concentrated orange

juice to reduce the volume used, so that storage and transportation costs are lower

(Wikipedia, 2012b).Total annual citrus production was estimated at over 105 million tons in

the period 2000-2004. Oranges constitute the bulk of citrus fruit production, accounting for

more than half of global citrus production in 2004. The rise in citrus production is mainly due

to the increase in cultivation areas and the change in consumer preferences towards more

health and convenience food consumption and the rising incomes (UNCTAD, 2005).

1.1.3 Nutrient and Phytochemical Content of Orange Fruit

Citrus fruits and juices serve as primary sources of our daily requirement of Vitamin C.

However, like most other whole foods, citrus fruits also contain an impressive list of other

essential nutrients, including glycaemic and non-glycaemic carbohydrate (sugars and fibre),

6

potassium, folate, calcium, thiamin, niacin, vitamin B6, phosphorus, magnesium, copper,

riboflavin, pantothenic acid and a variety of phytochemicals (Table 2). In addition, citrus

contains no fat or sodium and, being a plant food, no cholesterol. The average energy value

of fresh citrus is also low, which can be very important for consumers concerned about

putting on excess body weight. For example, a medium orange contains 60 to 80 kcal, a

grapefruit 90 kcal and a tablespoon (15 ml) of lemon juice only 4 kcal (Whitney and Rolfes,

1999).

Table 2: Nutrient and Phytochemical Contents of Citrus Fruits

Percentages are relative to US recommendation for adults.

(USA Nutrient Database, 2012)

Nutrient / Phytochemical Nutritional value per 100g

(3.5oz)

Energy

Carbohydrates

- Sugars

- Dietary fibre

Fat

Protein

Thiamine (vit. B1)

Riboflavin (vit. B2)

Niacin (vit. B3)

Pantothenic acid (B5)

Vitamin B6

Folate (vit. B9)

Vitamin C

Calcium

Iron

Magnesium

Phosphorus

Potassium

Zinc

192 kJ (46 kcal)

11.54 g

9.14 g

2.4 g

0.21 g

0.70 g

0.100 mg (9%)

0.040 mg (3%)

0.400 mg (3%)

0.250 mg (5%)

0.051 mg (4%)

17 μg (4%)

45 mg (54%)

43 mg (4%)

0.09 mg (1%)

10 mg (3%)

12 mg (2%)

169 mg (4%)

0.08 mg (1%)

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1.2 Pectic Substances

Pectic substance (also known as pectin) is the generic name used for the compounds that are

acted upon by the pectinolytic enzymes or pectinases. They are complex heterogeneous and

structural polysaccharides found in the primary cell wall and middle lamella of fruits and

vegetables where they function as hydrating agent and cementing materials of the cellulosic

network (Favella-Torres etal., 2006; Jarvis etal., 2003). They are largely responsible for the

structural integrity and cohesion of plant tissues (Alkorta et al., 1998). They are often

generally referred to as pectin.

Pectic substances are synthesized in the Golgi apparatus from UDP-D-galacturonic acid

during early stages of growth in young enlarging cell walls (Sakai et al., 1993). Lignified

tissues have a low content of pectic substances when compared with young, actively growing

tissues. The content of the pectic substances is also very low in higher plants usually less than

1%. They are mainly found in fruits and vegetables, constitute a large part of some algal

biomass (up to 30%) and occur in low concentration in forestry or agricultural

residues.

Figure 1: Schematic structureof the four different types of pectic polysaccharides

(Yadav et al., 2009).

8

1.2.1 Structure of Pectic Substances

Pectic substances are present in various forms in plant cells and this is the probable reason for

the existence of various forms of pectinolytic enzymes. Therefore, various pectic

polysaccharides have been detected in the cell wall, including homogalaturonan (HGA),

xylogalacturonan (XGA), rhamnogalacturonan 1 (RG1) and rhamnogalacturonan 11 (RG11)

(Figure 1) (Yadav et al., 2009).

In the plant cell wall, the ratio between the four polysaccharides, HGA, XGA, RG1 and

RG11 is variable, but typically HGA is the most abundant polysaccharide, consisting about

65% of the wall pectin, while RG1 constitute 20% to 35% (Mohnen, 2008). XGA and RG11

are minor component each, constituting less than 10% (Zandleven et al., 2007; Mohnen,

2008). Hence, HGA and RG1 are more abundant than the other components. These different

pectic polysaccharides are not separate molecules but covalently linked together.

1.2.1.1 Homogalacturonan(HGA)

Homogalacturonan HGA is a linear polymer formed by D-galacturonic acid which can be

acetylated and/or methyl esterified and is thought to contain 100 - 200 galacturonic acid

(GalA) residues (Zhang, 2006). HGA backbone is modified by esterification at C-6 carboxyl

position and /or O-acetylation O-2 or O-3 position (Ishii, 1995; Ishii, 1997). The degree of

methyl and acetyl esterification is variable and affects the physiochemical properties of the

pectin especially the formation of calcium-mediated interactions between HGA chains

(Liners et al., 1992).

1.2.1.2 Rhamnogalacturonan 1 (RG1)

It contains a backbone of the repeating disaccharide, α-(1→4) D-galacturonic acid and α-

(1→2)-L-rhamnose (Cosgrove, 1997). The predominant side chain contains linear and

branched α-L-arabinan and/or β-D-galactan linked to the C-4 atom of some of the rhamnose

residues (Sharma et al., 2006). Rhamnose (Rha) is a minor component of the pectin backbone

and introduces a kink into the straight chain (Figure 2) and other sugars such as arabinose,

galactose and xylose occur in the side chains (Oakenful, 1991). Some of the rhamnose

residues may also be O-acetylated at C-2 and or C-3 (Brent et al., 2001).

9

Figure 2: Schematic diagram showing how rhamnose (Rha) insertions cause kinking of

galacturonic acid (GalA) chain; S = neutral sugars(Sriamornsak, 2002).

1.2.1.3 Rhamnogalacturonan II (RGII)

Despite its name, RGII is a homogalacturonan chain with complex side chains attached to the

galacturonic residues (Willats et al., 2006). RGII is present in the primary cell walls as a

dimer that is mediated/cross-linked by a borate-diol ester which ensures the integrity of the

cell wall (O‟Neill et al., 2001). RG11 is not structurally related to RG1, as its backbone

contains stretches of at least seven 1,4 linked α-D-galacturonic acid residues than the

repeating disaccharides α-(1→2)-L-rhmnosyl-α-(1→4)-D-galacturonsyl found in RG1 (Brent

et al., 2001). In fact, clusters of complex side chains attached onto O-2 or O-3 position in the

galacturonan backbone give rise to RG11. These side chains are composed of 12 types of

glycosyl residues linked together by at least 22 different glycosidic bonds (Harholt et

al.,2010). Some of the glycosyl residues and glycosidic linkages found in RG11 side chains

are rare and considered unique in plant polysaccharides (e.g 2-O-methyl-L-fucose, L-aceric

acid, and α-1,3-xylofuranose) (Figure 1)

Conventionally, HG can be referred to as the smooth region of pectin molecule because it

consists of linear chain of homogalacturonan while RGI and RGII can also be called the hairy

region because of their branched chain network of rhamnogalacturonan 1 (RG1),

rhamnogalacturonan 11 (RG11 and another pectic structure, xylogalacturonan (XGA)

(Vincken et al., 2003).

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Figure 3: The structure of primary cell wall(Carpita and Gibeaut, 1993).

1.2.2 Classification and Nomenclature of Pectic Substances

The American Chemical Society classified pectic substances into four main types as reported

by Alkorta et al., (1998) as follows:

a) Protopectin; which is the water insoluble pectic substances present in intact tissue.

Pectic substances are found in the form of protopectin in plant cells (Luzio, 2004). On

restricted hydrolysis, protopectin yields pectin or pectic acids Protopectin is bound to

cellulose microfibrils conferring rigidity on cell walls. During ripening the fruit

enzymes alter the pectin structure by breaking the pectin backbone or side chains,

resulting in a more soluble molecule (Kashyap et al., 2001)

b) Pectic acid; which is the soluble polymer of galacturonans that contains negligible

amount of methoxy groups. Normal or acid salt of pectic acid are called pectates.

c) Pectinic acids; which are the polygalacturonan chain that contain less than 75%

methylated galacturonate units. Normal or acid salts of pectinic acid are called

pectinates.

d) Pectin; also called polymethylgalacturonate, is the polymeric material in which at

least, 75% of the carboxyl groups of the galacturonate units are esterified. It

represents the complex pectic substance extracted from plant and fruit walls which are

utilized for biochemical studies.

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1.2.3 Types of Pectin

Pectins are classified according to its degree esterification. They include: High methyl ester

(HM) pectin, low methyl ester (LM) pectin and amidated pectin.

1.2.3.1 High Methyl Ester (HM) Pectin

Pectin as extracted normally has more than 50% of the acid units esterified, and is classified

as high methyl ester (HM) pectin. The percentage of ester groups is called degree of

esterification (Figure 4).

Figure 4: HM pectin formular(IPPA, 2001).

1.2.3.2 Low Methyl Esther (LM) Pectin

When the high methy esters are modified at the extraction process, or continued acid

treatment, low methyl ester (LM) pectin will be formed. It has less than 50% methyl ester

group (Figure5).

Figure 5: LM pectin formular(IPPA, 2001).

1.2.3.3 Amidated Pectins

Some pectins are treated during manufacture with ammonia to produce amidated pectins,

which have particular advantages in some applications. (Figure 6)

Figure 6: Amidated Pectin(IPPA, 2001)

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1.2.4 Sources of pectin

Pectin is present in all plant but the content and composition varies on the specie, variety,

maturity of the plant, plant part, tissue and growing condition. Itis abundantly present in

apple, lemon, orange, mango, tomato, beet, and carrots (Pilnik and Voragen, 1993;

Girdharilal et al., 1998) (Table 3), especially in the peels of these fruits (Table 4). Pectin is

higher in legumes and citrus fruits than cereals.

Table 3: Sources of Pectin

(Pilnik and Voragen, 1993; Girdharilal et al, 1998)

1.2.5 Production of pectin

The first commercial production of a liquid pectin extract was recorded in 1908 in Germany,

and the process spread rapidly to the United States. This was followed by a rapid growth of

the pectin industry in the United States, and also somewhat later in Europe.

In recent years, the centre of production has moved to Europe and to citrus-producing

countries like Mexico and Brazil. Commercial pectins are almost exclusively derived from

citrus peel or apple pomace, both by-products from juice (or cider) manufacturing. Apple

pomace contains 10-15% of pectin on a dry matter basis. Citrus peel contains 20-30% (May,

1990). From an application point of view, citrus and apple pectins are largely equivalent.

Alternative sources include sugar beet waste from sugar manufacturing, sunflower heads

(seeds used for edible oil), and mango waste (Rolin, 1993).

Material % pectin in fresh material %pectin in dry wt basis

Apple Pomace

Lemon Pulp

Orange Pulp

Beet Pulp

1.5-2.5

2.5-4.0

3.5-5.5

1.0

15-18

30-35

30-40

25-30

13

Table 4: Pectin yield from various sources with respective Methoxylated and De-

esterified pectin

Source of pectin Yield (%) MeO (%) DE (%)

Apple pomace

Lime peels

Lemon peels

Sweet orange

Mandarin orange peels

17.0

32.0

27.7

17.8

18.4

8.9

8.6

9.2

7.7

9.5

74.9

63.2

73.4

57.0

64.9

(Rao and Maini, 1999)

1.2.6 Extraction of pectin

Commercially, pectin is extracted by treating the raw material with hot dilute mineral acid at

pH about 2. The precise length of extraction time varies with raw material, the type of pectin

desired, and from one manufacturer to another. The hot pectin extract is separated from the

solid residue as efficiently as possible. This is not easy since the solids are by now soft and

the liquid phase are viscous. The viscosity increases with pectin concentration and molecular

weight. There is a compromise between efficient extraction and solids separation and

operating cost. The pectin extract may be further clarified by filtration through a filter aid.

The clarified extract is then concentrated under vacuum.

Powdered pectin can be produced by mixing the concentrated liquid from either apple or

citrus with an alcohol (usually isopropanol). The pectin is separated as a stringy gelatinous

mass, which is pressed and washed to remove the mother liquor, dried and ground. This

process yields pectin of around 70% esterification (or methoxylation). To produce other

types, some of the ester groups must be hydrolysed. This is commonly carried out by the

action of acid, either before or during a prolonged extraction, in the concentrated liquid, or in

alcoholic slurry before separation and drying. This process can produce a range of calcium

reactive low methoxyl pectins. Hydrolysis using ammonia results in the conversion of some

of the ester groups into amide groups, producing „amidated low methoxyl pectins‟ (May,

1990).

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1.2.7 Properties of Pectin

1.2.7.1 General properties of pectin

1. Pectins are soluble in pure water.

2. Monovalent cation (alkali metal) salts of pectinic and pectic acids are usually soluble

in water; di- and trivalent cations salts are weakly soluble or insoluble.

3. Dry powdered pectin, when added to water, has a tendency to hydrate very rapidly,

forming clumps. Clump formation can be prevented by dry mixing pectin powder

with water-soluble carrier material or by the use of pectin improved dispersibility

through special treatment during manufacturing (Rolin, 1993; Hercules Incorporated,

1999).

1.2.7.2 Gel formation properties of pectin

The most important use of pectin is based on its ability to form gels. It has been suggested by

Oakenfull, (1991) that hydrogen bonding and hydrophobic interactions are important forces

in the aggregation of pectin molecules. Gel formation is caused by hydrogen bonding

between free carboxyl groups on the pectin molecules and also between the hydroxyl groups

of neighboring molecules.

In a neutral or only slightly acid dispersion of pectin molecules, most of the unesterified

carboxyl groups are present as partially ionized salts. Those that are ionized produce a

negative charge on the molecule, which together with the hydroxyl groups causes it to attract

layers of water. The repulsive forces between these groups, due to their negative charge, can

be sufficiently strong to prevent the formation of a pectin network.

When acid is added, the carboxyl ions are converted to mostly unionized carboxylic acid

groups. This decrease in the number of negative charges not only lowers the attraction

between pectin and water molecules, but also lowers the repulsive forces between pectin

molecules. Sugar further decreases hydration of the pectin by competing for water. These

conditions decrease the ability of pectin to stay in a dispersed state. When cooled, the

unstable dispersing of less hydrated pectin forms a gel, a continuous network of pectin

holding the aqueous solution.

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1.2.8 Uses of Pectin

1.2.8.1 Use of Pectin in Food Industry

In food industries, pectins are used as gelling, thickening or stabilizing agent to produce

confectioneries such as marmalade and jelly candies, fruit and berry preparations such as

fillings, fruit juices and jams; and fermented dairy products such as yoghurts and fruit

flavored milk deserts (Girdharilal et al., 1998). Other applications of pectin include use in

edible films, paper substitute, foams and plasticizers (Thakur et al., 1997).

1.2.8.2 Use of Pectin in Pharmacy and Medicine

Pectins in the diet of humans and laboratory animals have been shown to lower cholesterol

levels by raising the excretion of fecal bile acids and neutral sterols (Bali, 2003).

Administration of 15g/day of pectin for three weeks may result in a mean 13% reduction in

plasma cholesterol levels (Sharma et al., 2006). Pectin is a dietary fibre that functions in

mineral and ion absorption and exchange. Hence, it is effective in removing toxic cations like

lead and mercury from the gastro-intestinal tract and respiratory organs (Kohn, 1982). A

combination of pectin and their colloids can be used to treat diarrhoeal diseases and

constipation (Sharma et al., 2006).

In addition, pectin hydrogen is used in tablet formulation, owing to its binding ability and in

controlled-release matrix tablet formulations (Slany et al., 1981; Naggar etal., 1992). Also

the potential of pectin or its salt as a carrier for colonic drug delivery has been demonstrated

by Ashford et al. (1993) and Rubinstein et al. (1993).

1.3 Pectinases

Pectinases are group of enzymes that catalyze the breakdown of pectin or pectic substance.

They are also referred to as pectic enzymes or pectinolytic enzyme. These enzymes attack

pectin and deploymerize it by hydrolysis and transelimination as well as by de-esterification

reactions, which hydrolyses the ester bond between carboxyl and methyl groups of pectin

(Ceci and Loranzo, 1998). This process is called pectolysis. Pectolysis is one of the most

important processes for plant, as it plays a role in cell elongation and growth as well as fruit

ripening.

Pectinolytic enzymes are wide spread in nature and are produced by bacteria, fungi,

yeast, insects, nematodes and protozoa. For example bacteria like Bacillusspecies,

16

Clostridiumspecies, fungi like Aspergillusspecies, Penicillumspecies, yeast like

Saccharomyces, Candida etc. Microbial pectolysis is important in plant pathogenesis,

symbiosis and decomposition of plant deposits (Lang and Dornenburg, 2000). Thus by

breaking down pectin polymer for nutritional purposes, microbial pectinolytic enzymes play

important role in nature. These enzymes are inducible i.e. produced only when needed and

they contribute to the natural carbon cycle.

Microbial pectinolytic enzymes are not only enzymes available to attack plant

polysaccharides. However, pathogenic attack on plant tissue is normally initiated by pectic

enzymes because pectic substances are most readily accessible. Other carbohydrate enzymes

appear sequent and attack the available polysaccharides. Final result is a sequence of

appearance of microbial carbohydrate enzymes during microbial attack on plant cell walls

(Sakai et al., 1993).

1.4 Classification of Pectinases

Pectinases are classified according to their mode of secretion as extracellular or intracellular

pectinases. An extracellular enzyme is excreted outside the cell into the medium in which that

cell is living. Extracellular enzymes usually convert large substrate molecules into smaller

molecules that can then be more easily transported into the cell, whereas an intracellular

enzyme operates within the confines of the cell membrane (Bail, 2003).

Both extracellular and intracellular pectinases are classified into three broader groups based

on their mode of action on pectic substances to release different products (Sakai, 1992;

Palomaki and Saarilahti, 1997). The three classes include:

1. Protopectinases

2. Pectin esterases

3. Depolymerases

Depending upon the pattern of action, i.e. random or terminal, these enzymes are termed as

Endo or Exo enzymes, respectively (Table 5).

1.4.1 ProtopectinasesProtopectinases or pectinosinases catalyze the breakdown of

insoluble protopectin to form highly polymerized soluble pectin (Jayani et al., 2005). The

mode of reaction catalyzed by pectinosinase (Ppase ) is as shown in Reaction 1

Insoluble Protopectin + H2O →Soluble Pectin ------------------------- Reaction 1

17

Table 5: Classification of Pectinases

( Jayaniet al., 2005).

1.4.1.1 Classification of Protopectinases

There are two types of protopectinases (Ppase): A-type and B-type (Sakai et al., 1984). The

A-type reacts on the polygalacturonic acid region (inner site) of protopectin, whereas the B-

type reacts on the polysaccharide chains (outer site) that connect the polygalacturonic acid

chain and cell wall constituents (Jayani et al., 2005).

Types of Pectinases Enzyme

Code

number

Substrate Mode of

Action

Products

1.Protopectinases (Ppase)

a. A-type Ppase

b. B-type Ppase

Insoluble

protopectin

Insoluble

protopectin

Hydrolysis of

inner site.

Hydrolysis of

outer site.

Soluble pectin.

Soluble pectin.

2. Pectin Esterases (PE)

a. Pectin methylesterase

b. Pectin acetylesterase

3.1.1.11

3.1.1.6

Pectin

Pectin

Hydrolysis

Hydrolysis

Pectic acid + methanol

Pectic acid + ethanol

3. Depolymerases

a. Hydrolases

i. Endopolygalacturonase

ii. Exopolygalacturonase

iii.Endopolymethylgalacturonase

iv.Exopolymethylgalacturonase

b. Lyases

i. Endopolygalacturonate

(Endopectate) lyase

ii. Exodopolygalacturonate

(Exopectate) lyase

iii.Endomethylpolygalacturonate

(Endopectin) lyase

iv.Exomethylpolygalacturonate

(Exopectin) lyase

3. 2. 1. 15

3. 2. 1. 67

3. 2. 1. 41

-

4. 2. 2. 2

4. 2. 2. 9

4. 2. 2. 10

-

Pectic acid

Pectic acid

Pectin

pectin

Pectic acid

Pectin acid

Pectin

Pectin

Hydrolysis

Hydrolysis

Hydrolysis

Hydrolysis

Transelimination

Transelimination

Transelimination

Transelimination

Oligogalacturonates

Monogalacturonates

Oligomethylgalacturonates

Monomethylgalacturonates

Unsaturated

oligogalacturonates

Unsaturated digalacturonates

Unsaturated

methyloligogalacturonates

Unsaturated

methylmonogalacturonates

18

1.4.1.2 Occurrence of Protopectinases in Organisms

The presence of pectinosinases (Ppase) has been reported in fungi, yeast and Bacillus species.

The A-type Ppases have been isolated in the culture filtrates of yeast and yeast-like fungi

(Whitaker, 1990), whereas the B-type Ppase have been identified in the culture filtrate of

wide range of Bacillus species (Sakai, et al., 1984). In addition, the A-type Ppases have been

isolated from Kluyveromyces fragilisIFO 0288, Galactomyces reeseiL. and

Trichosporonpenicillantum and are referred to as Ppase-F, -L and -S, respectively (Whitaker,

1990). Also the B-type Ppases have been found in Bacillus subtilisIFO 12113,

BacillussubtilisIFO 3134 and Trametes species and are referred to as Ppase-B, -C and -T,

repectively (Jayani et al., 2005).

1.4.1.3 Determination of Protopectinase Activity

Protopectinase activity is assayed by measuring the amount of pectic substances released

from protopectin by carbazole-sulphuric acid method (Siebert and Anto, 1946). One unit of

Ppase activity is defined as the enzyme that liberates pectic substance corresponding to one

micromole (1µmole) of D-galacturonic acid per milliliter of reaction mixture under the assay

condition. The pectin concentration is measured as D-galacturonic acid from its standard

curve (Jayani et al., 2005).

1.4.1.4 Biochemical and Physicochemical Properties of Protopectinases

The F-, L- and S- forms of A-type Ppases are similar in biological properties and have similar

molecular weight of 30kDa (Jayani et al., 2005). Ppase-F is an acidic protein, while Ppase -L

and -S are basic proteins. The three enzymes, having pectin-releasing effects on protopectin

of various sources, catalyze the hydrolysis of polygalacturonic acid and decrease the

viscosity, thereby increasing the reducing value of the reaction medium containing

polygalacturonic acid.

Conversely, Ppase-B, -C and -T have molecular weights of 45, 30 and 55kDa, respectively

(Jayani et al., 2005). Ppase-B and -C have an isoelectic point (pI) of 9.0 while Ppase-T has a

pI of 8.1 (Sakai, 1992). However, the three enzymes act on protopectin from various citrus

peels and other plant tissues liberating soluble pectin (Sakai, 1992). In addition, the optimal

pH for protopectinase activity is within the range of 3.5 to 4.0 (Doby, 1965).

19

1.4.2 Pectin Esterases (PE)

Pectin esterases, formerly called the saponifying enzymes, catalyze the deesterification of

methyl or ethyl ester linkages of galacturonan backbone of pectic substances to release pectic

acids and methanol or ethanol respectively (Cosgrove, 1997; Yadav et al., 2009). The

resulting pectic substance from PE deesterification reaction is liable to attack by

polygalacturonase and lyase to release different products (Prade, et al., 1999).

1.4.2.1 Classification of Pectin Esterases

Pectin esterases are classified into two based on the type of alkyl group (methyl or acetyl

group) attached to the galacturonan backbone of the pectic substance they de-esterify. The

two classes include, pectin methylesterase (PME) (which is more common) and pectin acetyl-

esterase (PAE) (which is rare) (Yadav et al., 2009).

1.4.2.1.1 Pectin Acetylesterase (PAE)

Pectin acetylesterase (PAE) catalyzes the deesterification of ethyl ester linkages of

galacturonan backbone of highly acetylated pectin to release pectic acid and ethanol. In plant

tissues, acetyl esters are only very slowly deesterified (Deuel and Stutz, 1958) unlike the

plant methyl esters.

1.4.2.1.2 Pectin Methylesterase (PME)

PME is a specific enzyme that acts gradually, removing units of methanol and pectic acid

from the terminal pectin chains (Pilnik and Voragen, 1993). The mode of action of PME

varies depending on the origin of the enzyme. Pectin methylesterases of fungal origin act by a

multi-chain mechanism, removing the methyl groups at random, whereas pectin

methylesterases of plant origin act either on the non-reducing end or next the molecule by a

single chain mechanism (Jayani et al., 2005). Figure 7 shows the mode of action of PME.

20

Figure 7: Mode of Action of Pectin Methylesterase(Sathyanarayana and Panda, 2003).

1.4.2.2 Mechanism of Action of Pectin Esterase

Recently, the mechanism of pectin methylesterase has been illuminated by the crystal

structures of several catalytic mutants in complex with various substrates (Figure 8). Through

this elegant structural analysis, the nucleophilic aspartate was determined to be D199, which

attacks the carbonyl carbon of the C-6 ester and generates a tetrahedral intermediate (Fries

etal., 2007). The second proximal aspartate residue (D178) operates as the general acid-base

catalyst and forms a strong hydrogen bond with the carbonyl oxygen of the methyl ester. The

transition state is stabilized by the interactions formed on the carbonyl oxygen. Protonation of

the leaving group by D178 enables the release of methanol and the generation of covalently

bound anhydride intermediate (Abbott and Boraston, 2008). Subsequent hydrolysis of the

anhydride by D198 activated water molecule releases the aglycon group and regenerates the

active site.

The nucleophile (D199) attacks the carbonyl carbon, forming a tetrahedral intermediate that

is stabilized by Q177. The general acid-base catalyst D178 protonates the ester-linked

oxygen, and attack by a catalytic water releases methanol and polygalacturonate (Abbott and

Boraston, 2008). The majority of study on pectin methylesterase has focused on plant

enzymes that are operational during tissue development and fruit ripening (Prasanna etal.,

2007).

These enzymes operate in a processive fashion, in the same maner as bacterial enzymes to

produce blocks of demethylated subunits along the polysaccharide, for which both single-

chain and multi-chain mechanisms have been proposed (Grasdalen, et al, 1996). This process

contrasts with that for the fungal methylesterasees, which tend to demethylate pectin in

distinctive fashion by randomly selecting a substrate (van Alebeek etal., 2003).

Polygalacturonate

(Pectic Acid)

O O

OH

OH

OH

OH

COOCH3

O

O

O

COOCH3 Pectin

H

H

H H

H

H H

H

PME

PME

+ CH3OH

Methanol

O O

OH

OH

OH

OH

COOH

O

O

O

COOH

H

H

H H

H

H H

H

21

Figure 8.Generalized mechanism for demethylation of pectin by pectin

methylesterase.The nucleophile (D199) attacks the carbonyl carbon, forming a tetrahedral

intermediate that is stabilized by Q177. The general acid-base catalyst D178 protonates the

ester-linked oxygen, and attack by a catalytic water releases methanol and polygalacturonate,

recharging the active site(Abbott and Boraston, 2008).

1.4.2.3 Structure of pectin esterases

The crystal structure of pectin esterase contains amino acid sequence which is unrelated to

that of any other known protein (Figure 9). The enzyme adopts the parallel β-helix fold

described for both pectin lyase and polygalacturonase. Comparison of the tertiary structures

of these different enzyme classes indicates that the esterase is more structurally similar to

pectin lyases, in that it contains the same numbers of complete coils (eight) and β-sheets

(three) (Abbott and Boraston, 2008). The most noticeable difference in the enzyme is that the

T3 loops harnessing the putative catalytic site are shifted along the longitudinal axis of the

protein toward the C terminus. In addition, there is an extensive C-terminus tail with α-helical

character that packs antiparallel to the face of the β-helix (Abbott and Boraston, 2008).

The active site architecture of pectin esterase is unique and lacks the serine and histidine

residues of the Ser-His-Asp catalytic triad present in functionaly unrelated esterases (Jenkins

et al., 2001). A putative mechanism was originally predicated based upon structural analysis

(Johansson et al., 2002). The floor of the catalytic site is coated with aromatic residues:

Y158, Y181, F202, W269. These amino acid functions to dock the pectin substrate by

selectively stacking with the polar faces of individual residues. Of these, Y181, F202, and

W269 may be critical, as they are highly conserved among eukaryotic pectin esterase

(Jenkins et al., 2001). The de-esterification reaction is believed to be facilitated by two

aspartate residues (D178 and D199), which are positioned as suitable candidates for acid-base

catalysis (Figure 10). At their closet point the oxygen atoms from each carboxylate group are

within 4.2Åof each other, which is noticeably shorter than the 5.5Å typically observed in

retaining glycosidic hydrolases (GHs ) (Abbaott and Boraston, 2008).

22

Figure 9: The three dimensional structure of pectin metylesterase displayed in a

“cartoon” format with a transparent solvent-accessible surface (Abbott and Boraston,

2008).

Figure 10: The extracellular pectin metylesterase.Showing the active structure pectin

methylesterase displayed in wall-eyed stereo (Abbott and Boraston, 2008).

1.4.2.4 Occurrence of Pectin Esterase in Organisms

Activities of pectin esterases in cell wall metabolism including cell growth, fruit ripening,

abscission, ageing and pathogenesis have being reported by Gaffe et al. (1997) and Dorokhov

et al. (1999). Hence, PE is present in plant and also in plant pathogenic bacteria and fungi.

The enzyme have been identified in Rodotorula sp, Phytophthora infestans,

Erwiniachrysanthemi B 341, Saccharomyces cerevisae, Lachnospira pectinoschiza,

23

Pseudomonassolanacearum, Aspergillusniger, Lactobacillus lactissubsp. cremoris,

Penicillumfrequentans, E. chrysanthemi3604, Penicillumoccitanis, A. japanicus and so on

(Jayani etal., 2005).

There are also reports of PE occurrences in plants such as Caricapapaya (Innocenzo and

Lajalo, 2001), Citrussp (Arias and Burns, 2002), Pouteriasapota (Arenas-Ocampo et al.,

2003), Malpighia glabraL (Assis et al., 2004) and others. The enzyme has been found useful

in protecting and enhancing the firmness and texture of processed fruit juices and vegetables

as well as in the extraction and clarification of fruit juices (Fayyaz et al., 1993).

1.4.2.5 Determination of Pectin Esterase Activity

Lin et al. (1990) described a method for the determination of the methyl ester content of

pectin using the specific action of pectin esterase. The amount of NaOH consumed during the

enzyme reaction has been used for the assay comparable to acid-base titration used in

saponification reaction and it provides a simple rapid and selective procedure for measuring

the methoxyl content of pectin (Gummadi and Panda, 2003),

Also PE activity can be followed by gel diffusion assay described by Downie etal (1998).

Increased binding of ruthenium red to pectin, as the number of methyl ester attached to pectin

decreases is used in the assay. In addition, the activity of PE is highest on 65-75% methylated

pectin, since the enzyme is thought to act on methoxyl group adjacent to free carboxyl groups

(Whitaker, 1984).

1.4.2.6 Biochemical and Physicochemical Properties of Pectin Esterases

The activity of pectin esterases has a very little effect on viscosity of pectin containing

solutions unless divalent ions are present, which increase viscosity due to crosslinking

(Janyani et al., 2005). Pectin esterases are highly specific enzymes(Abbott and Borastan,

2008). Some PEs attack only ester groups next to a free carboxyl group (reducing chain) and

then continue to act along the molecules while others attack non-reducing end (Sakai et al.,

1993). The molecular weights of most PEs are in the range of 35-50KDa. pH values at which

PEs are active range from 4.0-8.0. Fungal pectin esterases have a lower pH optimum than that

of bacterial origin (Jayani etal., 2005). The author also reported that the optimum temperature

for maximal activity for majority of PEs ranges from 40-500C.

24

In addition, two different forms of pectin esterases, namely: PmeA, an extracellular enzyme

(Maldonaldo et al., 1998); and PmeB, an outer membrane protein (Shevchik et al., 1996),

have been isolated from Erwiniachrysanthemi3937and shows best activity at alkaline pH and

temperature of 50oC (Laurent et al., 2000).

1.4.3 Depolymerases

Depolymerases catalyze the hydrolysis of theα(1→4)-glycosidic bonds in the D-galacturonic

acid units of the pectic substances. These enzymes have been classified by Demain and Phaff

(1957) and Deuel and Stutz (1958) as glycosidases with specific activities pertaining to the

degree of esterification of the substrate and to random or terminal cleavage. Depolymerases

act on pectic substances by two different mechanisms: hydrolysis, in which they catalyze

hydrolytic cleavage with the introduction of water across the oxygen bridge and

transelimiation lysis, in which they break the glycosidic bond by a transelimination without

any participation of water molecule (Codner, 2001; Albersheim et al., 1960).

1.4.3.1 Classification of Depolymerases

Depolymerases are classified into hydrolases and lyases, depending on the preference of

enzyme for the substrate, the mechanism of cleavage and the splitting of the glycosidic

bonds. The hydrolases and lyases are further classified into endo-enzymes, and exo-enzymes,

depending upon their pattern of action which could be either random or terminal, respectively

(Table 5).

1.4.3.1.1 Hydrolases

Hydrolases catalyze the hydrolytic cleavage of pectic substances with the introduction of

water across the oxygen bridge to release free galacturonic acid and/or pectic substances of

lower molecular weights as end products. These enzymes comprise polygalacturonases and

polymethylgalacturonases that breakdown pectate and pectin, respectively by mechanism of

hydrolysis.

1.4.3.1.1.1 Polymethylgalacturonase (PMG)

PMG catalyze the hydrolysis of pectins (polymethylgalacturonates) to release

oligomethylgalacturonates or monomethylgalacturonates, depending on whether their pattern

of hydrolyses are either random or terminal (Jayani et al., 2005). Endo-

polymethlgalacturonase (endo-PMG) with the enzyme code number EC 3.2.1.41 catalyzes

the random hydrolytic cleavage of pectin to release oligomethylgalacturonate as end product,

25

whereas exo-polymethylgalacturonase catalyzes the terminal hydrolysis of pectin to release

monomethylgalacturonate units as end products (Figure 11).

Pectinase is a heterogeneous enzyme; therefore, if pectin esterase is present, its deesterifying

action on the pectin may prevent the correct evaluation of PMG activity (Pilnik and Voragen,

1970). Hence, PMG is less extensively studied than polygalacturonases. The mode of action

of PMG is shown Figure 11.

Figure 11: Mode of action of PMG(Sathyanarayana and Panda, 2003).

1.4.3.1.1.2 Polygalacturonase (PG)

Polygalacturonases are pectinolytic enzymes that catalyze the hydrolytic cleavage of the

polygalacturonic acid chain (pectic acid) with the introduction of water molecule across the

oxygen bridge. They are the most extensively studied among the family of pectinases. PG

exhibitendo and exo activities. Endo-PG is involved in random hydrolysis of O-glycosyl

bonds in 1, 4-α-galactosyluronic linkages in homogalacturonans or polygalacturonic acid

chain to release oligogalacturonates as end products. On the other hand, galacturonan 1, 4-α-

galacturonidase or Exo-PG are enzymes that degrade polygalacturonan by hydrolysis of the

glycosidic bonds from the non-reducing ends yielding the corresponding 1,4-α-D-

galacturonide and galacturonic acid (Favela-Torres etal., 2006). The mode of action of

polygalacturonases (PG) is shown in Figure 12.

H

O O

OH

OH

OH

OH

COOCH3

O

O

O

COOCH3 Pectin PMG

H

H

H H

H

H H

H

O

OH

OH

OH

OH

O

Oligo-/monomethylgalacturonate (for Endo or Exoenzyme, respectively)

(Pectic Acid)

COOCH3

COOCH3

O

OH

OH

HO H

H

H

H

H

H

H H

26

Figure 12. Mode of action of PG(Sathyanarayana and Panda, 2003)

1.4.3.1.1.2.1 Mechanism of Action of Polygalacturonase

The hydrolytic cleavage of glycosidic linkages within pectic fragments is catalyzed

exclusively by the family 28 polygalacturonases, also calledthe 28 glycoside hydrolases

(GH28s) (Abbott and Borastan, 2008). In addition to homogalacturonan, (GH28s) are also

involved in hydrolysis of heterogenous pectin derivatives such as rhamnogalacturonan and

xylogalacturonan. Studies on the reaction mechanism of enterobacteriaceae GH28s revealed a

catalytic cluster of three aspartate residues: D202, D223 and D224 (Figure 15). These amino

acids are positioned within 5Å of one another and approach the substrate in a “syn”

conformation. Interestingly, D202 and D223 are conserved within the catalytic sites of all

known GH28s, including rhamno- and xylo-galacturonases (Markovic and Jancek, 2001). The

hydrolysis reaction proceeds by a single-step inverting mechanism resulting in

stereochemical inversion around the anomeric carbon of the leaving group (Figure 13). Based

upon proximity to the scissile glycosidic oxygen and mutagenic studies, D223 is considered

to be the general acid (Shimizu et al., 2002). Presently, it is not known which of the

complementry aspartates operates as the general base by accepting a hydrogen atom and

charging the nucleophilic water. Further experiments are required to detail the role of D203

and D224 along the reaction coordinate (Abbott and Borastan, 2008).

In addition site-directed mutagenesis studies on the active site topology of Asperlligus niger

endoploygalacturonase II revealed the importance of Asp-180, Asp-201 and Asp-202 in

polygalacturonase catalysis. Polygalacturonase has been shown to hydrolyze glycosidic

bonds with an inverting mechanism that requires two carboxylic groups at a distance of 9 -

9.5Å from each other (McCarter and Withers, 1994). Armand et al. (2000) proposed that

Asp-180, with the assistance of Asp-202, acts as a base to activate the bound water molecule

whereas, Asp-201 acts as the general acid that protonates the product when it departs.

According to these authors, three arguments are in favour of this proposal:

O O

OH

OH

OH

H

OH

COOH

O

O

O

COOH Pectic acid PG

H

H H

H

H H

H

O OH

OH

OH

OH O

COOH

COOH

O

OH

OH

HO H

H

H

H

H

H

H H

Oligo/monogalacturonate

(for Endo or Exoenzyme, respectivey)

(Pectic Acid)

27

I. The mutation of Asp-201 residue led to an inactive enzyme;

II. Its replacement revealed the smallest effect on the “Bond Cleavage Frequencies‟‟

(BCFs) on oligogalacturonates, which suggest that Asp-201 does not directly

interact with the substrate; and

III. His-223, which is also important for catalysis most likely shares proton with Asp-

201, allowing this later amino acid to be in the proper ionization state to protonate

the product. Hence, Asp-180 and Asp-201 in ploygalacturonase II (PG11) are

directly involved in catalysis (Pickersgill et al., 1998) whereas, His-223 plays an

indirect role in catalysis.

Figure 13: Generalized reaction mechanism for inverting family 28 GHs(Abbott and

Borastan, 2008).

1.4.3.1.1.2.2 Overall Structure of Polygalacturonase

Endopolygacturnoase II folds into right-handed parallel β-helical structure comprising 10

complete turns with overall dimensions of approximately 65Å X 35ÅX 35Å(van Santen et

al., 1999). The number of amino acid per turn varies from 22 to 39, averaging to 29 residues

per turn. This variation is caused by the diversity of lengths of the loops connecting the β-

strands. The average rise per turn is 4.8Å, a value typical for β-helix is formed by four

parallel β-sheets, named PB1, PB2a, PB2b and PB3 (van Santen et al., 1999). This naming of

the β-sheets in pectate lyase structure, the first right-handed parallel β-helical structure that

was solved (Yoder et al., 1993). The prominent structural difference between

endopolygalacturonase II and pectate lyase is that the tertiary structure of

28

endopolygalacturonase II comprised four β-sheets which is one more than in the lyase

(Abbott and Borastan, 2008). PB1, PB2b and PB3 are the endopolygalacturonase II

conterparts of PB1, PB2 and PB3, respectively, of pectate lyase.The open-ended

endopolygalacturnoase II active site has a well designed topography for the recognition of

polygalacturonate, an observation that is in agreement with its previously described mode of

activity (Schevchik et al., 1999). Attack of internal galacturonide residues is enabling the

freedom of the substrate to extend out into solvent at either end. The electrostatic potential of

the solvent-accessible surface within the active site of the enzyme reveals two loops with

basic patches composed primarily of lysine (Pickersgill et al., 1998). These residues are

suitable candidates for involvements in substrate recognition events, as the formation of salt

bridges has been reported to be critical for catalysis by endopectate lyase (Charnock et al.,

2002) and predicated by modeling of an octagalacturonate-polygalacturonase complex in

Aspergillus aculeatus (Cho et al., 2001).

Figure 14: Three dimensional structure of polygalacturonase is displayed in a “cartoon”

format with a transparent solvent-accessible surface (Abbott and Borastan, 2008).

1.4.3.1.1.2.3 Occurrence of Polygalacturonases in Organisms

Polygalacturonases are frequently found in yeasts, moulds and bacteria (Favela-Torres, etal

2006; Luh and Phaff, 1951). They are also found in higher plants and parasitic nematodes

(Sakai etal, 1993). The endo-PGs have been isolated in microogranisms such as

29

Rhizoctoniasolanikulin (Marcus etal., 1986), Aureobasidiumpullulans (Sakai, 1984),

Fusariummoniliforme (De Lorenzo etal., 1987), Rhizopusstoloniforme (Manachini etal.,

1987), Thermomyces lanuginosus (Kumar and Palanivelu, 1999) and Aspergillussp (Nagai

etal., 2000). Endo-polygalacturonases have been cloned genetically in a number of microbial

strains (Raymond etal., 1994; Centis etal., 1996). The exo-PG, have been reported in

Erwiniacorotovora, Agrobacteriumtumefaciens, Bacteroidesthettaitamicron, E. chrysathemi,

Alternariamali, Fusariumoxysporum and Ralstoloniasalanacearium (Jayani etal., 2005).

Exo-polygalacturonases are distinguished into two types: fungal exo-PG, which produces

monogalacturonic acid as the main end product; and the bacterial exo-PG, which produces

digalacturonic acid as the main end product (Sakai etal., 1993).

Figure15: The extracellular endopolygalacturonase. Showing Superimposed catalytic sites

of closely related endopolygalacturonase(green) and periplasmic

exopolygalacturonase(yellow) displayed in wall-eyed format. The residues from both

endopolygalacturonase (D202, D223, and D224) and periplasmic exopolygalacturonase

(D381, D402, and D403) are labeled. The digalacturonate product from the

exopolygalacturonase complex is shown in beige, and subsites −1 and −2 are labeled in

red(Abbott and Borastan, 2008).

1.4.3.1.1.3 Determination of Pectin Hydrolase Activity

The type of substrate (Pectic acid or pectin) used for hydrolase assay makes it easier for one

to differentiate between PG and PMG. However, the presence of pectin esterase in a reaction

medium may interfere with the evaluation of PMG activity (Pilnik and Voragen, 1970). The

activity of pectin esterase in the medium will give rise to a product (polygalacturonic acid or

30

pectic acid) which is a specific substrate for polygalacturonase (PG). Polygalacturonase

activity and also polymethylgalacturonase activity can be quantified, and therefore expressed

in different units, whether by the reduction of viscosity in the reaction mixture or by the

release of reducing groups during the enzymic reaction under established conditions (Favela-

Torres etal., 2006). The amount of reducing sugar can be readily measured by colorimetiric

methods like 3, 5-dinitrosalicylate reagent (Miller, 1959) and asenomolybdate-copper reagent

method (Somogyi, 1952). One unit of enzyme activity is defined as the enzyme that releases

1mole ml-1

min-1

galacturonic acid under standard assay conditions.

1.4.3.1.3 Biochemical and Physicochemical Properties of Hydrolases

Hydrolases (PMG and PG) isolated from different microbial sources differ markedly from

each other with respect to their physicochemical and biochemical properties, and their mode

of action. The optimum pH for polymethylgalacturonase isolated from Apergillusniger was

around 4.0 (Koller and Neukom, 1967). Also, the optimal pH and temperature for

polygalacturonase were found to be 5.0 and 45ºC, respectively in Rhizopusstolonifer

(Manachini et al., 1987).

Among polygalacturonases obtained from different microbial sources, most have the optimal

pH range of 3.5- 5.5 and optimal temperature range of 30-50 ºC (Jayani etal., 2005). Two

endo-PGs (PG I and PG II), isolated from Aspergillusniger have optimal pH ranges of 3.8-4.3

and 3.0-4.6, respectively (Singh and Rao, 2002). In addition, few alkaline polygalacturonases

have been reported in Bacilluslineniformis(Singh etal., 1999) andFusariumoxysporumwith

optimum pH of 11.0(Pietro and Roncero, 1996). Barnby etal. (1990) identified four

isozymes, viz PGI, PGII, PGIII and PGIV with same molecular weight but differing in their

isoelectric points in Kluyveromycesmarxianus. Table6summarizes the biochemical and

physicochemical properties of polygalacturonases obtained from different sources.

31

Table 6: Biochemical and Physicochemical Properties of some Polygalacturonases (PG)

(Jayani et al., 2005)

Source of PG Nature Molecular

weight

(KDa)

pI Specific

activity

(µ/mg)

Km

(mg/ml)

Optimum

Temperature

Optimum

pH

Temperature

Stability

pH

Stability

Mucorflavus Endo 40 8.3 - - 45 3.5-5.5 40 2.5-6.0

Aspergillus

niger

Endo 61(PGI) - 982 0.12 43 3.8-4.3 50 -

Endo 38(PGII) - 3750 0.72 45 3.0-4.6 51 -

Thermococcus

auraniacus

Endo 35 5.9 5890 0.13 55 5.0 60 4.0-6.5

Aspergillusjap

anicus

Endo 38(PGI) 5.6 - - 30 4.0-55 - -

Endo 65(PGII) 5.3 - - 30 4.0-55 - -

Aspergillusaw

omori

Endo 41 6.1 487 - 40 5.0 50 4.0-6.0

BacillusSpKS

M-P410

Exo 45 5.8 54 1.3 60 7.0 50 7.0-12.0

Penicillium

frequentans

Exo 63 - 2571 1.6 50 5.0 - -

Exo 79 - 185 0.059 50 5.8 - -

Yersiniaentero

clitica

Exo 63 6.6 - - - - - -

Bacilluslichen

iformis

Exo 38 - 209 - 69 11.0 - 7.0-11.0

Saccharomyce

scerevisiae

- 43 - - - 45 4.5 - -

Fusariumoxys

porum

Exo 38 - 209 - 69 11.0 - 7.0-11.0

Kluyveromyce

s marxianus

Endo 496(PGI) 6.3 102.6 - - - - -

Endo 496(PGII) 6.0 102 - - - - -

Endo 496(PGIII) 6.3 107.8 - - - - -

Endo 496(PGIV) 5.7 97.6 - - - - -

32

1.4.3.1.2 Lyases

Lyases (or transeliminases) catalyze the non-hydrolytic breakdown of pectates or pectinates,

characterized by a trans-eliminative split of the pectic polymer (Sakai etal, 1993). These

enzymes break the glycosidic linkages at C-4 and simultaneously eliminate H from C-5

producing a 4:Δunsaturated product (Albersheim etal, 1960). The reaction mode of lyases is

shown in Figure 16.

Figure 16: Mode of action of lyases.

R = H for PGL and CH3 for PL where PGL = Polygalacutronate (Pectate) lyase and PL =

Polymethylgalacturonate (pectin) lyase (Sathyanarayana and Panda, 2003).

1.4.3.1.2.1 Classification of Lyases

Lyases are classified into two: Pectate lyase (PGL) and Pectin lyase (PL), depending on the

type of substrate they attack (pectate or Pectin). Pectate lyase breakdown pectic acid chains

by β-elimination, whereas pectin lyase breakdown pectin by β-elimination. Both enzymes

release unsaturated uronide along with units of oligo-/monogalacturonate as their end

product. Also these two enzymes (PGL and PL) can each be classified as endo and exo-

enzymes, depending on their pattern of attack on pectic structures (random or terminal,

respectively)

1.4.3.1.2.2 Mechanism of Action of Pectin Lyase

The two most common pectin lyase families (Families 1 and 9) operate by a common

mechanism to cleave glycosidic linkages between two neighboring galacturonic acid

monosaccharide. Generally, they utilize a two-step E1cb β-elimination, producing a planar

product with an unsaturated bond between C-4 and C-5 at the non-reducing end (Chanock et

al., 2002) (Figure 17). In the first step, the C-5 hydrogen is abstracted by a catalytic arginine

(Bronstead base). This process is coupled to H-5 resulting from Ca2+

coordination by the C-5

uronate group. Not surprising, due to the specialized chemistry of this catalytic base, the

O O

OH

OH

OH

OH

COOR

O

O

O

Pectic acid

or pectin PGL/PL

OH

COOR

O

OH

OH

OH

OH

O OH

OH

COOR

COOR

H H

H

H

H

H H

H H H

H

H H

H H

Unsaturated uronide

33

optimal pH of these enzymes is alkaline and ranges between 7.5 and 10 (Tardyetal., 1997).

Following H-5 abstraction, the transition state is stabilized by electron delocalization to the

C-5 carboxylate “sink”. In the second step of the reaction, product resolution results from

electron shuttling to O-4, triggering elimination of the leaving group (Abbott and Boraston,

2008). In a nutshell, the two main components of β-elimination are Brostead base and

divalent cation binding pocket.

Figure

17:Generalized reaction coordinates for calcium assisted β-elimination(Abbott and

Borastan, 2008).

1.4.3.1.2.3 Structure of Pectin Lyase

The structure of Erwinia chrysanthemi family I pectate lyase C was the first structure of a

pectinolytic enzyme ever described (Yoder, et al., 1993) (Figure 18). The enzyme adopts a

parallel β-helix topology with three distinct β-sheets formed from eight complete β-strand

turns. The center of the enzyme is stabilized by a ladder stacking residues, including a rich

hydrogen bond network between repeating asparagines, and hydrophobic stacks between

aliphatic and aromatic side chains. This architecture of intramolecular bonds generates a very

stable protein fold, presumably enabling persistence of the virulence factor within the harsh

extracellular environment during infection (Abbott and Boraston, 2008). When visualized

from the side, the enzyme is asymmetrical with a noticeable protrusion formed by several

loops (called the T3 loops) that are contributed from different β-strands. This region of the

molecule contains the catalytic center of the enzyme, which is a noticeable structural

heterogeneity compared to other β-helix enzymes (Abbott and Boraston, 2008). What is truly

remarkable about the β-helix is that following its initial discovery in 1993 (Yoder et al.,

1993), the topology has proven to be well-consumed scaffold for pectinases in general, as

other pectin lyases from sequence divergent families (pectate lyases 1, 3, and 9), and

enzymes harnessing distinct catalytic machinery (glycoside hydrolases 28, GH28s and

34

Carbohydrate esterases 8, CE8s) have been described (Jenkins and Pickersgill, 2001). The

superimposition of two family I pectate lyases (pectate lyase A and pectate lyase C) has a

calculated core root mean square deviation value of 1.72Å for 260 aligned Cα (Figure 19).

Beyond similarities in overall folds, closer analysis of the active site reveals that there is a

striking conservation of catalytic residue architecture. When the active site of pectate lyase C

is compared to other family I pectate structures (pectate lyases A and E) from Erwinia

chrysanthemi , there is a strigent conservation of catalytic amino acid. In pectate lyase A

structure, both catalytic base (R241) and calcium-coordinating residue (D184) are conserved

(Abbott and Boraston, 2008). The lack of calcium complexes for this enzyme precludes any

direct comparison of the metal coordination chemistries. Structural analysis of more distantly

related enzymes, however, does reveal subtle structural differences between them.

Superimposition of pectate lyase 1 and the family 9 pectate lyases has a root mean square

deviation of 2.31Å2

for 218 matched Cα . Analysis of the catalytic site architecture indicates

that in the family 9 enzyme the catalytic base, a lysine in this enzyme (K273), is shifted to

two position toward the reducing end of the sugar and the Ca2+

coordination site (D209, D237

and D233) is rotated around the substrate axis. Superimposition of the catalytic bases

revealed that the Ca2+

coordination pockets are in fact structurally conserved (Jenkins etal.,

2004). Hence, this observation is reflective of pectate lyases in general, as even diverse fold

families have similar active-site architecture (Abbott and Boraston, 2008).

Figure 18 :Three dimensional structure of pectate lyase is displayed in a “cartoon”

format with a transparent solvent-accessible surface (Abbott and Borastan, 2008).

35

Figure 19: The extracellular endo-pectin lyases: Showing thesuperimposed active sites of

pectin lyase C and pectin lyase Adisplayed wall-eyed format (Abbott and Borastan, 2008).

1.4.3.1.2.4 Occurrence of Lyases in Organisms

Activities of lyases (PGL and PL) have been reported in microorganisms and higher plants

(Whitaker, 1990). Though there are scanty reports on production of lyases in plants and

animals, most studies on lysaes have been from microorganisms (Whitaker, 1990). PGLs are

produced by a number of bacteria and some pathogenic fungi with endo-PGLs being more

abundant than exo-PGLs (Jayani etal., 2005).

Lyases are mainly produced by fungal genera Aspergillus, Penicillum and Fusarium but there

are reports on bacteria and yeasts (Yadav etal., 2009). Pectate lyase (PL) has been isolated

from Colletorichumlindemuthianum, Bacteroidesthataiotaomicron, Erwiniacarotovora,

Amucala Sp, Colletrichummaga, Erwiniachrysanthemi, C. gleosporioides (Jayani etal.,

2005). Also PLs have be reported in very few organisms such as Aspergillussojae (Ishii and

Yokotuka, 1972), Erwiniaaroideae (Kaminiya etal., 1974), Aspergillusniger (Kester and

Visser, 1994), Phythiumsplendens (Chen etal., 1998),Crystofilobasidiumcapitatum

(Nakagawa etal., 2005),Rizopousoryzae (Hamdy, 2005),Penicillumcanescens (Sinitsyna etal,

2007)and so on.

1.4.3.1.2.5 Determination of Lyase Activity

The best method for assaying lyase activity is by measuring the increase in absorbance at

235nm due to formation of 4:5 double bonds produced at the non-reducing ends of the

unsaturated products (Albersheim, 1966; Whitaker, 1990). The molar extinction coefficients

for PGL and PL are 4.6 x 103 and 5.5 x 10

3 M

-1CM

-1, respectively. One unit of enzyme

36

activity is defined as the amount of enzyme that releases 1µ mole of unsaturated product per

minute under assay conditions.

Reducing group methods are also useful in determining the lyase activity (Miller, 1959;

Collmer etal., 1988). Viscosity reduction method (Roboz etal., 1988), in conjunction with a

reducing group method or along with intermediate product analysis by high performance

liquid chromatography or gas chromatography, can be used to distinguish between endo and

exo splitting enzymes (Albersheim, 1966). Another method based on transformation of the

unsaturated uronic ester into a colored species possessing UV absorption at 550nm is also

used. Finally, for the detection of this unsaturated compound, thiobarbituric acid is claimed to

be the colorimetric test specific for the quantification of the pectin lyase activity (Albersheim

etal., 1960; Nedjma etal., 2001).

1.4.3.1.2.6 Biochemical and Physicochemical Properties of Lyase

In bacteria, lyases are the largest group of pectinolytic enzymes and are directly involved in

plant pathogenicity (Dixit etal., 2004). Generally, lyases have high optimum pH and are

activated by Ca2+

(Pilnik and Voragen, 1970). Pectate lyases (PGL) have absolute

requirement for calcium ions (Ca2+

) (Margo et al., 1994) and hence chelating agents such as

EDTA act as their inhibitors whereas pectin lyases (PL) do not have an absolute requirement

of Ca2+

but are stimulated by Ca2+

and other cations (Whitaker, 1990). Endo-pectin lyase is

the only enzyme known to be able to cleave, without the prior action of other enzymes, the α-

1,4-glycosidic bonds of highly esterified pectins (Alana etal., 1990).

Most lyases have molecular weights ranging between 30 and 40kDa, with isoelectric point

ranging from 7.0 to 11.0 (Jayani etal., 2005). They have pH optima in the alkaline range (7.5

-10.0) and temperature optima of 40-500C. Thermostable lyases have also been reported from

BacillusspTS47 and Thermoascusauratniacus (Martins etal., 2002). A thermostable exo-PGL

from Bacillussp showed maximum activity at pH 11.0 and Ca2+

for its activity (Singh etal.,

1999).

1.5 Sources of Pectinases

Pectinolytic enzymes are widely distributed in nature. They have been reported in plants,

bacteria, fungi, yeasts, insects, nematodes and protozoa (Zhong and Cen, 2005). Microbial

37

sources of pectinases became prominent in microorganisms such as bacteria, yeast and fungi

(Gummadi and Panda, 2003).

Microorganisms are currently the primary source of industrial enzymes: 50% originate from

filamentous fungi and yeast; 35% from bacteria, while the remaining 15% are either of plant

or animal origin (Bali, 2003). The filamentous fungi are most often used in the commercial

production of pectinases. Microbial pectinases have been extensively produced from several

fungi, including Aspergillussp (Angayarkanni etal., 2002), Aspergillusniger (Kumpoun and

Motomura, 2002), Penicillumexpansum (Cardoso etal., 2007), Penicillumroqueforti (Pericin

etal., 2007), Penicilliumchrysogenum (Banu etal., 2010), Rhizopusstolonifer (Manachini

etal., 1987), Aspergillusflavus (Mellon and Cotty, 2004) and pectinolytic moulds (Fawole and

Odunfa, 1992).

1.6 Production of Fungal Pectinases

Microorganisms are widely accepted as best producers of pectinases (Patil and Dayanand,

2006). They have a number of advantages: through the application of selection methods,

increase of biosynthesis via the conditions of substrates, wide spectrum of enzyme complex

and their application in genetic engineering via gene cloning (Kutateladze etal., 2009).

Bacteria are known to produce industrial enzymes, but filamentous fungi are desired for the

production of enzymes because their nature is generally regarded as safe (GRAS) (Sumantha

etal., 2005). The mycelial fungi are distinguished for such ability, as they are eukaryotic

organisms in comparison with prokaryotic organisms, have wide spectrum of genetic

information, and are able to perform microbial conversion (Kutaleladze etal., 2009).

In the course of time, several reports have been given on the optimization of fermentation and

microbiological parameters and different fermentation strategies for the production of

pectinases (Friedrich etal, 1989; Panda and Naidu, 1999; Panda and Naidu, 2000). With the

advent of molecular biology, vigorous research has been carried out on cloning and

expression of pectinase genes in various hosts such as Aspergilusflavus (Whitehead etal.,

1995). Saccharomycescerevisiae (Gognies etal., 1999), Erwiniachrysanthemi (Surgey etal.,

1996) and Colletotrichumgloesosporioides (Gognies etal., 2001). In addition, the expression

of cloned fungal genes including the pectinases genes Aspergillusaculeatus in yeast was

reviewed by Dalbogre (1997).

38

1.6.1 Substrate for the Production of Pectinase

Substrates that are employed in the production of enzyme should be solid as solid substrate

can give good encourage to the growing cells. Substrates should provide all needed nutrients

to the microorganisms for its growth. The synthesis of pectinase is induced or stimulated by

the presence of pure pectin, and for economic reasons, this is normally supplied by adding

pectin-rich agrowastes to the culture medium (Rombouts and Pilnik, 1980). Different

agrowastes,apple pomace (Hours etal., 1988), coffee pulp (Boccas etal., 1994), orange

bagasse and sugar cane bagasse (Martin etal., 2004), wheat bran and sugar cane bagasse

(Suresh and Viruthagiri, 2010) and mango peel have been utilized for the production of

pectinases. These agricultural wastes are most commonly used as substrate for solid state

fermentation process (Rangarajan etal., 2010). For instance, apple pomace has been reported

to be an attractive raw material for production of pectinases by Aspergillusfoetidus in solid

state cultures (Hours etal., 1988).

1.6.2 Methods used for Pectinase Production

There are two methods used commercially for pectinase production as well as other

enzymes(Murad and Foda, 1992). They include:

1. Solid state fermentation and

2. Submerged fermentation (Aquilar and Huitron 1990).

1.6.2.1 Solid State Fermentation (SSF) Technique

Solid state fermentation or SSF is generally defined as the cultivation of microorganisms on

solid materials under aerobic condition and in the absence or near-absence of free

waterbetween substrate particles (Sanzo etal., 2001). The metabolites obtained by SSF are

more concentrated and operational costs on downstream processing are minimized (Kumar

and Lonsane, 1987). Also, simple reactor designs with minimum controls and low moisture

content of the fermenting medium makes SSF system economical with less risks of bacterial

contamination (Singh etal., 1999). In spite the aforementioned qualities of SSF technique, the

technique still suffer difficulties in the control of pH, temperature and oxygen tension

compared to the SmF technique (Castilho etal., 2000; Coasta, 1998).

39

1.6.2.2 Submerged Fermentation Technique

Submerged fermentation (SmF) technique is the cultivation of microorganisms on liquid

broth. SmF system for enzyme production are generally conducted in stirred reactors under

aerobic conditions or fed batch systems (Bali, 2003). The fermentation system requires large

volumes of water, continuous agitation and generates lot of effluents. Also, the

physicochemical properties such as pH, temperature and oxygen tension are easier to control

in SmF than in solid state fermentation (Canel and Moo-Young, 1980; Costa etal., 1998;

Castilho etal., 2000).

However, high capital investment and energy costs, and the infrastructural requirements for

large-scale production make the application of SmF technique in enzyme production,

impractical in a majority of developing country environments (Bali, 2003). Inspite these

limitations, SmF have been used for pectinase production by several authors (Aguilar and

Huitron, 1990; Galiotou-Panayotou etal., 1993; Shivakumar and Krishnand, 1995; Blandino

etal., 2001;Patil and Dayanad, 2006). Table 7 shows the major differences between SSF and

SmF.

Approximately 90% of all industrial enzymes are produced in SmF, frequently using

specifically optimized, genetically manipulated microorganisms. In this respect SmF

processing offers an insurmountable advantage over SSF. On the other hand, almost all these

enzymes could also be produced in SSF using wild-type microorganisms (Filer, 2001; Pandey

et al., 2001). Interestingly, fungi, yeasts and bacteria that were tested in SSF in recent

decades exhibited different metabolic strategies under conditions of solid state and

submerged fermentation.The aim of SSF is to bring the cultivated fungi or bacteria into tight

contact with the insoluble substrate and thus to achieve the highest substrate concentrations

for fermentation. This technology results, only on a small scale, in several processing

advantages of significant potential economic and ecological importance as compared with

SmF (Table 7).

However, there are also several disadvantages of SSF, which have discouraged use of this

technique for industrial production. The main obstructions are due mainly to the build-up of

gradients of temperature, pH, moisture, substrate concentration or CO2 during cultivation,

which are difficult to control under limited water availability.

40

Table 7: Comparison of Solid State and Submerged Fermentation for Pectinase

Production.

Factor Submerged Fermentation Solid State Fermentation

Substrates

Aseptic conditions

Water

Metabolic heating

pH control

Oxygen tension

Soluble substrates (pectin or

Pectin-rich substrates)

Heat sterilization and aseptic

control

High volumes of water

consumed and effluents

discarded

Easy control of temperature

Easy pH control

Easy to control

Insoluble substrates (mainly

pectin-rich agrowastes)

Vapor treatment, non-sterile

conditions

Limited consumption of water;

limited effluent.

Low heat transfer capacity

Buffered solid substrates

Not easy to control

(Raimbault, 1998; Bali, 2003).

1.6.3 Factors Affecting Microbial Pectinases Production

Environmental and nutritional factors are known to have marked effects on enzyme

production by microorganisms. There are, therefore, variations in optimum conditions for

pectic enzyme production. Some of the cultural factors that affect the production of pectic

enzymes are presented in this study.

1.6.3.1 Initial pH of growth medium: According to Shoichi et al. (1985) the initial pH of

the medium has a great effect on the growth of the organism, on the membrane permeability,

also on the biosynthesis and stability of the enzymes (Murad, 1998; Murad and Salem, 2001).

Optimum production of pectic enzymes from many moulds has been reported to be within the

acidic pH range (Zetelaki-Horvath, 1980; Shin et al., 1983). Zheng and Shetty (1999),

reported that, polygalacturonase produced from Lentinusedodes has a relatively lower

optimum pH (pH 5.0) in addition, Piccoli-Valle et al. (2001) observed that a high

polygalacturonase and pectin esterase activity was showed by P. griseoroseum in more acid

pH of 4.5 and 5. Also, Silva et al. (2002) found that P. viridicatum showed maximum

production of polygalacturonase and pectinlyase at a pH of 4.5 and 5, respectively.

41

Fawole and Odunfa (2003) reported that the optimum pectolytic activity observed was at pH

5. Phutela et al. (2005) concluded that the thermophilic fungi A. fumigatus Fres expressed

maximum pectinase (1116 Ug-1) activity at pH 4.0 while polygalacturonase was active at pH

5.0 (1270 Ug-1). Also, Debing et al. (2005) found that the pH 6.5 was the optimal pH for

pectinase production fromAspergillus. niger by solid state fermentation. Reda et al. (2008)

found that the polygalacturonase productivity by Bacillus firmus-I-10104 reached its

maximum at initial pH 6.0 and 6.2. Rasheedha et al. (2010) found that P. chrysogenum

exhibited maximum polygalacturonase production at initial pH of 6.5. However, the

mechanism by which the pH acts on the production pectic enzyme is not known.

1.6.3.2 Incubationperiod: The time of fermentation had a profound effect on microbial

product formation (Murad and Foda, 1992; Murad, 1998; Murad and Salem, 2001).

Maximum production of pectic enzyme from different moulds varies from 1 to 6 days

(Ghildyal et al., 1981). Castilho et al. (2000) reported that the highest polygalacturonase

activities were obtained by Aspergillusnigerafter 70 h of fermentation period. In addition,

Fawole and Odunfa (2003) reported that optimum production of pectinmethylesterase was

obtained after 4 days of fermentation under submerged fermentation condition. Moreover,

Sarvamangala and Dayanand (2006) observed a gradual increase in the production of

pectinase from deseeded sunflower head by Aspergillusniger after 72 h of fermentation

period in submerged and up to 96 h in solid-state conditions. Reda et al. (2008) found that the

level of polygalacturonase increased gradually with increasing the incubation period up to a

maximum of 96 h by Bacillus firmus-I-10104 under solid state fermentation conditions.

1.6.3.3 Nitrogen Source: The effects of organic and inorganic nitrogen sources on the

production of pectinase were extensively studied. The observations of Hoursetal. (1988)

suggested that lower levels of (NH4)2SO4 (0.16%), or K2HPO4 (0.1%) added to the growth

medium as inorganic nitrogen sources did not influence pectinase yield. In addition Galiotou-

Panayotou and Kapantai (1993) observed that ammonium phosphate and ammonium sulphate

did influence production of pectinase positively but also recorded the inhibitory effects of

ammonium nitrate and potassium nitrate on pectinase production. Moreover, Sarvamangala

and Dayanand (2006) revealed that both ammonium phosphate and ammonium sulphate did

influence production of pectinase positively in both submerged and solid-state conditions. In

contrast, Sapunova (1990) found that ammonium salts stimulated the pectinolytic enzyme

production in AspergillusalliaceusBIM-83. Moreover, Sapunova et al. (1997) has also

42

observed that (NH4)2SO4 stimulated pectinase synthesis, as in its absence fungus displayed a

slight proteolytic activity and did not produce extracellular pectinases. In addition, Fawole

and Odunfa (2003) found that ammonium sulphate and ammonium nitrate were good

nitrogen sources for pectic enzyme production from Aspergillusnigerwhile glycine and

tryptophan did not support enzyme production. Also, Phutela et al. (2005) reported that

(NH4)2SO4 stimulated pectinase production, as in its absence fungus displayed a slight

proteolytic activity and did not produce extracellular pectinases. In addition, Rasheedha et al.

(2010) found that ammonium sulphate has enhanced the production of P. chrysogenum

pectinase.

On the other hand, report of Aguilar et al. (1991) showed yeast extract (organic nitrogen

source) as the best inducer of exopectinases by Aspergillus sp. Moreover Kashyap et al.

(2003) found that, yeast extract, peptone and ammonium chloride were found to enhance

pectinase production up to 24% and addition of glycine, urea and ammonium nitrate inhibited

pectinase production. Also, Reda et al. (2008) found that the maximum value of

polygalacturonase productivity by Bacillus firmus-I-10104 reached up to 350 U mL-1 in the

presence of peptone as a nitrogen source in the growth medium. In addition, Vivek et al.

(2010) found that organic nitrogen sources showed higher endo, exo pectinases activities than

inorganic nitrogen sources. Also the increasing trend in the enzymes activity with the

increase in nitrogen source content was observed in the case of organic nitrogen sources

while decreasing trend observed for inorganic nitrogen sources

1.6.3.4 Carbon Source: An adequate supply of carbon as energy source is critical for

optimum growth affecting the growth of organism and its metabolism. Aguilar and Huitron

(1987) reported that the production of pectic enzymes from many moulds is known to be

enhanced by the presence of pectic substrates in the medium. Fawole and Odunfa (2003)

found that pectin and polygalacturonic acid promoted the production of pectic enzyme and

they observed the lack of pectolytic activity in cultures with glucose as sole carbon source

reflects the inducible nature of pectic enzyme from the strain of Aspergillusniger. However,

when different concentrations of glucose were added to the medium containing pectin,

production of pectic enzymes was inhibited at high glucose concentration while low glucose

concentrations (0.5% w/v) stimulated enzyme production. Also, the reducing sugar content

of the culture filtrate increased with increase in the amount of glucose added to the growth

medium. The ability of high concentrations of glucose in the medium to meet growth

43

requirement of the organism probably made the breakdown of pectin in the medium

unnecessary or minimal and thus the low pectic activities observed in cultures. Phutela et al.

(2005) stated that wheat bran supported maximum pectinase production (589 U g-1) while

pure pectin give the maximum production of polygalacturonase (642 U g-1). Sarvamangala

and Dayanand (2006) reported that glucose (4-6%) increase the production of pectinase in

submerged condition whereas 6-8% sucrose gives better yield of pectinase in solid-state

condition. Reda et al. (2008) reported that Solanumtuberosum (ST) peels was the best carbon

source for polygalacturonase production by Bacillus firmus-I-10104 under solid state

condition.

1.6.4 Purification of Microbial Pectinases

In order to characterize and study the properties of microbial pectinases the enzymes must be

purified. Important purification methods for the isolation of different pectinases are briefly

summarized in this section. Pectinases from various sources of microorganisms have been

purified to homogeneity. An exo-PG has been separated from mycelial extracts of

Aspergillusniger by eluting from DEAE cellulose with 0.2M sodium acetate buffer at pH 4.6.

Purification was efficient with 209-fold increase in specific activity with a recovery of 8.6%

and the enzyme displayed its full activity only in the presence of Hg2-

ions (Mill, 1966). A

second PG was isolated with 205-fold increase in specific activity with a recovery of 1%.

These two PGs are differentiated by their optimum pH and PG II was not inhibited by

chelating agents and did not require Hg2-

for activity (Mill, 1966).

Benkova and Slezarik (1966) developed a purification strategy for the isolation of

extracellular PMG, PG and PE. The enzyme was salted out with ammonium sulphate and

precipitated with ethanol after gel filtration through Sephadex G-25. Repeated

chromatography on DEAE-cellulose column yielded a homogeneous preparation of enzyme.

Exo-PG, Endo-PG and pectinesterase have been separated from the culture filtrate

ofTrichoderma reesei by Sephadex chromatography (Markovic et al., 1985).

Polygalacturonase from Rhizopus stolonifer has been purified up to 10-fold by ethanol

precipitation followed by CM-Sepharose 6B ion exchange chromatography and gel filtration

by Sephadex G-100 (Manachini et al., 1987). PG and PL (pectinlyase) from Aureobasidium

pullulans LV10 have been separated by CM-Sepharose 6B followed by column

chromatography (DEAE-cellulose column) and gel filtration on Sephadex G-100 (Manachini

44

et al., 1988). PG and PL (pectinlyase) have been separated into PG I and PG II and PL I and

PL II, respectively.

Pectatelyase (PGL) was synthesized by Amycolata species and the extracellular crude

enzyme has been purified to homogeneity by both cation and anion exchange columns and

hydrophobic interaction chromatography (Bruhlmann, 1995). It has been observed that

purification resulted in a 4-fold increase in specific activity with 37% recovery. Pectinases

from Clostridium aectobutylicum ID 91-36 a UV mutant, has been purified by cation

exchange chromatography on a Sepharose column by eluting with NaCl (Seethaler and

Hartmeier, 1992). Endopectate lyase synthesized by Bacillus macerans has been purified by

ammonium sulphate precipitation followed by DEAE-Sephadex A-50 chromatography and

CM-cellulofine chromatography (Miyazaki, 1991). Similarly endopectate lyase I/IV has been

isolated from the culture filtrate of Erwiniacarotovora by CM-Sepharose CL 6B

chromatography, Sephadex S-200 gel filtration and isoelectric focusing (Tanabe et al., 1984).

Kobayashi et al. (2001) purified the first bacterial; exo-PG from Bacillus sp. strain KSM-

P443 to homogeneity. This enzyme releases exclusively mono-galacturonic acid from

polygalacturonic acid (PGA), di-, tri-, tetra-and penta-galacturonic acids. They also

determined the N-terminal sequence and concluded that no sequence matched with other

pectinases reported to-date. An extracelluar endo-PG produced by Aspergillus awamori IFO

4033 was purified homogeneity using cation-exchange and size-exclusion chromatographic

columns (Nagai et al., 2000). Sakamoto et al. (1994) isolated protopectinase-N (PPN) and

protopectinase-R (PPR) from the culture filtrate of Bacillus subtilis IFO3134. These enzymes

have been purified by hydrophobic interaction chromatography on butyl-toyopearl 650 M,

cation exchange chromatography on CMtoyopearl650 M and gel filtration on sepharose

12HR. These enzymes have been found to be stable over a wide range of pH and temperature.

Endopectate lyase produced by Erwinia caratovara FERM P-7576 has been selectively co-

sedimented with an extracellularly produced lipopolysaccharide lipid complex (Fukoka et al.,

1990). The cell free broth was precipitated and the enzyme separated by gel chromatography

with a specific activity of 710 U mg-1 of protein. Co-sedimentation has been affected by pH

and ionic strength. Denis et al. (1990) studied the effect of shear stress on purification of five

isozymes of pectate lyase produced by Erwinia chrysanthemi 3937 in ultrafiltration

equipment. Activity was not affected during 7 h of pumping and 36% activity was lost after

25 000 passes. New affinity matrices have been developed for the purification of pectinases,

which possess better mechanical and chemical stability than those cross-linked one with

45

pectic acid (Lobarzewski et al. 1985). The culture filtrate was desalted on a Sephadex G-25

column. The supports used were silanized controlled pore glass, silica gel silanized with 5-

aminopropyl triethoxysilane. All supports were activated with 3-(3-dimethylaminopropyl)

carbodiimide and best results were obtained with silanized controlled pore glass. Gupta et al.

(1996) developed an affinity precipitation technique for separation of selective proteins using

heterobifunctional ligands. They used a soluble form of the ligand for affinity binding and

then precipitation was induced for separating the protein complex. Alginate was used as

successful ligand for pectinases.

1.7 Biotechnological applications of microbial pectinases

1.7.1 Fruit juice extraction

The largest industrial application of pectinases is in fruit juice extraction and clarification

(Figure 20). Pectins contribute to fruit juice viscosity and turbidity. A mixture of pectinases

and amylases is used to clarify fruit juices. It decreases filtration time up to 50%(Blanco et

al., 1999). Treatment of fruit pulps with pectinases also showed an increase in fruit juice

volume from banana, grapes and apples (Kaur et al., 2004). Pectinases in combination with

other enzymes, viz., cellulases, arabinases and xylanases, have been used to increase the

pressing efficiency of the fruits for juice extraction (Gailing and Guibert, 2000). Vacuum

infusion of pectinases has a commercial application to soften the peel of citrus fruits for

removal. This technique may expand in future to replace hand cutting for the production of

canned segments (Baker and Wicker, 1996). Infusion of free stone peaches with

pectinmethylesterase and calcium results in four times firmer fruits. This may be applied to

pickle processing where excessive softening may occur during fermentation and storage

(Baker and Wicker, 1996).

1.7.2 Liquefactionof pulp

Instead of pressing, pulps are better liquefied enzymatically using pectinolytic enzymes.

Pectinases in combination with other enzymes like, cellulases, arabinases and xylanases, have

been used to increase the pressing efficiency of fruits for juice extraction (Gailing etal.,

2000). Enzymatic hydrolysis of the fruit cell wall increases the extraction yield, reducing

sugars, soluble dry matter content and galacturonic acid content and titrable acidity of

products (Joshi etal., 1991; Drilleau, 1994).

46

1.7.3 Textile processing and bioscouring of cotton fibers

Pectinases have been used in conjunction with amylases, lipases, cellulases and

hemicellulases to remove sizing agents from cotton in a safe and ecofriendly manner,

replacing toxic caustic soda used for the purpose earlier (Hoondal etal., 2000). Bioscouring is

a novel process for removal of non-cellulosic impurities from the fiber with specific enzymes.

Pectinases have been used for this purpose without any negative side effect on cellulose

degradation (Hoondal etal., 2000).

Figure 20: Pectinases at different phases of fruit juice manufacturing

47

Pectinases at different phases of fruit juice manufacturing

1.7.4 Degumming of plant bast fibres

Bast fibres are the soft fibres formed in groups outside the xylem, phloem or pericycle, e.g.

Ramie and sunn hemp. The fibers contain gum, which must be removed before its use for

textile making (Hoondal etal., 2000). The chemical degumming treatment is polluting, toxic

and non-biodegradable. Biotechnological degumming using pectinases in combination with

xylanases presents an ecofriendly and economic alternative to the above problem (Kapoor et

al., 2001).

1.7.5 Retting of Plant Fibres

Pectinases have been used in retting of flax to separate the fibres and eliminate pectins

(Hoondal etal., 2000). In recent years, a few fundamental studies have been initiated on the

enzymatic retting process. These employ purified enzymes on defined substrates and

characterization of the resulting products. Apectinase from Rhizomucorpumilis was used for

flax retting (Henriksson et al., 1999). To ensure maximum strength of the thread

manufactured from retted flax, only a small fraction of the pectinases belonging to the fibre

bundles needs to be hydrolyzed. In developing nation and particularly in countries where

forest lands are endangered from over exploitation, better use might be made of herbaceous

fibres for paper production. Such feedbacks should be amenable to enzymatic pulping and the

resulting processes should give together yields with fewer environmental problems.

1.7.6 Waste water treatment

Pectinolytic enzymes are applied in the treatment of pectin containing waste water. For

instance, waste waters from fruit juice and vegetable food industries contain pectinaceous

materials as by-products. Pretreatment of these waste waters with pectinases facilitate

removal of pectins and renders it suitable for decomposition by activated sludge treatment

(Hoondal etal., 2000).

1.7.7 Coffee and tea fermentation

Pectinases have been used to eliminate pectin in coffee and tea processing plants (Boccas

etal., 1994; Rashmi etal., 2008). These enzymes facilitate tea fermentation and also destroy

the foam forming property of instant tea powders by destroying pectins (Jayani etal., 2005).

They are also used in coffee fermentation to remove mucilaginous coat from coffee bean.

48

1.7.8 Paper and pulp industry

During paper making, pectinases can deploymerize pectins and subsequently lower the

cationic demand of pectin solutions and the filtrate from peroxide bleaching (Reid and

Richard, 2004).

1.7.9 Animal feed

Pectinases are used in the enzyme cocktail, used for the production of animal feeds. This

reduces the feed viscosity, which increases absorption of nutrients, liberates nutrients, either

by hydrolysis of non-biodegradable fibers or by liberating nutrients blocked by these fibers,

and reduces the amount of faeces (Hoondal etal., 2000).

1.7.10 Purification of plant viruses

A virus prior to purification is very limited. Very pure preparations of viruses are required in

order to carry out chemical, physical, and other biological studies. The need numerous

purification that can be adapted to many of the virus that infect plants. However, there are

several different purification systems that can be selected for use according to the type of

virus. In those cases in which the virus is restricted to phloem, certain enzymes, such as

alkaline pectinases and cellulases can be used to liberate the virus from the tissues (Salazar

and Jayasinghe, 1999)

1.7.10 Oil extraction

Pectinases have been used in the extraction of vegetable oils (Rashmi etal., 2008) and citrus

oils such as lemon oil (Jayani etal., 2005). The enzyme destroys the emulsifying properties of

pectin, which interfers with the collection of oils from citrus peel extracts (Scott, 1978).

1.7.11 Improvement of chromaticity and stability of red wines

Pectinolytic enzymes added to macerated fruits before the addition of wine yeast in the

process of producing red wine resulted in improved visual characteristics (colour

andturbidity) as compared to the untreated wines. Enzymatically treated red wines presented

chromatic characteristics, which are considered better than the control wines. These wines

also showed greater stability as compared to the control (Revilla and Ganzalez-san, 2003).

49

1.8 Aim and Objectives of Study

Works on pectinase production by SmF system using pectin extracted from orange peels as

carbon source are scanty. Hence, this work reports on the production, partial purification and

characterization of pectinases isolated from Aspergillus nigerunder submerged fermentation

condition using pectin extracted from orange peels as carbon source.

This work is therefore designed to achieve the following objectives:

1. Extraction of pectin from ground orange peels.

2. Isolation of pectinase producing fungi from natural source.

3. Production of extracellular pectinase using the isolated fungal population.

4. Partial purification of the pectinases through ammonium sulphate precipitation and

dialysis.

5. Characterization of the pectinases produced with respect to pH, temperature and

substrate concentration.

50

CHAPTER TWO

2.0 MATERIALS AND METHODS

2.1 Materials

2.1.1 Reagents

The chemicals used in the study were sourced as follows:

3, 5-dinitrosalicylic acid (DNS) - Sigma Chemical company (USA)

Bovine serum albumin (BSA) - Bio Rad Laboratories (India)

D-(+)-Galacturonic acid monohydrate - Sigma-Aldrich (USA),

Folin-Ciocalteau - Sigma-Aldrich (USA).

All other chemicals used in this work were of analytical grade and were products of Merck

(Germany), BDH chemical limited (England), May and Baker limited (England), Riedel-

DeHaen Hannaves (Germany), Hopkins and Williams Essex (England), Fluka chemical

company (Germany), Kermel chemicals (China) and Lab.Tech Chemicals, Avighkar (India),

unless otherwise stated.

2.1.2 Apparatus/ Equipments

Autoclave: UDAY BURDON‟s Patent Autoclave, made in India.

Centrifuge: Finland Nigeria 80-2B.

Glass wares: Pyrex

Incubator: B and T Trimline incubator.

Magnetic stirrer: AM-3250B Surgi Friend Medicals, England.

Microscope: WESO microscope.

Milling machine: Thomas Willey laboratory Mill Model 4, Anthor H (Thomas.

Company, Philadelphia, USA)

Oven: Gallenkamp Hotbox, England.

pH meter: Ecosan pH meter, Singapore.

Sensitive weighing balance: B2404-5 mettler Toledo, Switzerland.

Water bath: Model DK.

Weighing balance: Ohaus Dial-O-Gram, Ohaus Cooperation, N. J. USA.

51

Uv/visible spectrophotometer: Jenway 6405

2.1.2 Collection of Orange Fruits

Fresh orange fruits(Citrussinensis) were obtained from orange sellers at Ogige market,

Nsukka, Enugu State. It was ensured they came from the same source so as to maintain

experimental homogeneity.

2.1.3 Collection of Microorganism

Three Aspergillus species were isolated from soil containing decaying fruits and

vegetablesusing the method described by Martin et al.(2004). The soil samples were collected

in clean dry plastic containers and transported to the laboratory.

2.2 Methods

2.2.1 Processing of the Orange Peels

The fresh orange fruits were washed with water to reduce microbial load. The fruits were

peeled, cut into small bits and treated with 96% ethanol to disinfect the peels. The ethanol

treated peels were washed again with water and sun dried for seven days. The dried peels

were ground to powder with a milling machine.

2.2.2 Extraction of Pectin

Pectin was extracted using the method described by McCready (1970). 100g of ground

orange peels were weighed into a 2000ml beaker containing 800ml of distilled water. 12g of

freshly ground sodium hexametaphosphate was added and the initial pH was adjusted with

3N HCL to 2.2 ± 0.1. The mixture was heated in a water bath at 70oC for 1 hour and stirred

with a stirrer and the pH checked at intervals of 15mins. The water lost was replaced at

intervals except in the last 20mins of the extraction. The extract was vacuum filtered through

a muslin cloth and the residue was washed with 200ml of distilled water, and the washings

were added to the filtrate. The filtrate was concentrated by evaporation on a hot plate to

approximately 1/5 of the initial volume.

The concentrated pectin was cooled to 50oC and poured into a volume of ethanol in the ratio

of 1:3 the ethanol contained 0.5M HCL. The mixture was stirred for 30mins and allowed to

stand for 1 hour. The mixture was vacuum filtered using vacuum filter and washed with 20ml

ethanol-HCL solution. The extract was finally washed with acetone to remove traces of HCL

52

and ethanol. The extract was dried in an oven at 40 oC for few hours to constant weight and

ground finely.

Percentage yield of pectin was calculated by the following formula:

%Yield of pectin = Amount of pectin obtained

Total amount of orange peel powder used × 100

2.2.3 Isolation of Pectinolytic Fungi

2.2.3.1 Preparation of Liquid Broth

Samples (2g) from agricultural soil and decaying orange fruits were pooled and homogenized

in sterile medium containing 1% orange pectin; 0.14% of (NH4)2 SO4, 0.2% of K2HPO4,

0.02% of MgSO4.7H2O, 0.1% of nutrient solution containing; 5g/L FeSO4.7 H2O, 1.6mg/L

MnSO4.H2O, 1.4mg/L ZnSO4.7H2O, 2.0mg/L CoCl2. The mixture was incubated at 30oC for

24 hours.

2.2.3.2 Preparation of Solid Medium

The medium contained 1% orange pectin, 0.14% of (NH4)2SO4, 0.2% of K2HPO4, 0.02% of

MgSO4.7H2O, 0.1% of nutrient solution containing; 5mg/L FeSO4.7 H2O, 1.6mg/L

MnSO4.H2O, 1.4mg/L ZnSO4.7H2O, 2.0mg/L CoCl2 and 3% agar-agar (the gelling agent)

(w/v). The medium was autoclaved at 121oC for 15min. It was allowed to cool to about 45

oC

and then poured into Petri dishes and allowed to gel. The plates were then incubated in a B &

T Trimline incubator at 37oC overnight to check for sterility.

2.2.3.3 Inoculation of Plates and Sub-culturing

A loop of homogenized extract from the liquid broth was streaked onto the solid medium

under the flame of bunsen burner. Streaks were made from each side of the plate, marking an

initial point, with sterilization of the wire loop after each side has been completed. The plates

were thereafter incubated at 35ºC till visible colonies were observed. All morphological

contrasting colonies were purified by repeated streaking and sub-culturing on separate plates.

This process was continued till pure fungal cultures were obtained.

2.2.3.4 Storage of Pure Fungal Isolates

Pure fungal isolates were maintained on Potato Dextrose Agar (PDA) slopes or slants as

stock cultures. PDA media were prepared according to the manufacture‟s description.

53

2.2.3.5 Macroscopic Features of the Isolated Fungi

Three days old pure cultures were examined. The color, texture, nature of mycelia or spores

and growth patterns were also observed. Photographs of the cultures were also taken.

2.2.3.6 Fungal identification

The three days old pure culture was used in preparing microscopic slides. A little bit of the

mycelia was dropped on the slide and a drop of lactophenol blue was added to it. A cover slip

was placed over it and examination was performed under the light microscope at X400

magnification. Identification was carried out by relating features and the micrographs to

“Atlas of mycology” by Barnett and Hunter (1972).

2.2.4 Fermentation Experiments

2.2.4.1 The Fermentation Broth

Submerged fermentation (SmF) technique was employed using a 250ml Erlenmeyer flask

containing 100ml of sterile cultivation mediumoptimized for pectinase with 0.1% NH4NO3,

0.1% NH4H2PO4, 0.1% MgS04.7H2Oand 1% orange pectin. The flask was covered with

aluminum foil and autoclaved at 121oC for 15mins.

2.2.4.2 Inoculation of the Broth

From the PDA slants, fresh plates were prepared as described in section 2.2.3.2 and

inoculated. Three days old cultures were used to inoculate the flasks. In every sterile flask,

two discs of the respective fungal isolates were added using a cork borer of diameter 10mm

and then plugged properly. The culture was incubated for 7 days at room temperature (30oC).

2.2.4.3 Harvesting of the Fermented Broth

At each day of harvest, flasks were selected from the respective groups and mycelia biomass

separated by filtration. Each day, the fiterate was analyzed for pectinase activity and

extacellular protein concentration till the 7th day of fermentation.

2.2.4.4 Mass Production of Enzyme

54

After the 7 days pilot studies under SmF, the day of peak pectinase activity was chosen for

mass production of enzyme from the respective fungal isolates. Several 250ml Erlenmeyer

flasks were used to produce upto 2.5litres of the enzyme using the method described in

sections 2.2.4.1 and 2.2.4.2. Harvesting was carried out on the respective peak days of

enzyme activity.

All the experiments for isolation, screening and pectinolytic enzyme production were done

under sterile conditions and adequate safety measures were undertaken (Appendix two)

2.2.5 Pectinase Assay

Pectinase activity was evaluated by assaying for polygalacturonase (Pg) activity of the

enzyme. This was achieved by measuring the release of reducing groups from orange pectin

using a modification of the 3,5 dinitrosalicylic acid (DNS) reagent assay method described by

Miller (1959) as contained in Wang et al.(1997) with little modifications.

The reaction mixture containing 0.5ml of 0.5% orange pectin in 0.05M sodium acetate buffer

pH 5.0 and 0.5ml of enzyme solution was incubated for 1 hour. 1ml of DNS reagent was

added and the reaction was stopped by boiling the mixture in a boiling water bath for 10mins.

The mixture volume was made up to 4ml with 1ml of Rochelle salt solution and 1ml of

distilled water. The reaction mixture was allowed to cool and then absorbance read at 575nm.

One unit of enzyme activity was defined as the amount of enzyme that catalyzes the release

of one micromole of galacturonic acid per minute.

2.2.6 Protein Determination

Protein content of the enzyme was determined by the method of Lowry et al. (1951), using

bovine serum albumin as standard as outlined in sections 1.6 and 1.7 (Appendix One).

2.2.6.1 Procedure for Protein Determination

For protein standard curve, the reaction mixture contained 0.0-1.0ml of protein stock solution

(2mg/ml BSA) in test tubes arranged in triplicates. The volume was made up to 1ml with

distilled water. For the test, 0.1ml of enzyme was mixed with 0.9ml of distilled water. In

either case, 5ml of solution E was added and allowed to stand at room temperature for about

10min. Then 0.5ml of solution C (dilute Folin-Ciocalteau phenol reagent) was added with

rapid mixing. After standing for 30min, absorbance was read at 750nm using

55

spectrophotometer. Absorbance values were converted to protein concentration by

extrapolation from the standard curve (Appendix four).

2.7 Partial Purification of Protein

2.2.7.1 Ammonium Sulphate Precipitation Profile

Ammonium sulphate precipitation profile was carried out to determine the concentration of

ammonium sulphate suitable for pectinase production. This was done at different ammonium

sulphate saturation ranging from 20 – 100% sulphate at intervals of 10% in each test tube

containing 10ml of crude enzyme. These were allowed to stand at cold temperature of about

4oC for 30hours. The test tubes were centrifuged at 3500 rpm for 30mins and pellets re-

dissolved in equal volumes of 0.05M acetate buffer pH 5.0. Pectinase activity of the

precipitates was assayed to determine the percentage ammonium sulphate saturation that has

the highest activity.

2.2.7.2 Ammonium Sulphate Precipitation of Pectinase

From the studies in section 2.2.7.1, maximum enzyme activity was achieved at eighty percent

(80%) ammonium sulphate saturation and was therefore used to precipitate one (1) litre of

crude enzyme. This was done by adding 516g of ammonium sulphate in 1000ml of crude

enzyme and stirring gently till the salt dissolves completely. The precipitate were re-

dissolved in 10ml of 0.05M acetate buffer pH 5.0 after centrifugation and then kept under

cold condition for further studies.

2.2.7.3 Dialysis

The precipitate was desalted by dialysis following the standard protocol; the 10cm pretreated

dialysis bag was used and activated by rinsing in distilled water. One end of the dialysis bag

was tightly tied and the precipitate recovered was introduced inside the bag. The other end of

the dialysis bag was tightly tied to prevent the leakage. After that, dialysis bag was suspended

in a beaker containing 0.05M sodium acetate buffer pH 5.0. Dialysis was carried out for

12hours with continuous stirring and buffer changed every 6 hours with a view to removing

low molecular weight substances and other ions that may interfere with enzyme activity

(Dixon and Webb, 1964). The dialyzed enzyme was also assayed for pectinase activity

(Miller, 1959) and protein concentration (Lowry etal., 1951) while the remaining sample was

stored frozen at -24oC.

56

2.2.8 Studies on Partially Purified Enzyme

2.2.8.1 Effect of pH Change on Pectinase Activity

The optimum pH for enzyme activity was determined using 0.05M sodium acetate buffer pH

3.5 - 5.5, phosphate buffer pH 6.0 - 7.5 and Tris-HCl buffer pH 8.0 - 9.5 at intervals of 0.5.

0.1% orange pectin solution was prepared by dissolving 0.1g pectin in 100ml of 0.05M of the

respective buffers. Also partially purified enzymes were dispersed in the various buffers and

0.5ml of the enzyme mixed with 0.5ml pectin solution at the corresponding pHs for pectinase

assays using the method described in section 2.2.5.

2.2.8.2 Effect of Temperature Change on Pectinase Activity

The optimum temperature was determined by incubating the enzyme with pectin solution at

25-70oC interval of 5

oC for 1hour and at the pH 5.0. The activity was then assayed using the

method described in section 2.2.5.

2.2.8.3 Effect of Substrate Concentration on Pectinase Activity.

The effect of substrate concentration on the activity of pectinase was determined by

incubating the enzyme with 10, 20, 30, 40, 50, 60, 70, 80, 90 and 100mg/ml of orange pectin

at pH 5.0 and temperature of 40oC.The Vmax and Km values of the enzyme were determined

using the double reciprocal plot.

2.2.8.4 Further Studies with Partially Purified Enzyme

Protein concentration and pectinase activity of partially purified enzymes were determined

using the methods described by Lowry et al. (1951) and Miller (1959) as shown in sections

2.2.6.1 and 2.2.5 respectively.

57

CHAPTER THREE

3.0 RESULTS

3.1 Orange Pectin Extraction

3.1.1 Orange Pectin Extraction Yield

Pectin extraction yield was found to be 15.5% at pH 2.2, temperature of 70oC and extraction

time of 1hour.

3.1.2 Photograph of Orange Pectin Extract

Figure 21shows the photograph of pectin extracted from ground orange peels

Figure 21: Pectin Extracted From Ground Orange Peels

3.2 Microorganisms

3.2.1 Selectionof Pectinolytic Fungi

Three fungal isolates were obtained from soil containing decaying fruits and vegetables. The

criterion for the selection process was based on isolation of species with similar

morphological features in both a test culture containing orange pectin and a standard culture

containing apple pectin as carbon sources, respectively.These isolates were qualitatively

screened for pectinolytic activity on selective media.Spectrophotometric assay of exudates

secreted by the fungal isolates indicated that they were pectinolytic.

58

3.2.2 Macroscopic and Microscopic Examination of Fungal Isolates

Genus identification was by examining both macroscopic and microscopic features of a three

day old pure culture. The colour, texture, nature of mycelia and/or spores produced, growth

pattern in addition to microscopic features such as separation, spore shapes and so on were

examined. Based on these characteristics,Aspergillus niger, Aspergillus fumigatus and

Aspergillus flavus were identified and confirmed as the three pectinolytic fungal isolates.

Among these three isolates, Aspergillus niger showed relatively higher pectinase activity and

was therefore selected for further studies. Figures 22, 23 and 24shows the photograph of the

pure culture of Aspergillus niger, Aspergillus fumigatus and Aspergillus flavusrespectively.

Figure 22: Pure culture of AspergillusNiger

59

Figure 23: Pure culture of Apergillusfumigatus

Figure 24: Pure culture of Aspergillusflavus

60

3.3 Production of Pectinases by Submerged Fermentation(SmF)

After seven days pilot study, the day of maximum pectinase production and maximum

protein production from the selected fungal specie (Aspergillusniger)was found to be on day

4 and 5 respectively. Day 4 which depicts the day of maximum enzyme secretion was

therefore used for massproduction.

Table 8: Pectinase Production from Aspergillusspecies

Source of Enzyme Day of Maximum

Pectinase Production

Day of Maximum Protein

Production

Aspergillus niger 4 5

Aspergillus fumigatus 4 5

Aspergillus flavus 5 5

3.4 Mass Production

2.5 litres of crude enzyme was harvested after 4days of submerged fermentation with

Aspergillusniger. This was stored at -24oC prior to its use.

3.5 Studies on Crude Enzyme

3.5.1 Protein Concentration of Crude Enzyme

Protein concentration of the crude enzymes produced by Aspergillus niger was found to be

0.486mg/ml (Table 8)

3.5.2 Pectinase Activity of Crude Enzyme

Pectinase (polygalacturonase) activity of the crude enzyme produced by Aspergillus nigerwas

found to be 25.73U/ml (Table 8).

3.6. Ammonium Sulphate Precipitation Profiles of Pectinases

80% ammonium sulphate saturation was observed to have the highest polygalacturonase

activity of 78.55U/ml.Hence,80% ammonium sulphate saturation was chosen for

precipitation of pectinases from Aspergillus niger. (Figure 25)

61

3.7 Studies on Partially Purified Enzymes

3.7.1 Purification Fold of Partially Purified Enzymes

Pectinases isolated from Aspergillus niger was partially purified approximately 2-fold with

specific activity of 92.08U/mg protein and 26% recovery (Table 8). Purification fold

increased from 1 to 1.74 after dialysis.

Figure 25: Purification folds of the partially purified enzyme

Figure 26: Ammonium sulphate precipitation profile for pectinases isolated from

Aspergillusniger

1

0.63

1.74

Crude Enzyme

After Ammonium sulphate precipitation

After Dialysis

0

10

20

30

40

50

60

70

80

90

0 20 40 60 80 100 120

Act

ivit

y(U

/ml)

% Ammonium Sulphate Saturation

62

Table 9: Summary of Purification step of Pectinases from Aspergillus niger

Purification

Step

Volume

(ml )

Protein

Conc.

(mg/ml)

Activity

(U/ml)

Specific

Activity

(U/mg)

Total

Activity

(U)

Purification

fold

%Yield

Crude enzyme

filtrate 1000 0.486 25.73 52.94 25,73 1.00 100

80% (NH4)2SO4

precipitation 62 2.164 72.63 33.56 4,503 0.63 18

Dialyzed enzyme 70 1.041 95.86 92.08 6,710 1.74 26

63

3.7.2 Changes in Protein Concentration of Partially Purified Enzyme

The protein concentration increased to 2.164mg/ml after ammonium sulphate precipitation

and decreased to 1.041mg/ml after dialysis (Figure 26).

Figure 27: Changes in Protein concentration after partial purification

0.486

2.164

1.041

0

0.5

1

1.5

2

2.5

Crude Enzyme After Ammonium sulphate precipitation

After Dialysis

Pro

tein

con

cen

trati

on

(m

g/m

l)

Purification step

64

3.7.3 Changes in Pectinase Activity During Purification

Pectinaseactivity of the crude enzyme was found to be 25.73. After ammonium sulphate and

dialysis, the value increased to72.63U/ml and 95.86U/ml respectively (Figure 27).

Figure 28: Changes in Pectinase Activity after partial purification

25.73

72.63

95.86

0

20

40

60

80

100

120

Crude Enzyme After Ammonium sulphate precipitation

After Dialysis

Act

ivit

y (

U/m

l)

Purification step

65

3.7.4 Specific Activity of Partially Purified Enzymes

Specific activity of the crude enzyme was found to be 52.94U/mg protein. This value

decreased to 33.56U/mg protein after ammonium sulphate precipitationand increased to

92.08U/mg proteinand dialysis respectively (Figure 28)

Figure 29: Changes in Specific Activity after partial purification

52.94

33.56

92.08

0

10

20

30

40

50

60

70

80

90

100

Crude Enzyme After Ammonium sulphate precipitation

After Dialysis

Spe

cifi

c A

ctiv

ity

(U/m

g)

Purification step

66

3.8 Enzyme Characterization

3.8.1 Effect of pH Change on Pectinase Activity

Results shown in Figure 29indicate that pH 5.0 was more suitable for pectinaseactivityand

has the highest activity of 60.45U/ml. This is known as the optimum pH of the enzyme.

Either increase or decrease in pH beyond the optimum value showed decline in enzyme

activity.

Figure 30:Effect of pH on pectinase activity

0

10

20

30

40

50

60

70

3 3.5 4 4.5 5 5.5 6 6.5 7 7.5 8 8.5 9 9.5

Act

ivit

y (U

/ml)

pH

67

3.8.2 Effect of Temperature Change on Pectinase Activity

At pH 5.0,an increase in temperature was accompanied by an increase in pectinase activity up

to the optimal temperature of 40oC after which the enzyme activity decreased steadily.

Figure 31: Effect of Temperature on Pectinase Activity

0

10

20

30

40

50

60

70

80

90

20 25 30 35 40 45 50 55 60 65 70 75

Act

ivit

y (U

/ml)

Temperature (0C)

68

3.8.3 Effect of Substrate Concentration on Pectinase Activity

The study on the effect of substrate concentrationrevealed that pectinase activity gradually

increases as the substrate concentration increases up to 70mg/ml, after which the enzyme

activity decreases(Figure 31).

Figure 32: Effect of Substrate Concentration on Pectinase Activity

0

20

40

60

80

100

120

140

160

180

0 10 20 30 40 50 60 70 80 90 100

Act

ivit

y (μ

mo

le/m

in)

Substrate concentration (mg/ml)

69

3.8.4 Determination of Kinetic Parameters

The result obtained from the effect of substrate concentration on pectinase activity was used

to plot Lineweaver-Bulks plot. Kinetic parameters such as Vmax and Km of the enzyme were

calculated from the Lineweaver-Burk plot (Figure 32). The Vmax and Km values were found to

be 200µmole/minand 18mg/ml respectively.

Figure 33: Lineweaver-burk plot of pectinases from Aspergillus niger

y = 0.088x + 0.005R² = 0.863

0

0.002

0.004

0.006

0.008

0.01

0.012

0.014

-0.12 -0.1 -0.08 -0.06 -0.04 -0.02 0 0.02 0.04 0.06 0.08 0.1 0.12 0.14

1/V

(μ

mo

le/m

in)-1

1/[S ] (mg/ml)-1

70

3.8.5 Summary of Pectinase Characterization

The summary of the isolated pectinase activity is shown in Table 10. pH and temperature optimum of

5.0 respectively. Vmax of 200μmole/min and km value of 18mg/ml.

Table 10: Summary of Pectinase Characterization

Properties A. niger

pH 5.0

Temperature (0C) 40

Vmax (μmole/min) 200

Km (mg/ml) 18

71

CHAPTER FOUR

4.0 DISCUSSION

Pectinase production constitutes 25% of global enzyme production because of its wide

industrial application. Hence, a lot of research is ongoing for efficient and economical

production of this enzyme. Thus, in the present study, pectin extracted from orange peels was

used to produce extracellular pectinasesunder submerged fermentation system using newly

isolated fungal specie from natural source.Only extracellular pectinases were targeted

because in comparison to intracellular pectinases, extracellular are easier to harvest and

scaling up work can be more easily attempted. Furthermore, as the isolated speciewas to

thrive only on pectin, pectin content of orange fruit was determined.

The percentage yield of pectin from ground orange peels was found to be 15.5% at pH 2.2,

temperature of 70oC and extraction time of 1hour. This yield is to an extent comparable to

17.8% and 18.4% of pectin obtained by Rao and Miani (1999) from sweet orange peels and

Mandarin oranges peels respectively.The slight variations in the yield may have resulted from

environmental differences and many other factors which include extraction technique,

changes in pH, temperature and extraction time (Kertesz 1951; Rehman et al., 2004). This

result indicates that orange peels contain substantial amount of pectin and thus can be used as

substrate for the production of pectinolytic enzymes by microorganisms. The pectin extracted

from orange peels act as the inducer for the production of pectinolytic enzymes. The selection

of orange peels as substrates for the process of enzyme biosynthesis was not only based on

the pectin content but also on the following factors:

1) They represent one of the cheapest agro-industrial wastes.

2) They are available at any time of the year.

3) Their storage constitutes no problem in comparison with other substrate.

From the three pectinolytic fungal species isolated from natural sources (Aspergillus flavus,

Aspergillus fumigatus andAspergillus niger), one of the species producedrelatively higher

pectinase activity on a selective media containing orange pectin. Based on colony

morphology and microscopic examination of the fungal isolates, the specie was identified as

Aspergillus niger (Figure 22)This specie was able to degrade pectin by producing pectinase

enzyme and was selected for further studies. This result indicates that Aspergillusniger is

highly pectinolytic and can be used effectively in pectinase production. Filamentous fungi

72

was chosen for the present studybecause their nature is generally regarded as safe (GRAS)by

United States Food and Drugs Administration (USFDA)(Sumanthaetal., 2005). Also, they are

eukaryotic organisms in comparison with prokaryotic organisms, have wide spectrum of

genetic information, and are able to perform microbial conversion (Kutaleladze etal.,

2009).Aspergillus sp. represents the most common source of commercial pectinases

(Castilhoet al., 1999), likewise many other industrial enzymes. Bacteria are also known to

produce industrial enzymes, but filamentous fungi are desired for the production of enzymes.

The pectin extracted from orange peels was used to induce pectinase synthesis

usingAspergillus nigerunder submerged fermentation. The entire fermentation process was

carried out at room temperature (30oC). Fungi are extracellular organisms and as such,

secrete extracellular enzymes that convert large substrate molecules into smaller molecules

that can be more easily transported into their system. Therefore, the presence of pectin as the

only carbon source induces the organism to secrete extracellular pectinases into the

medium.The accumulation of maximal extracellular enzyme activity was observed after 4

days of fermentation. Similar observationwas also obtained during pectinase production in

submerged fermentation (Yogesh et al. 2009) and solid-state fermentation (Martin et al.,

2004) using Aspergillusniger. Banu etal. (2010) reported maximum polygalacturonase

activity on the 5th

day of fermentation with Penicilliumchrysogenum.Apart from the effect of

the inducers on pectinases production, various other factors related to environment affect the

production of pectinases. Some of them are; concentration of nutrients, pH, temperature,

moisture content and influence ofextraction parameters on recovery of pectinases.These

factors are easier to control in submerged fermentation systemthan in solid state fermentation

(Canel and Moo-Young, 1980; Costa etal., 1998; Castilho etal., 2000), making submerged

fermentation system more suitable for pectinases production.

Highest degree of precipitations was achieved by 80% ammonium sulphate saturation (Figure

25).This precipitate has the highest pectinase activity. Buga et al. (2010) reported 70%

ammonium sulphate saturation for pectinase from Aspergillusniger (SA6) while Adejuwon

and Olutiola (2007) reported 90% ammonium sulphate saturation for pectinase from

Lasidioplodia theobromae. The precipitation occurs in that, proteins in aqueous solutions are

heavily hydrated because of their hydrophilic interaction with water molecules and with the

addition of ammonium sulphate, the water molecules become more attracted to the salt than

to the protein due to the higher charge. This competition for hydration is usually more

favorable towards the salt, which leads to interaction between the proteins, resulting in

73

aggregation and finally precipitation or salting out. The salt concentration at which a protein

precipitates differs from one protein to another. Hence, salting out can be used to fractionate

proteins (Markus and Aaron, 2007).The precipitate gotten after ammonium sulphate

precipitation was desalted by subjecting it to dialysis using buffer solution. After dialysis, it

was observed that the volume of the enzyme inside the dialysis bag increased due to the

diffusion of the buffer solution into the dialysis bag.

The protein concentration of the crude enzyme was assayed and was found to be 0.468mg/ml.

the value rose to 2.164mg/ml after ammonium sulphate precipitation indicating that much

protein were precipitated. After 12hours dialysis the value decreased to 1.041mg/ml (Figure

26). The decrease in protein concentration may be attributed to the diffusion of buffer

solution into the dialysis bag which was observed during dialysis.

The pectinase activity increased after ammonium sulphate precipitation and dialysis because,

more of the enzymes of interest were precipitated with ammonium sulphate (Figure 27).

Secondly, dialysis serves to remove low molecular weight substances and ions e.g.

ammonium sulphate salt that may interfere with the enzyme activity and may account for the

increase in pectinase activity after dialysis.

The specific activity of the crude enzyme was found to be 52.94U/mg protein. This value

decreased to 33.56 U/mg proteinafter ammonium sulphate precipitation and finally increased

to 92.08U/mg proteinafter dialysis (Figure 28). This result agrees with the report by Lukong

et al. (2007) that for a purification procedure to be successful, the specific activity of the

desired enzyme must be greater after the purification procedure than as it was before. The

increase in specific activity is a measure of purification achieved. Pectinases isolated from

Aspergillus niger was partiallypurified approximately 2-fold with 26% recovery (Figure 24).

Purification fold increased from 1 to 1.74 after dialysis showing that the enzyme has received

some level of purification.

The partially purified enzyme was characterized based on effects of pH, temperature change

and substrate concentration on pectinase activities. From the pH studies, as the pH was

increased from pH 3.5 to pH 5.0, the pectinase activity was found to increase. Further

increase in pH beyond pH 5.0, resulted in decrease the pectinase activity(Figure 29). The

optimum pH was therefore found to be 5.0. The optimum pH obtained in this study is

comparable with the polygalacturonase from Aspergillus awamori, Thermococcusauraniacus

and Penicillium frequentans (Jayani et al., 2005). It was also reported that the optimum pH

74

for pectinase activity from thermotolerant Aspergillus sp. N12 was 5.5 (Ramakrishna, et al.,

1982).

The optimum temperature for pectinase activity was found to be at 40°C (Figure 30). Further

increase in temperature beyond 40oC decreased the pectinase activity. The decrease in

enzyme activity at higher temperature may be due to enzyme denaturation. Similar results

were also reported for polygalacturonase by Aspergillusawamori and Aspergillus

niger(Jayani et al., 2005). Exo-polygalacturonase from Monascus and Aspergillussp. (Freitas

et al.,2006) exhibited maximum activity at 60 and 50°C, respectively. The

endopolygalacturonase from Mucour rouxii NRRL 1894 exhibited maximum activity at 35°C

(Saad et al., 2007).

Temperature and pH are highly influential to enzyme activity. This is because proteins fold

into particular shapes that are vital for (and determine) their function. The shape a protein

will fold into is determined by its amino acid sequence, since different amino acids have

different properties. Each amino acid has a side chain sticking out of the main polypeptide

chain, which will have specific chemical properties capable of forming certain interactions

with other amino acids in the protein (as well as with water and other molecules). So,

increasing the temperature increases the energy of the bonds and atoms in the protein, to the

point at which there is enough energy to overcome the force of the intramolecular reactions,

resulting in their breaking. Disruption of the interactions in any case will lead to some of the

protein losing its ability to be held in a certain shape, which then reduces it's catalytic activity

(as catalytic activity relies on the shape).The dual effects of increase in temperature and

protein denaturation beyond the optimum temperature give rise to the bell-shaped nature of

activity curves of most enzymes (Anosike, 2001).

Altering the pH above or below its optimum pH will also reduce the enzyme's activity, and at

extremes the enzyme may be permanently denatured. pH is a measure of the concentration of

hydrogen ions, which are positively charged. If there were more hydrogen ions in the solution

than the protein was designed for, these ions would compete for the interactions holding the

protein together, as well as protonating groups that need to be deprotonated to form important

intramolecular interactions. Equally, if there were too few hydrogen ions in the solution, the

same interactions would be disrupted by the relatively high concentration of hydroxide (OH)

ions, and important protonated groups may become deprotonated. The loss of activity will be

75

proportional to the extent of the disruptions, which will in turn be proportional to the extent

of the change in pH or temperature.

From the double reciprocal plot (Figure 32), the values for the Km and Vmax were found to be

18mg/ml and 200µmole/minrespectively. According to Anosike (2001),Km values provide a

parameter for comparing enzymes from different organisms and also, establish approximate

value of intracellular level of substrate. It establishes a relationship between the enzyme and

its affinity with the substrate. A small Km indicates that the enzyme requires only a small

amount of substrate to become saturated; hence, the maximum velocity is reached at

relatively low substrate concentration while a large Kmindicates the need for high substrate

concentrations to achieve reaction velocity.

The Vmax or maximum velocity gives information on the turnover number of an enzyme

(Anosike, 2001). The turnover number of an enzyme is the number of moles of substrate

converted into product per active site of the enzyme per second, when the enzyme is fully

saturated with substrate. This implies that, pectinase obtained from Aspergillus niger converts

about 200 micromoles of substrates into products per minute.Vmaxalso gives us information

on how efficient a given enzyme is as a catalyst.

76

CONCLUSION

Results obtained from this work indicate that orange peels can be utilized effectively in the

production of pectinases, under submerged fermentation system using Aspergillusniger.

Pectinases produced from this fungal specie has an optimum pH and temperature of 5.0 and

40oC respectively. Km and Vmax were also found to be 18mg/ml and 200µmole/min

respectively.This can be applied industrially for the production of pectinases used for various

purposessuch as fruit processing, vegetable oil extraction, coffee and tea fermentation, paper

making and cotton fabric processing etc. Since orange peels utilized in this process are

readily accessible as waste with little or no cost and also contain an appreciable amount of

pectin, they can be regarded as a low-cost substrate for efficient and economical production

of pectinases usingAspergillusniger. This will not only lead to the reduction in the

production cost of pectinases but also help to decrease the pollution load resulting from these

wastes.

FUTURE RECOMMENDATIONS

Based on the findings in this work, the following recommendations are made:

1. Genetic and environmental modification of the organism used in this work will

equally help to increase the yield in pectinase production. This should be achieved

through genetic engineering.

2. A comparative study between pectin extracted from orange peels and other cheap

agro-waste materials or peels will be beneficial in determining which carbon source is

better for pectinase production.

3. Also, studies on other physicochemical properties of pectinase such as thermal and

pH stability should be conducted to understand their effects on the enzyme activity.

4. Further purification of the enzyme using gel filtration, ion-exchange chromatography,

gel electrophoresis etc. will be essential in understanding properly its biochemical

functions. An adage of biochemistry is, „Never waste pure thoughts on an impure

protein‟.

77

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APPENDIX ONE

1.0 Preparation of 3N HCl

Normality of the stock HCl in Winchester bottle was calculated using the formulae

outlined below:

Normality (N) = % assay x 1000 x specific gravity

100 x Equivalent weight

For HCl, % assay = 37, specific gravity = 1.19 and equivalent weight (in this case) = 36.5.

The volume of the stock solution required to prepare 3N HCl was calculated using the

formula

N1V1 = N2V2

Where: N1 = Normality of stock HCl, V1 = volume of stock HCl required for the preparation,

N2 = required normalty (3N) and V2 = working volume (1000ml). V1 obtained from the

calculation was diluted in 1000ml of diluent (water) to make the required 3N HCl.

1.1 Preparation of Ethanol-HCl Solution

The ethanol-HCl solution contained 0.5M HCl prepared using similar formula as that in 1.0

except that molecular weight was used instead of eqvivalent weight to generate the molarity

of stock HCl. The volume of the stock solution required to prepare 0.5M HCl was calculated

using the formula C1V1 = C2V2;whereC1 = Molarity of stock HCl, V1 = volume of stock HCl

required for the preparation, C2 = required molarity (0.5M) and V2 = working volume

(1000ml). V1 obtained from the calculation was diluted in 1000ml of diluent (ethanol) to

make the required ethanol-HCl solution.

1.2 Preparation of Buffers

The standard buffers used in study were pH 4.0, pH 7.0 and pH 9.2. These buffers were used

to standardize the pH meter. The working buffers were prepared as thus: 0.05M sodium

acetate and 0.05M Tris-HCl buffers were prepared by dissolving 4.10g sodium acetate salt

and 6.01g Tris base, respectively in 1000ml of distilled water and stirred with a magnetic

stirrer till a homogenous solution was formed. The solutions were titrated against acetic acid

and HCl, respectively till the required pHs were obtained. Also 0.05M phosphate buffer was

prepared by dissolving 7.10g disodium hydrogen phosphate salt in 1000ml stirred as for

95

sodium acetate and phosphate buffers and then tritrated againt the solution of its conjugate

acid, sodium dihydrogen phosphate till the required pHs were obtained.

1.3 Preparation of Dinitrosalicylic Acid (DNS) Reagent

A modification of DNS reagent method of Miller (1959) as contained in Wang et al. (1997)

was used in the assay. The reagent contains 44mM dinitrosalicylic acid, 4mM sodium

sulphite, and 375mM sodium hydroxide.

1.4 Preparation of 20mM Galacturonic Acid

0.42g D-(+)-Galaturonic acidmonohydrate (molecular weight 212.15g/mole) was dissolved in

100ml 0.05M sodium acetate buffer stirred over a magnetic stirrer until a homogenous stock

solution was obtained.

1.5 Galacturonic Acid Standard Curve

The reaction mixture contained 0.0-1.0ml of galacturonic acid stock solution in test tubes

arranged in triplicates. Each test tube was made up to 1ml using freshly prepared 0.05M

sodium acetate buffer of pH 5.0. 1ml of DNS reagent was added to each of the test tubes and

placed in a boiling water bath for 10min. 1ml of 1.4M Rochelle salt (sodium potassium

tartarate) was added to the test tube immediately after heating and the total volume of the

solution was adjusted to 4ml with distilled water. The mixture was cooled to room

temperature and the absorbance read at 575nm. The concentration of reducing sugar in each

of the tubes was calculated using the formula

C1 V1 = C2 V2

Where: C1 = initial concentration of reducing sugar (mM)

C2 = final concentration of reducing sugar (mM)

V1 = initial volume of 20mM galacturonic acid preparation measured into the tube

V2 = final volume of the preparation.

Using the values obtained from above the calculations, the plot of optical density was

constructed and the concentration of galacturonic acid released at a given absorbance was

extrapolated (Appendix Two).

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1.6 Preparation of the Component Reagents for Protein Determination

Solution A: An alkaline sodium carbonate (Na2CO3) was prepared by dissolving 2g of

Na2CO3 in 100ml of 0.1M NaOH (0.4g of sodium hydroxide pellets were dissolved in 100ml

of distilled water).

Solution B: A copper tetraoxosulphate IV - sodium potassium tartarate solution was prepared

by dissolving 0.5g of CuSO4 in 1g of sodium potassium tatarate, all in 100ml of distilled

water. It was prepared fresh by mixing stock solution, and so was done whenever required.

Solution C: Folin-Ciocalteau phenol reagent was made by diluting the commercial reagent

with water in a ratio of 1:1 on the day of use.

Solution D: Standard protein (Bovine Serum Albumin, BSA) solution.

Solution E: Freshly prepared alkaline solution was made by mixing 50ml of solutions A and

1ml of solution B.

1.7 Preparation of 2mg/ml Bovine Serum Albumin (BSA) Standard Protein

0.2g of BSA was dissolved in 100ml of distilled water and then used as a protein stock

solution.

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APPENDIX TWO

BIOSAFETY PRECAUTION WHILE PERFORMING EXPERIMENTS

1. Hand gloves and mouth covers were worn while performing all fungal experiments in

sporulating rooms.

2. Sporulating room was properlyfumigated before performing experiments.

3. The benches are also disinfected with ethanol before any experiment.

4. The fungi were discarded by properly autoclaving the flasks after

performingexperiment.

98

APPENDIX THREE

Appendix 2: Galacturonic Acid Standard Curve, Using 20mM D-(+)-Galacturonic Acid

Monohydrate

.

y = 0.146xR² = 0.994

0

0.02

0.04

0.06

0.08

0.1

0.12

0.14

0 0.2 0.4 0.6 0.8 1

Ab

sorb

ance

at

57

5 n

m

Galacturonic acid concentration in mM

Galacturonic acid standard curve

99

APPENDIX FOUR

Appendix 3: Protein Standard Curve, Using 2mg/ml Bovine Serum Albumin (BSA)

y = 0.796xR² = 0.994

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0 0.2 0.4 0.6 0.8 1

Ab

sorb

ance

at

75

0n

m

Protein concentration in mg/ml

Protein standard curve