PRODUCTION AND CHARACTERIZATION OF α-AMYLASE FROM ...

i

PRODUCTION AND CHARACTERIZATION OF α-AMYLASE FROM Aspergillus

nidulans UNDER SUBMERGED FERMENTATION SYSTEM

BY

ENE, CHIBUEZE KELECHI

(PG/MSc./16/81439)

DEPARTMENT OF BIOCHEMISTRY

FACULTY OF BIOLOGICAL SCIENCES

UNIVERSITY OF NIGERIA, NSUKKA

NOVEMBER, 2018.

i

TITLE PAGE

PRODUCTION AND CHARACTERIZATION OF α-AMYLASE FROM Aspergillus

nidulans UNDER SUBMERGED FERMENTATION SYSTEM

A DISSERTATION SUBMITTED TO THE DEPARTMENT OF

BIOCHEMISTRY, UNIVERSITY OF NIGERIA, NSUKKA, IN PARTIAL

FULFILLMENT OF THE REQUIREMENTS FOR AWARD OF DEGREE OF

MASTER OF SCIENCE (M.Sc.) IN BIOCHEMISTRY (ENZYMOLOGY AND

PROTEIN CHEMISTRY) OF THE UNIVERSITY OF NIGERIA, NSUKKA.

BY

ENE, CHIBUEZE KELECHI

(PG/MSc./16/81439)

SUPERVISOR: PROF. F. C. CHILAKA

ii

CERTIFICATION

This is to certify that Ene, Chibueze Kelechi a postgraduate student of the Department of

Biochemistry, University of Nigeria, Nsukka with Registration Number, PG/MSc/16/81439, has

satisfactorily completed the requirements of the award of the degree of Master of Science (M.Sc.)

in Enzymology and Protein Chemistry. The work embodied in this dissertation is original and has

not been submitted in part or full for any other diploma or degree of this or any other University.

…………………..….. ……………..…….. …………………….………………..

Prof. F. C. Chilaka Prof. F. C. Chilaka

(Head of Department) (Supervisor)

---------------------------------------------------

External Examiner

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DEDICATION

This work is dedicated to my beloved parents.

iv

ACKNOWLEGDEMENT

I wish to express my profound gratitude to the Almighty God for the gift of life and good health

and for thus far He has brought me.

I sincerely thank my supervisor Prof. F. C. Chilaka whose support, encouragement, dedication,

patience and constructive criticism made this work a success.

My gratitude goes to all the lecturers and members of staff of Department of Biochemistry,

University of Nigeria, Nsukka for their dedication and selflessness in discharging their duties.

I want to specially thank Dr A. L. Ezugwu, Abugu Sylvester, Ezike Tobechuckwu and Chigbu

Michael for their immense help and contributions during this research work. I am equally very

grateful to Ukwuenu Prosper, Ottah Victoria, Iroha Okechukwu, Omeje Kingsley, Okagu

Innocent, Udeh Jerry, and many other senior friends for their useful advice, contributions and

encouragement in the course of this work. I wish to express my thanks to every member of my

research group under the supervision of Prof. F. C. Chilaka as well as my colleagues for their

contagious drive for excellence.

To my wonderful parents, Mr and Mrs Ignatius Enembagu and caring siblings, Henry, Onyedika,

Onuora and Ezinne, I say thank you for your unquantified love and support. You are simply the

best.

I cease this opportunity to appreciate my brethren at Graduate Student Fellowship (GSF). Indeed,

you have been a home away from home.

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ABSTRACT

Three fungi isolates were obtained from soil collected from a laundry waste water disposal site.

These organisms were identified morphologically as Aspergillus niger, Aspergillus nidulans and

Trichoderma hazianum. Ten days pilot study on these organisms indicated that A. nidulans

produced α-amylase of highest activity on day 6 and was therefore chosen for further studies. The

internal transcribed spacer (ITS) region of the organism showed an amplicon size of approximately

500-600 bp from agarose gel electrophoresis. Optimum pH for α-amylase production was observed

to be pH 5.0 with an activity of 140.706 µmolmin-1. Forty percent (40%) Ammonium sulphate

saturation was found suitable to precipitate protein with maximum α-amylase activity. The enzyme

was further purified through dialysis and gel filtration, giving purification fold of 5.99 and

percentage yield of 19.62. The specific activity increased from 371. 43 to 2226.37 U/mg showing

that the purification processes were ideal for the enzyme. The total protein decreased with the

purification processes showing that low molecular weight proteins and other contaminants were

removed with purification steps. The optimum pH and temperature of the purified α-amylase were

5.0 and 60°C, respectively. The Michaelis constant, KM (18.28mg/ml) and maximum velocity, Vmax

(144.927µmol/min) were obtained from the Lineweaver-Burk plot at different substrate

concentrations. The study revealed that Co2+, Cu2+, Mg2+ and Fe2+ inhibited α-amylase activity

whereas Ca2+ and Mn2+enhanced its activity. From the result of pH stability study, more than 50

% of the enzyme’s initial activity was retained at pH range of 4.0 to 9.0 after 60 min of incubation

and was most stable at pH 5.0, with residual activity of 99 % after 60min and 90 % at 120 min.

The thermal stability study revealed that the enzyme was most stable within 40 to 50°C with

residual activity of 89 % at 40°C and 84% at 50°C after 120 min of incubation. At the optimum

temperature, more than 50% of its initial activity was maintained after 90 min. The half-life and

D-value decreased with temperature increase. The activation energy of denaturation Ea and Z-

values were 66.793 Kjmol-1 and 31.25°C, respectively. At optimum temperature, the results of the

thermodynamic parameters, ΔH0, ΔG0, and ΔS0 were 64.024, 53.242 and 0,039 Kjmol-1,

respectively. The results of the study showed that Aspergillus nidulans can be exploited as a cheap

and efficient source of α-amylase for many biotechnological industries that operate within these

observed conditions.

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TABLE OF CONTENTS

Title Page i

Certification ii

Dedication iii

Acknowledgement iv

Abstract v

Table of Contents vi

List of Figures xii

List of Tables xiii

List of Plates xiv

List of Abbreviations xv

CHAPTER ONE 1

INTRODUCTION 1

1.1 Alpha amylases (α-amylase) 2

1.2 Cassava starch as a substrate for α-amylase 2

1.3 Sources of α-amylase 5

1.3.1 Plant α-amylases 5

1.3.2 Animal α-amylases 6

1.3.3 Bacterial α-amylases 7

1.3.4 Fungal α-amylases 8

1.3.4.1 Fungi of Aspergillus genus as an enzyme source 9

1.3.4.2 Morphological identification of Aspergillus species 10

1.3.4.3 Molecular identification of Aspergillus species 10

1.3.4.4 Taxonomy of Aspergillus 11

1.4 Structural and functional characteristics of α-amylase 11

1.5 α-Amylase active site structure and function 13

vii

1.6 Mode of action of α-amylase 14

1.7 Reaction mechanism of α-amylase 14

1.8 Production of α-amylase 15

1.8.1 Submerged fermentation 16

1.8.2 Solid state fermentation 16

1.9 Process parameters affecting α-amylase production 18

1.9.1 Temperature 18

1.9.2 pH 19

1.9.3 Carbon sources 20

1.9.4 Nitrogen source 20

1.9.5 Duration of fermentation 20

1.9.6 Metal ions 21

1.9.7 Moisture 21

1.10 Isolation and purification of fungal α-amylase 22

1.10.1 Ammonium sulphate precipitation 22

1.10.2 Gel filtration 22

1.11 α-Amylase assay 23

1.12 Biochemical properties of α-amylase 23

1.12.1 Substrate specificity 23

1.12.2 Temperature optima and stability 24

1.12.3 pH optima and stability 24

1.12.4 Effect of metal ions 24

1.12.5 Effect of inhibitors 26

viii

1.12.6 Molecular weight 26

1.12.7 Stabilization of α-amylase 26

1.13 Applications of α-amylase 26

1.13.1 Production of fructose and glucose syrup 27

1.13.2 Bakery industry 27

1.13.3 Detergent industry 28

1.13.4 Desizing of textiles 28

1.13.5 Paper industry 29

1.13.6 Feed industry 29

1.13.7 Bioethanol production 29

1.14 Aim and Objectives 30

1.14.1 Aim 30

1.12 Specific Objectives of the Study 30

CHAPTER TWO 31

2.0 MATERIALS AND METHODS 31

2.1 Materials 31

2.1.1 Reagents 31

2.1.2 Apparatus 31

2.1.3 Collection of plant material 32

2.1.4 soil samples 32

2.2 Methods 32

2.2 1 Processing of cassava starch 32

2.2.2 Isolation of α-amylase producing fungi 32

ix

2.2.2.1 Serial dilution agar plate technique for isolation of fungi 32

2.2.2.2 Storage of pure fungal isolates 33

2.2.2.3 Examination of isolated fungi 33

2.2.2.4 Fungal identification 34

2.2.3 Molecular characterization of fungal isolates 34

2.2.3.1 DNA extraction from the microorganism 34

2.2.3.2 PCR amplification of the ITS region 34

2.2.3.3 DNA sequencing 35

2.2.3.4 BLAST analysis of DNA sequences 35

2.2.3.5 Phylogenetic analysis 35

2.2.4 Fermentation experiments 35

2.2.4.1 Preparation of fermentation broth 35

2.2.4.2 Inoculation of broth 35

2.2.4.3 Harvesting of fermented broth 36

2.2.4.4 Mass production of enzyme 36

2.2.5 Assays for α-amylase activity using dinitrosalicyclic acid (DNSA) reagent 36

2.2.6 Effect of pH on α-amylase production 36

2.2.7 Determination of protein content 37

2.2.7.1 Procedure for protein determination 37

2.2.8 Purification of the enzyme 37

2.2.8.1 Determination of percentage ammonium sulphate saturation suitable

for ɑ-amylase precipitation 37

2.2.8.2 Mass ammonium sulphate precipitation of ɑ-amylase 37

x

2.2.8.3 Dialysis 38

2.2.8.4 Gel filtration chromatography 38

2.2.9 Studies on purified enzyme 38

2.2.9.1 Effect of pH on α-amylase activity 38

2.2.9.2 Effect of temperature on α-amylase activity 39

2.2.9.3 Effect of metal ions on α-amylase activity 39

2.2.9.4 Effect of substrate concentration on α-amylase activity 39

2.2.10 pH stability 39

2.2.11 Thermal inactivation studies 39

2.2.11.1 Determination of percentage residual activity 40

2.2.11.2 Denaturation constants (Kd) of the enzyme fraction 40

2.2.11.3 Half-life (t1/2) of the enzyme solution 40

2.2.11.4 Activation energy of denaturation Ea(D) 40

2.2.12 Thermodynamics parameters 41

CHAPTER THREE 42

RESULTS 42

3.1 Microorganisms 42

3.1.1 Selection of amylolytic fungi 42

3.1.2 Macroscopic and microscopic examination of fungal isolates 42

3.2 Production of α-amylase and study on crude 42

3.2.1 Effect of incubation period on α-amylase production 42

3.2.2 Molecular characterization of selected fungal isolate 42

3.2.3 Effect of pH of medium on α-amylase production 43

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3.2.4 Protein concentration of crude enzyme 41

3.2.5 α-Amylase activity of crude enzyme 41

3.3 Purification of crude α-amylase 43

3.3.1 Ammonium sulphate precipitation profiles of α-amylase 43

3.3.2 Gel Filtration of α-amylase 43

3.4 Characterization of α-amylase 43

3.4.1 Effect of pH on α-amylase activity 43

3.4.2 Effect of temperature on α-amylase activity 44

3.4.3 Effect of metal ions on α-amylase activity 44

3.4.4 Effect of substrate concentration on α-amylase activity 44

3.4.5 Determination of kinetic parameters for substrate concentration 44

3.4.6 Effect of pH stability on α-amylase activity 44

3.4.7 Thermal denaturation study of the enzyme 44

3.4.7.1 Percentage residual activities of the enzyme 44

3.4.7.2 Calculation of denaturation constants (Kd) of the enzyme 45

3.4.7.3 Calculation of half-life (t1/2) of the enzyme 45

3.4.7.4 Calculation of D-values of the enzyme 45

3.4.7.5 Calculation of Z-values of the enzyme 45

3.4.7.6 Calculation of activation energy of inactivation of the enzyme 45

3.5 Thermodynamics parameters of the enzyme 45

CHAPTER FOUR 65

4.0 Discussion 65

4.1 Conclusion 73

xii

4.2 Suggestions for further studies 73

REFERENCES 74

APPENDIX 84

xiii

List of figures

Figure 1: Structure of A) amylose and B) amylopectin 3

Figure 2: Modes of action of amylolytic enzymes 4

Figure 3: Structure of α-amylase 13

Figure 4: Structural organization of subsites of α-amylase active sites 15

Figure 5: The α-amylase reaction mechanism 16

Figure 6: Photographs of pure cultures of A. niger, A. nidulans and Trichoderma harzianum 46

Figure 7: Comparison of α-amylase production by the three organism based on incubation

Period 47

Figure 8: Agarose gel electrophoretogram of amplified ITS region 48

Figure 9: Effect of pH of medium on α-amylase production 59

Figure 10: Ammonium sulphate precipitation profile for α-amylase activity from A. nidulans 50

Figure 11: Gel filtration elution profile for α-amylase 51

Figure 12: Effect of pH on α-amylase activity 53

Figure 13: Effect of temperature on α-amylase activity 54

Figure 14: Effect of some divalent metal ions on α-amylase activity 55

Figure 15: Effect of substrate concentration on α-amylase activity 56

Figure 16: Lineweaver-Burk plot of α-amylase from Aspergillus nidulans 57

Figure 17: % Residual activity of α-amylase at pH 3.0 to 10.0. 58

Figure 18: % Residual activity of α-amylase at pH 5.0 and temperatures 40 to 80°C 59

Figure 19: Log % Residual activity of α-amylase temperature of 40 to 80°C 60

Figure 10: A graph of log of D-value against temperature (°C) for the determination of the Z-

value 61

Figure 21: Arrhenius plot of ln Kd against 1/T (K) for the determination of the Activation

energy of thermal deactivation (Ea) 62

Figure 22: Protein standard curve 86

Figure 23: Glucose standard curve 87

xiv

List of tables

Table 1: α-Amylase producing bacteria isolated from different sources 8

Table 2: Amylase producing fungi isolated from different sources 9

Table 3: Bacterial sources and the mode of fermentation used for enzyme production 17

Table 4: Fungal sources and the mode of fermentation used for enzyme production 18

Table 5: Concentration effect of metal ions on the activity of α-amylases 25

Table 6: The purification table of α-amylase from Aspergillus nidulans 52

Table 7: Kinetic parameters for thermal inactivation of α-amylase 63

Table 8: Thermodynamic parameters for thermal inactivation of α-amylase 64

xv

List of plates

Plate A: Pure culture of Aspergillus niger 42

Plate B: Pure culture of Aspergillus nidulans 42

Plate C: Pure culture of Trichoderma harzianum 42

xvi

List of abbreviations

µl microliter

µmol micromole

bp Base pair

cm centimeter

DNA Deoxyribonucleic acid

gDNA genomic deoxyribonucleic acid

g gram

h hour

K kelvin

kDa kiloDalton

KJ kilojoule

l litre

ITS Internal transcribed spacer

M molar

mg miligram

mM milimole

min minutes

nm nanometer

PCR Polymerase Chain Reaction

PDA Potato Dextrose Agar

rDNA Ribosomal deoxyribonucleic acid

rRNA Ribosomal ribonucleic acid

xvii

rpm Revolution per minute

s seconds

sp. Species

Uv Ultraviolet light

1

CHAPTER ONE

INTRODUCTION

Owing to their specificity and catalytic efficiency, enzymes are today applied in many industries

to obtain a valuable final product. This has stimulated interest in the exploration of known and

potentially useful enzymes for biotechnological applications (Saranraj and Stella, 2013). Amylase

was the very first enzyme to be produced industrially in the year 1833 and was used for the

treatment of digestive disorders (Akansha and Varsha, 2013). Alpha Amylase (E.C.3.2.1.1)

catalysis the random hydrolysis of internal α-1,4-glycosidic linkages in starch to low molecular

weight products, such as glucose, maltose and maltotriose units (Paula and Pérola, 2010). It is an

enzyme with diverse applications and constitutes a class of industrial enzymes of approximately

25 % of the world enzyme market (Paula and Pérola, 2010; Rajendra et al., 2016). Alpha Amylase

(α-amylase) has found application in food industries, textile, paper, detergent, and pharmaceutical

industries. However, with the advances in biotechnology, its application has expanded to many

fields such as clinical, medicinal and analytical chemistry (Paula and Pérola, 2010; Saranraj and

Stella, 2013).

Today, amylases are commercially available and this has caused an almost complete shift from

chemical hydrolysis of starch in starch processing industries to eco-friendlier enzyme hydrolysis.

Microorganisms in recent years have been a biotechnological source of α-amylase because of their

many benefits over plant and animal sources. The use of microorganisms for the production of α-

amylase is mainly because of economical bulk production capacity and the fact that microbes are

easily engineered to obtain enzymes of desired characteristics (Saranraj and Stella, 2013). Among

the microorganisms capable of secreting α-amylase, fungi are preferred because of their more

acceptable GRAS (generally regarded as safe) status. In addition, the hyphal mode of growth, good

tolerance to low water activity (𝑎𝑤) and acidity which allows the avoidance of bacterial

contamination make fungi most efficient for enzyme production (Jiby et al., 2016).

Solid state fermentation (SSF) method has in recent years gained attention for production of

enzymes. Compared to submerged fermentation, Solid state fermentation proves to be more

simple; requiring lower capital, simple fermentation media, absence of rigorous control of

2

fermentation parameters, uses less water and so produces lower wastewater (Saranraj and Stella,

2013; Priya and Mrunal, 2016). However, submerged fermentation (SmF) which was the very first

method adopted for industrial enzyme production, is still very relevant for enzyme production

because of better monitoring, ease of sterilization of the medium and purification process (Saranraj

and Stella, 2013). Virtually all the large-scale enzyme producing facilities uses the proven

technology of SmF (Jiby et al., 2016).

With constant rise in demand of α-amylase in different industries, there is need for producing one

with better characteristics suitable for industrial applications. Therefore, the current study focuses

on the production, purification and characterization of α-amylase from organism isolated from a

laundry waste water disposal site.

1.1 Alpha amylases (α-amylase)

Alpha amylases (E.C.3.2.1.1) are extracellular enzyme group that hydrolysis the interior α-1,4-

glycosidic linkages in starch. They belong to the enzyme class of hydrolases and they are a calcium

metalloenzyme, completely unable to function in the absence of calcium (Ajita and

Thirupathihalli, 2014). Hydrolysis of internal α-1,4-glycosidic linkages by α-amylase occurs

randomly and generates maltotriose and maltose from amylose, or maltose, glucose and "limit

dextrin" from amylopectin (Saranraj and Stella, 2013). α-Amylases being endo-amylases act along

the chain length of its substrate and therefore, tends to be faster-acting than glucoamylase which

is the exo-amylase (Jiby et al., 2016). Terminal glucose residues and α-1,6-linkages cannot be

cleaved by α-amylase (Gupta et al., 2008; Ajita and Thirupathihalli, 2014). In human physiology,

both salivary and pancreatic amylases function as digestive enzymes and operate at optimum pH

of 6.7 - 7.0 (Tiwari et al., 2015).

1.2 Cassava starch as a substrate for α-amylase

Starch is a polymer of anhydrous glucose units which are typically accumulated in the unique and

independent granules (Zia et al., 2017). It is the most abundant carbohydrate reserve in plants and

is distributed in different parts such as leaves, flowers, fruits, seeds, stems and roots (Sandoval and

Fernández, 2013). Structurally, as shown in Figure 1, starch consists of two polysaccharides;

amylose (20 – 30 %) and amylopectin (70 – 80 %). Both consist of chains of α-(1,4)-linked D-

glucose residues, which are interconnected through α-(1,6) glucosidic linkages, thus forming

branches in the polymers (Eric, 2017). In biotechnological application, α-amylases display highest

3

specificity to starch more than any other type of polysaccharide (Saranraj and Stella, 2013).

Cassava (Manihot esculenta) starch is predominantly found in the roots of the crop and is a highly

suitable material for industrial use. This is because of its high carbohydrate content (80–90 % dry

basis), consisting almost entirely of starch (Zia et al., 2017). Industrially, cassava starch plays an

important role as raw material for bioethanol production among other applications. One of its

advantages as an industrial choice is the possibility of being isolated in a pure form, having low

contamination by non-starch components. Its functional properties include; low gelatinization

temperature, non-cereal flavor, high viscosity, high water binding capacity, bland taste, translucent

paste and a relatively good stability (Oluwatoyin et al., 2018).

A

B

Figure 1: Structure of A) amylose and B) amylopectin (Dipak, 2016)

4

Other enzymes as shown in Figure 2 participate in starch conversion. They include; β-amylase, γ-

amylase, debranching enzymes and transferases. β-Amylases (EC 3.2.1.2) unlike α-amylases are

exo-hydrolases acting from the non-reducing end of a polysaccharide chain (Ajita and

Thirupathihalli, 2014; Ritu et al., 2017). β-amylase catalyzes the hydrolysis of the second α-1,4-

glycosidic bond, cleaving off two glucose units (maltose) at a time. β-amylase cannot cleave at the

branched point and therefore generates limit dextrin unit on hydrolysis of highly branched

polysaccharide such as glycogen or amylopectin (Ajita and Thirupathihalli, 2014). The sweetness

and flavor that accompany fruit ripening is as a result of starch breakdown to maltose by β-amylase

(Ritu et al., 2017). γ-amylase (EC 3.2.1.3) in addition to cleaving the terminal α-1,4-glycosidic

linkages at the non-reducing end of amylose and amylopectin, will also cleave α-1,6-glycosidic

linkages yielding glucose (Ajita and Thirupathihalli, 2014). Unlike the other forms of amylase, γ-

amylase is most efficient in acidic environments and has an optimum pH of 3 (Ajita and

Thirupathihalli, 2014).

Figure 2: Modes of action of amylolytic enzymes (Djekrif et al., 2016)

5

1.3 Sources of α-amylase

Plants, animals and microorganisms are all good sources of α-amylase, but the production from

plants and animals are rather limited for several reasons (Asad et al., 2011). The concentrations of

enzymes in plant materials are generally low and usually, starch processing industries require large

quantities of enzymes (Asghar et al., 2007). α-Amylases from animal origin are usually limited to

by-product of the meat industry and therefore its supply is also small (Hmidet et al., 2010).

Microbial sources, mainly fungi and bacteria, are presently the preferred sources for its industrial

production due to advantages such as cost effectiveness, consistency, less time and space required

for production and ease of process modification and optimization (Swetha et al., 2006;

Mohammadabadi and Chaji, 2012; Ahmadi, 2012; Saranraj and Stella, 2013). In plants, animals

and microbes, α-amylases play an important role specifically for carbohydrate metabolism. There

are many reports on its isolation, identification and characterization from these sources (Asghar et

al., 2007; Bakri et al., 2009; Hmidet et al., 2010; Asad et al., 2011; Ahmad et al., 2013).

1.3.1 Plant α-amylases

There are three families of α-amylase genes that play a role in carbohydrate metabolism in plants

and the relative involvement of α-amylases and other enzymes in starch degradation probably vary

between plant species (Duncan et al., 2005).

The family one α-amylase is characterized by a secretory signal peptide. All the well-characterized

cereal grain α-amylases fall within this family. This family of α-amylase is involved in the

breakdown of extracellular starch in the endosperm of cereal grains and dicotyledonous seeds and

probably in diseased tissue where cell death has occurred (Nakajima et al., 2004; Duncan et al.,

2005). The induction of family one α-amylase gene expression is promoted in response to the

phytohormone gibberellic acid and repressed by abscisic acid (Arpana et al., 2012). Similarly, at

the beginning of seed germination, there is active metabolism and a rise in respiration rate which

causes rapid sugar exhaustion in embryo. This triggers the expression of α-amylase genes in

general and degradation of starch in this tissue (Arpana et al., 2012). Since sugar reduces quantity

of gibberellic acid, sugar starvation is suggested to be primary factor in initiating the synthesis of

phytohormone gibberellic acid in the embryo which in turn brings about a remarkable up-shoot in

6

sugar and α-amylase level. Therefore, the embryo’s nutritional requirement at early phase of its

growth is met (Arpana et al., 2012).

The second family of α-amylase gene has no signal peptide and therefore is thought to be localized

in the cytoplasm. They are thought to degrade a supposed cytosolic α-glucan, or a heteroglycan

(Fettke et al., 2004). It is possible that family two α-amylase is involved in general stress responses.

For example, cold acclimation in plants includes an increase in soluble sugars and

hexosephosphates in the cytosol, and could lead to a transitory increase in cytosolic α-glucan,

maintaining a high concentration of sugar within the cytosol, and limiting the need to transport

sugar across plastid membranes (Duncan et al., 2005).

The third family is characterized by a large N-terminal domain, typically 400-500 amino acids in

length, which contains a predicted chloroplast transit peptide (Duncan et al., 2005). Family three

α-amylases may also be responsible for degrading plastid bound starch in starch storage tissues

and in leaves in other plant species (Blennow et al., 2002).

1.3.2 Animal α-amylases

In animals, α-amylase occurs in pancreas, parotid, serum, urine and occasionally in smaller

amounts in other tissues or tumors (Whitcomb and Lowe, 2007). Animal α-amylase consists of a

single chain protein to which a carbohydrate is attached. Although, some species have an isoform

with no carbohydrate attached (Ferey-Roux et al., 1998). Salivary amylase initiates carbohydrate

digestion in the mouth and pancreatic amylase is the main enzyme for luminal digestion of

carbohydrate in the small intestine. Human pancreatic α-amylase is a protein of about 57 kDa and

512 amino acids which includes a signal sequence (Whitcomb and Lowe, 2007). Human pancreatic

juice amylase has no sugar groups and exists as two isoforms termed HPA I and HPA II (Ferey-

Roux et al., 1998). Salivary amylase is coded for by the AMY1 gene and pancreatic amylase by

AMY2; a third form present in some tumors is termed AMY2B (Ferey-Roux et al., 1998).

The animal α-amylase contains three domains termed A, B, and C from the amino terminal with

C being a globular domain of unknown function. The active site is located in a cleft between the

A and B domains (Whitcomb and Lowe, 2007). Calcium and chloride ions bind to the A domain

and may stabilize the active site. The active center contains 5 subsites which bind different glucose

residues in the substrate (Ferey-Roux et al., 1998; Whitcomb and Lowe, 2007).

7

1.3.3 Bacterial α-amylases

α-Amylase has been produced by bacterial of genus such as Bacillus (Muralikrishna and Nirmala,

2005; Asghar et al., 2007), Pseudomonas (Haq et al., 2002) and Clostridium (Asghar et al., 2007).

However, bacterial α-amylase has been derived majorly from the genus Bacillus. Bacillus

licheniformis, Bacillus stearothermophilus, and Bacillus amyloliquefaciens have all been used to

produce α-amylase and applied in industries such as food, textile and paper industries (Konsoula

and Liakopoulou-Kyriakides, 2007; Paula and Pérola, 2010).

In most industrial processes requiring enzymes, thermostability is a desired characteristic. As

enzymatic liquefaction and saccharification of starch are performed at high temperatures (100–

110oC), thermostable amylolytic enzymes have been currently investigated to improve industrial

processes of starch degradation in order to produce valuable products like glucose, crystalline

dextrose, dextrose syrup, maltose and maltodextrins (Asghar et al., 2007). Bacillus subtilis,

Bacillus stearothermophilus, Bacillus licheniformis, and Bacillus amyloliquefaciens are known to

be good producers of thermostable α-amylase, and these have been widely used for commercial

production of α-amylase for various applications (Paula and Pérola, 2010).

Also, α-amylase produced by some halophilic microorganisms which have optimal activity at high

salinities are employed in many harsh industrial processes where the concentrated salt solutions

would otherwise inhibit enzymatic conversions (Prakash et al., 2009). Usually, halophilic bacteria

are fairly thermotolerant and remain stable at mild temperature of about 25-300C over long periods.

Halophilic α-amylases have been characterized from halophilic bacteria such as

Chromohalobacter sp., Halobacillus sp., Haloarcula hispanica, Halomonas meridiana, and

Bacillus dipsosauri (Deutch, 2002; Amoozegar et al., 2003; Hutcheon et al., 2005; Prakash et al.,

2009).

Table 1: α-Amylase producing bacteria isolated from different sources (Dipak, 2016)

Bacteria Sources

8

Bacillus thermooleovorans Hot spring

Bacillus pseudofirmus, Bacillus cohnii, Bacillus vedderi,

Bacillus agaradhaerens, Nesterenkonia halobia

Soil and water of soda lakes

Halobacillus sp. Traditional fermented foods

Bacillus subtilis Saline soil

Caldimonas taiwanensis Traditional fermented food

Anoxybacillus amylolyticus Hot spring

Bacillus sphaericus Geothermal soil of active

fumaroles

Serratia marcescens Hot spring

Chromohalobacter sp. Seas and lake

Bacillus licheniformis, Bacillus subtilis Solar evaporated saltern pond

Bacillus megaterium Digestive tract of fish

Bacillus agaradhaerens Soil

Bacillus licheniformis, Gracilibacillus sp. Salt-enriched soil

Pseudomonas luteola Salt lake

Bacillus amyloliquefaciens Olive washing wastewater

contaminated soil

Corynebacterium alkanolyticum Rhizosphere of plant

1.3.4 Fungal α-amylases

In addition to availability, high productivity and suitability for genetic manipulations, fungi have

gained much interest as source of α-amylase as a result of its GRAS (generally regarded as safe)

status (Iftikhar et al., 2013). Similarly, tolerance to acidity (pH < 4) which allows the avoidance

of bacterial contamination has increased the focus on fungi as the choice for α-amylase production

(Paula and Pérola, 2010). Industrial production of α-amylase has concentrated on fungi species

that thrive at moderate temperatures and attempts have been made to specify the cultural conditions

and superior strains of fungi to produce α-amylase on a commercial scale (Gupta et al., 2008).

Also, fungal sources have mostly been confined to terrestrial isolates such as Aspergillus and

Penicillium (Paula and Pérola, 2010). The Aspergillus species produce a large variety of

9

extracellular enzymes, and amylases are the ones with most significant industrial importance

(Polizeli et al., 2016). Filamentous fungi, such as Aspergillus oryzae and Aspergillus niger,

produce considerable quantities of enzymes that are used extensively in the industry (Paula and

Pérola, 2010).

Thermostable α-amylase have been purified from thermophilic fungus such as Thermomyces

lanuginosus, Hemicola insolens, H. lanuginosa, H. stellata, Mucour pusillus and Talaromyces

thermophilus (Jensen et al., 2002; Kunamneni et al., 2005; Iftikhar et al., 2013)

Table 2: Amylase producing fungi isolated from different sources (Dipak, 2016)

Fungi Isolated from

Penicillium fellutanum Rhizosphere soil of

mangrove plant

Penicillium rugulosum Soil

Penicillium janthinellum Soil

Penicillium expansum Waste fruit kernels

Aspergillus niger Contaminated field soil

Pestalotiopsis microspora, Aspergillus oryzae Leaves of mangrove plants

Penicillium chrysogenum, Fusarium

incarnatum, Penicillium polonicum

Saltwater lake

Aspergillus fumigatus Soil

Aspergillus flavus Soil sample of solid waste

dump site

1.3.4.1 Fungi of Aspergillus genus as an enzyme source

Fungi from Aspergillus genus comprises of a diverse group of species sharing morphological,

physiological and phylogenetic features. They are widely distributed on the soil. In biochemistry

and biotechnology, they serve as an essential tool because they can be easily isolated and studied

(Giraldo et al., 2014). The number of recognized species of Aspergillus genus continues to increase

as a result of the construction of phylogeny by biochemical and molecular tools, and the application

of the phylogenetic species concept. Presently, about 399 species of Aspergillus genus have been

identified by ITS, calmoduli, β-tubulin and RPB2 sequences (Samson et al., 2014). Aspergillus

10

species are used in the fermentation industry and several of them such as Aspergillus niger, A.

oryzae, A.flavus have already established status as cell factories for the production of enzymes and

other metabolites (Polizeli et al., 2016).

1.3.4.2 Morphological identification of Aspergillus species

Morphological identification of Aspergillus species is based on macro-morphology and micro-

morphology (Hina et al., 2013). Macroscopic identification of Aspergillus species is done with the

naked eyes by using distinct features such as the diameter of the colony, conidial colour, colony

reverse, exudates. On the other hand, microscopic characterization is based on their structural

properties as seen under the microscope. These microscopic features include, conidiophore,

vesicle, metulae, phialides and conidia. Although molecular method of identification of organism

continues to improve and is becoming more readily available, microscopic and macroscopic

methods are still relied upon as essential tools for identification of Aspergillus sp (Hina et al.,

2013).

1.3.4.3 Molecular identification of Aspergillus species

At the specie level, morphological character may be contentious or problematic as they may not

always provide accurate groupings within an evolutionary framework (Huzefa et al., 2017).

Factors such as hybridization, convergent evolution, cryptic speciation and limited phenotypic

character makes this method inaccurate (Hughes et al., 2013; Lücking et al., 2014).

Molecular identification of fungi began about two decades ago. It is a method of identification

based on the DNA sequence. The molecular methods are divided into; DNA barcoding which

makes use of the internal transcribed spacer (ITS) region and DNA taxonomy method which makes

use of one or multiple genes in sequence alignment, after which, a tree building tool is used to

compare and estimate phylogenetic relationship (Huzefa et al., 2017).

Three nuclear ribosomal genes are commonly used for molecular fungal identification; nrLSU-

26S or 28S, nrSSU-18S and ITS1, 5.8S, ITS2 for higher taxonomic level, intermediate taxonomic

level and specie taxonomic level identification respectively (Schoch et al., 2012). The ITS is the

most useful for specie-level identification, because it is the fastest evolving portion of the rRNA

cistron. Other factors such as ease of amplification, widespread use, and appropriately large

11

barcode gap (the difference between interspecific and intraspecific variation), makes ITS the

official barcode for fungi by a consortium of mycologists (Schoch et al., 2012; Huzefa et al., 2017).

1.3.4.4 Taxonomy of Aspergillus

Domain: Eukaryota

Kingdom: Fungi

Phylum: Ascomycota

Subphylum: Pezizomycotina

Class: Eurotiomycetes

Order: Eurotiales

Family: Trichocomaceae

Genus: Aspergillus

Source: (Samson et al., 2007)

1.4 Structural and functional characteristics of α-amylase

The amino acid residues of α-amylase in different organisms share approximately 30 % homology

to all glycosyl hydrolase family 13 (GH-13 family) of protein (Tiwari et al., 2015). The three

dimensional (3D) structures of α-amylase revealed a calcium containing enzyme of single

polypeptide chain folded into three domains A to C as shown in Figure 3 (Tiwari et al., 2015).

Domain A is the most conserved domain and the amino acid residues of the α-amylase family that

are involved in catalysis and substrate binding are located in loops at the carboxyl terminal of β-

strands in this domain (Arpana et al., 2012; Tiwari et al., 2015). It (domain A) consists of a highly

symmetrical fold of eight parallel β-strands arranged in a barrel encircled by eight α-helices which

is common to all enzymes belonging to the α/β barrel family (Tiwari et al., 2015).

Domain B of α-amylase is linked to central domain A by a disulfide bond and has a segment that

protrudes between β-sheet number 3 and α-helix number 3. It ranges from 44 to 133 amino acid

12

residues and participate in substrate or calcium binding (Nielsen et al., 2004; Tiwari et al., 2015).

The sequence of this B-domain varies most. In Bacillus α-amylases, it is relatively long and folds

into a complex structure of β-strand, whereas in barley α-amylase, there is an irregularly structured

domain of 64 residues (Tiwari et al., 2015). Common to most α-amylases is a conserved Ca2+

binding site which is located at the interface between domains A and B (Paula and Pérola, 2010;

Arpana et al., 2012; Tiwari et al., 2015). Additionally, α-amylase produced by Bacillus

thermooleovorans was found to contain a chloride binding site close to the active site (Malhotra

et al., 2000), which proved to enhance the catalytic efficiency of the enzyme, presumably by

elevating the pKa of the hydrogen donating residue in the active site (Prakash and Jaiswal, 2009).

The C domain consists of β-sheets connected to the A domain by a small polypeptide chain and

appears to be a separate domain with unknown function. The orientation of the C-domain relative

to the A-domain varies depending the type and/or source of the amylase (Tiwari et al., 2015).

Figure 3: Structure of α-amylase (Paula and Pérola, 2010).

13

Domain A is shown in red, domain B in yellow and domain C in purple. At the catalytic center,

the calcium ion is shown as the blue sphere and the chloride ion as the yellow sphere. The green

structures are bound to the active site and to the surface binding sites.

1.5 α-Amylase active site structure and function

The active site of α-amylase is formed between domains A and B. Therefore, residues from domain

B are also involved in substrate binding (Paula and Pérola, 2010). The substrate binding sites are

commonly lined with aromatic residues (Phenylalanine, Tyrosine and Tryptophan) which make

hydrophobic stacking interactions with the sugar rings. In addition, the active sites contain many

residues which form hydrogen bonds with the substrate either directly or through water molecules

(Jackson et al., 2008; Tiwari et al., 2015).

In TAKA α-amylase (an α-amylase produced by Aspergillus oryzae), the first examined protein α-

amylase by X-ray crystallography, three acidic residues (one glutamic acid- Glu230 and two

aspartic acids- Asp206 and Asp297) were found at the center of the active site, and subsequent

mutational studies have shown that these residues are essential for catalysis (Tiwari et al., 2015).

Further studies have revealed that the glutamic acid residue is the proton donor, while the first of

the two conserved aspartic acids appearing in the amino acid sequence of the α-amylase family

member is thought to act as the nucleophile. The role of the second aspartic acid is less certain,

but it has been suggested to be involved in stabilizing the oxocarbenium ion-like transition state

and also in maintaining the glutamic acid in correct state of protonation for activity (Uitdehaag et

al., 1999; Tiwari et al., 2015). These residues occur near the ends of 3, 4, 5, and 7 strands of the

α/β-barrel and are found in four short sequences. They are known to be conserved in α-amylase

family enzymes (Tiwari et al., 2015).

1.6 Mode of action of α-amylase

Several models for α-amylase action pattern have been proposed, such as the random action and

the multiple attack action. Random action has also been described as either a single attack or a

multi-chain attack action (Tiwari et al., 2015). In single attack, the polymer molecule is

successively hydrolyzed completely before dissociation of the enzyme-substrate complex. While

in the later, only one bond is hydrolyzed per effective encounter. The second model, the multiple

attack action is an intermediate between the single-chain and the multi-chain action where the

14

enzyme cleaves several glycosidic bonds successively after the first (random) hydrolytic attack

before dissociation from the substrate. It is observed that the multiple attack action is generally an

accepted concept to explain the action pattern of amylases (Tiwari et al., 2015). However, most of

the endo-amylases displays a low to very low level of multiple attack action with temperature

increase depending on the amylase itself (Tiwari et al., 2015).

1.7 Reaction mechanism of α-amylase

Glutamic acid and aspartic acid are two catalytic residues present in the active site. The glutamic

acid acts as acid/base catalyst, supporting the stabilization of the transition states during hydrolysis

and the aspartic acid as nucleophile during the formation of the intermediate (Arpana et al., 2012).

As shown in Figure 4, once substrate is bound to the active site, proton transfer from the glutamic

acid to the glucosidic oxygen (the oxygen between two glucose molecules at the subsites -1 and

+1) takes place and the nucleophilic aspartate attacks the C1 of glucose at subsite -1 (Arpana et

al., 2012). These two events lead to disruption of glucosidic bond and an oxocarbonium ion-like

transition state converts to a covalent intermediate. The +1 subsite protonated glucose molecule

leaves the active site and is replaced by water or a new glucose molecule in the active site, which

then attacks the glycosyl-enzyme intermediate to form again an oxocarbonium ion-like transition

state (Arpana et al., 2012). Glutamate pulls off the proton from the water or newly entered glucose

molecule. The oxygen of the incoming water or newly entered glucose molecule forms a hydroxyl

group at C1 or a new glycosidic bond between -1 and +1 subsite.

Apart from the glutamic acid and aspartic acid residues described, another aspartic acid residue

has an indirect role in the catalytic process and thought to contribute its role in distortion of

substrate by binding to the OH (2) and OH (3) of the substrate (Uitdehaag et al., 1999). Apart from

these active site residues, conserved residues like histidine, arginine and tyrosine play an important

role in properly orienting the substrate into the active site. They also direct proper orientation of

the nucleophile and electronic polarization of substrate structure (Muralikrishna and Nirmala,

2005).

15

Figure 4: Structural organization of subsites of α-amylase active sites (Pascale et al., 2006). (○)

glucose unit; (↓) cut position; and (●) reduced glucose unit

Figure 5: The α-amylase reaction mechanism (Arpana et al., 2012).

1.8 Production of α-amylase

Two methods exist for production of enzymes and there are reports of α-amylase production with

these methods. These are submerged fermentation and solid state fermentation. The latter is a

relatively new method while the former is a traditional method of enzyme production from

microbes which has been in use for a longer period of time (Ajita and Thirupathihalli, 2014).

16

1.8.1 Submerged fermentation

Submerged fermentation involves the production of microorganisms in liquid medium containing

appropriate nutrients. In submerged fermentation, there is easy control of different parameters such

as pH, temperature, aeration and oxygen transfer. In addition, SmF supports the utilization of

genetically modified organisms to a greater extent than SSF and the sterilization of the medium

and purification process of the end products can be done easily (Paula and Pérola, 2010; Saranraj

and Stella, 2013; Ajita and Thirupathihalli, 2014). SmF system for enzyme production are

generally conducted in stirred reactors under aerobic conditions or fed batch systems (Paula and

Pérola, 2010). This fermentation technique is suitable for microorganisms that require high

moisture content for their growth. Fungi and yeast are known to grow under SSF as well as SmF

whereas bacteria have been considered unsuitable on SSF. Bacterial cultures are better managed

and manipulated in a submerged fermentation (Paula and Pérola, 2010).

1.8.2 Solid state fermentation

Solid state fermentation is defined as fermentation process occurring in the absence or near absence

of free liquid, employing an inert substrate or a natural substrate as a solid support (Noraziah et

al., 2017). In the solid state fermentation process, the solid substrate not only supplies the nutrients

to the culture, but also serves as an anchorage for the microbial cells. The moisture content of the

medium changes during fermentation as a result of evaporation and ultimately the moisture content

of the substrates becomes an important factor to consider (Saranraj and Stella, 2013). The solid

substrates commonly used in this method include; bran, bagasse, and paper pulp. The main

advantage is that nutrient-rich waste materials can be easily recycled and used as substrates in this

method (Noraziah et al., 2017). Negligible foaming problems during the process, reduction of

quantity of reagents used and waste water produced are other advantages that SSF has over SmF

(Ajita and Thirupathihalli, 2014).

Table 3: Bacterial sources and the mode of fermentation used for enzyme production (Ajita and

Thirupathihalli, 2014)

Bacterial source Method

B.amyloliquefaciens

Bacillus licheniformis

SSF

SSF

SSF

17

Bacillus coagulans

Chromohalobacter sp

B. polymyxa

B. mesentericus

B. vulgarus

B. megaterium

Bacillus licheniformis GCB U8

Bacillus sp. PS-7

Bacillus licheniformis M27

Halobacillus sp MA-2

Halomonas meridiana

Rhodothermus marinus

Bacillus cereus MTCC 1305

SSF

SSF

SSF

SSF

SSF

SmF

SSF

SSF

SmF

SmF

SmF

SSF SmF

SmF

.

Table 4: Fungal sources and the mode of fermentation used for enzyme production (Ajita and

Thirupathihalli, 2014)

Fungal source Method

Aspergillus oryzae SSF

Penicillium fellutanum SmF

Thermomyces lanuginosus SSF

Aspergillus niger SSF, Smf

Penicillium roquefortii SSF

Streptomyces rimosus SSF, Smf

Aspergillus kawachii SSF, Smf

Penicillium chrysogenumm SSF

Penicillium janthinellum (NCIM 4960) SSF

Aspergillus awamori SmF

Pycnoporus sanguineus SSF

1.9 Process parameters affecting α-amylase production

For maximal secretion of extracellular enzymes by microorganisms, optimization of some

parameters of their growth medium may be necessary. This is important as enzymes are required

in large quantities to meet industrial demand (Dey and Banerjee, 2015). Production of α-amylase

in fungi is known to depend on both morphological and metabolic state of the culture. Growth of

18

mycelium is crucial for extracellular enzymes like α-amylase (Ajita and Thirupathihalli, 2014; Dey

and Banerjee, 2015). Various factors have been known to affect the production of α-amylase such

as temperature, pH, and time of incubation, carbon sources, surfactants, nitrogen sources and

different metal ions. Moisture and agitation with regards to SSF and SmF, respectively can also

affect enzyme production. Interactions of these parameters are reported to have a significant

influence on the production of the enzyme. The optimum process control parameters vary

depending on the microbial source, desired end product, method of fermentation employed and

many other factors (Ajita and Thirupathihalli, 2014).

1.9.1 Temperature

The optimum temperature for production of enzymes normally depends on whether the source

organism is mesophilic or thermophilic (Lawal et al., 2014). Among the fungi, most α-amylase

production studies have been done with mesophilic fungi within the temperature range of 25–37

°C (Francis et al., 2003; Ramachandran et al., 2004). A raw starch degrading α-amylase was

produced by Aspergillus ficuum at 30 °C (Varalakshmi et al., 2013). Yeasts such as Saccharomyces

kluyveri and Saccharomyces cerevisiae were reported to produce α-amylase at 30 °C (Moller et

al., 2004). α-Amylase production at optimum level has been reported between 50–55°C for the

thermophilic fungal cultures such as Talaromyces emersonii, Thermomonospora fusca and

Thermomyces lanuginosus (Swetha et al., 2006). Bacterial amylases are produced at a much wider

range of temperature. Bacillus amyloliquefaciens, B. subtilis, B. licheniformis and B.

stearothermophilus are among the most commonly used Bacillus species reported to produce α-

amylase at temperatures 37–60 °C (Mendu et al., 2005; Swetha et al., 2006). Even with

thermophilic organisms, the α-amylase production increases with increase in temperature till it

reaches the optimum. With further increase in temperature the enzyme production decreases. This

may be due to the loss of moisture in the substrate which adversely affects the metabolic activities

of the microbes leading to reduced growth and decline in enzyme production (Ajita and

Thirupathihalli, 2014).

1.9.2 pH

Enzymes are pH sensitive and hence care must be taken to control the pH of the production process.

Earlier studies have revealed that fungi required slightly acidic pH and bacteria required neutral

19

pH for optimum growth (Sidkey et al., 2011). pH is known to affect the synthesis and secretion of

α-amylase just like its stability (Sethi et al., 2016). Aspergillus oryzae, A. ficuum and A. niger have

been found to give significant yields of α-amylase at pH 5.0 to 6.0 in SmF (Djekrif et al., 2005).

The production of extracellular α-amylase by the thermophilic fungus Thermomyces lanuginosus

was investigated in solid state fermentation (SSF). The maximum enzyme activity when wheat

bran was used as the substrate was recorded at pH condition of 6.0 (Bhargav et al., 2008). α-

Amylase producing yeast strains such as Saccharomyces cerevisiae and S. kluyveri exhibited

maximum enzyme production at pH 5.0 (Moller et al., 2004). Bacterial cultures such as B. subtilis,

B. licheniformis, and B. amyloliquefaciens required an initial pH of 7.0 (Haq et al., 2005;

Tanyildizi et al., 2005).

1.9.3 Carbon sources

Carbon sources such as galactose, glycogen and inulin have been reported as suitable substrates

for the production of α-amylases by Bacillus licheniformis (Swetha et al., 2006). Starch and

glycerol are known to increase enzyme production in B. subtilis (Tanyildizi et al., 2005). Soluble

starch has been described as the best substrate for the production of α-amylase by B.

stearothermophilus (Swetha et al., 2006). Aspergillus tamarii actively synthesizes α-Amylase

when cultured on maltose, starch and glycogen under static conditions. This yield is about four-

fold increase when compared to shaking cultures (Ajita and Thirupathihalli, 2014). Currently,

agricultural wastes are being used for both liquid and solid fermentation to reduce the cost of

fermentation media. This makes the process of enzyme production environmental friendly (Swetha

et al., 2006; Ajita and Thirupathihalli, 2014). These wastes consist of carbon and nitrogen sources

necessary for the growth and metabolism of organisms (Haq et al., 2005)

1.9.4 Nitrogen source

The nitrogen source used for production of α-Amylase may be organic or inorganic. Few of the

inorganic nitrogen sources include ammonium sulphate, ammonium chloride and ammonium

hydrogen phosphate. Most commonly used organic sources of nitrogen include peptone, yeast

extract and soyabean meal (Ajita and Thirupathihalli, 2014). Penicillium fellutanum and

Thermomyces lanuginosus showed maximum activity when peptone was used as the nitrogen

source (Erdal and Taskin, 2010; Ajita and Thirupathihalli, 2014). In Penicillium fellutanum, this

20

was about 9 % higher than the activity obtained with yeast, meat and casein (Erdal and Taskin,

2010). Peptone is usually the most high-yielding nitrogen source when employed in solid state

fermentation. Among the inorganic nitrogen sources, ammonium salts showed better results when

compared to sodium nitrate (Raul et al., 2014).

1.9.5 Duration of fermentation

This is a crucial factor in fermentation process. Normally, enzyme activity increases with increase

in incubation time till it reaches the optimum duration. In most cases, the production of enzyme

begins to decline if the incubation time is further increased. This could be due to the depletion of

nutrients in the medium or release of toxic substances (Lévêque et al., 2000; Raul et al., 2014).

1.9.6 Metal ions

Supplementation of salts of certain metal ions provides good growth of microorganisms and better

enzyme production whereas some will reduce enzyme production (Wang et al., 2018). Calcium

(Ca2+) ions are reported to be present in most α-amylases and studies have shown an increase in

enzyme production with addition of CaCl2 to the fermentation media (Francis et al., 2003). Strains

of Bacillus species grown with LiSO4 (20 mM) and MgSO4 (1 mM) showed increase α-amylase

production, but FeCl3 and MgSO4 exhibited negative influence on a-amylase production (Swetha

et al., 2006).

1.9.7 Moisture

Optimum levels of initial moisture content may vary depending on the microbial source used. For

fungal sources, the moisture content required is less compared to bacterial sources for high yield

of enzyme (Lévêque et al., 2000; Ajita and Thirupathihalli, 2014). The optimum level can be

known by determining the enzyme yield within a range of initial moisture content (Lévêque et al.,

2000). For solid state fermentation, low and high moisture levels of the substrate can affect the

growth of the microorganism and may result in variation in enzyme production. High moisture

content leads to reduction in substrate porosity, changes in the structure of substrate particles and

reduction of gas volume (Swetha et al., 2006). In a study of production of α-amylase by Penicillium

janthinellum using wheat bran as the substrate, the moisture content was varied within a range of

20 – 80 % by varying the amount of salt solution used in moistening the substrate particles. The

21

study revealed a maximum yield of enzyme at 60 % of initial moisture content. The yield decreased

at 80 % (w/w) moisture content (Prakasham et al., 2007). Generally, enzyme yield usually

increases with increase in initial moisture content until an optimal level, followed by decrease in

enzyme yield with further increase in moisture content. At low moisture content there is high water

tension and low solubility of the nutrients which causes the low yield of enzyme. With increase in

moisture content, the swelling of substrate takes place which ensures better uptake of nutrients by

the microbes. But with increase of moisture content further the enzyme yield decreases owing to

many reasons. The inter-particle distances reduce and can result in agglomeration of substrate

particles. Also the reduction in gas volume and gas diffusion results in impaired oxygen transfer

(Prakasham et al., 2007).

1.10 Isolation and purification of fungal α-amylase

Most fungi secrete α-amylase extracellularly for use in the digestion of carbohydrate in their

environment (Ajita and Thirupathihalli, 2014). Isolation of the extracellular enzyme after

fermentation is achieved by filtration and/or centrifugation process which removes the biomass

while the crude enzyme is left in the filtrate or supernatant. The crude enzyme obtained may be

subjected to various purification steps such as ammonium sulphate precipitation and gel filtration,

depending on the intended use (Renge et al., 2012).

1.10.1 Ammonium sulphate precipitation

Some proteins are soluble in water while others are not easily soluble. With little increase in salt

concentration, most proteins become soluble (Krisna and Sandra, 2014). Generally, positively and

negatively charged regions of proteins often self-aggregate. However, when salt is present, the

charges on the protein surface are neutralized by the salt’s anions and cations (Burges, 2009).

Further increase in the salt concentration makes the surface of the protein charged once again so

that the protein molecules begin to aggregate and precipitate out of solution (Krisna and Sandra,

2014). Ammonium sulphate is often used for the precipitation of proteins because of its low price,

high solubility which allows solution of high ionic strength, availability in pure state and ability to

stabilize protein structure (Markus and Aaron, 2007; Burges, 2009). The primary disadvantage of

salting out as a means of purifying enzyme is that contaminants often precipitate with the protein

of interest. Also, the enzyme precipitated is left in high concentration of the salt which can interfere

22

with the enzyme activity (Krisna and Sandra, 2014). Desalting is generally achieved by the method

of dialysis.

1.10.2 Gel filtration

Gel filtration chromatography, also known as size exclusion chromatography, is used to separate

molecules of different sizes. In addition, oligomeric forms of a particular protein can be resolved

as well as exchange of the buffer of a sample for a different one with this method (Krisna and

Sandra, 2014). Gel filtration column is comprised of a matrix of beads that contain sieves of a

particular size. These beads consist of cross-linked polyacrylamide, agarose, dextran, or a

combination of any of these (Scopes, 1993). Proteins and polymers may or may not enter the beads,

depending on their sizes. The large molecules will elute before small compounds such as ions and

buffer salts, which can enter the sieves in the matrix of the stationary phase (Ajita and

Thirupathihalli, 2014). Size exclusion columns are characterized by their cutoff size, void volume,

and column volume. Cutoff size refers to the approximate size of the largest molecule that can

possibly enter the beads. The void volume refers to the volume of the column that is not occupied

by the matrix while the column volume refers to the total accessible volume of the column to

solvent (Krisna and Sandra, 2014). Gel filtration chromatography serves as a final purification step

after applying at least one other purification step. It is not to be used as an initial protein

purification step after cell lysis because there will too many proteins with similar sizes (Krisna and

Sandra, 2014).

1.11 α-Amylase Assay

The enzyme is assayed spectrophotometrically (540 nm) by measuring the reducing sugars

released as a result of the action of α-amylase on starch. The amount of enzyme releasing one

μmole of the product (reducing sugars) per minute per ml gives the α-amylase activity unit (Ajita

and Thirupathihalli, 2014).

1.12 Biochemical properties of α-amylase

With increasing diversity of α-amylase application, there is always the need to search out for novel

α-amylase with unique properties such as optimal temperature for activity and stability and pH

profile (Swetha et al., 2006). The rate of hydrolysis of starch by α-amylase depends on many

process conditions in addition to temperature and pH such as the nature of substrate, substrate

23

concentration, enzyme concentration, presence of Ca2+ ions and other stabilizing agents. Hence,

α-amylases with characteristics suitable for the industrially relevant process conditions and

applications have to be appropriately selected as required (Saranraj and Stella, 2013).

1.12.1 Substrate specificity

The substrate specificity of α-amylase shows slight variation from one microorganism to another.

Generally, α-amylases display highest specificity towards starch followed by amylose,

amylopectin, cyclodextrin, glycogen and maltotriose (Saranraj and Stella, 2013).

1.12.2 Temperature optima and stability

Most enzymes are protein and denatures at high temperatures. The optimum temperature for α-

amylase activity ranges from 25 °C to 130 °C (Arpana et al., 2012). The lowest temperature

optimum of 25 to 30 °C has been reported for Fusarium oxysporum amylase and the highest of

100 and 130 °C from Pyrococcus furiosus and Pyrococcus woesei (Saranraj and Stella, 2013). In

most starch processing industries, it is important that α-amylase used is active at high temperature

of gelatinization (100-110 °C) and liquefaction (80-90 °C) to economize the process. Therefore,

there has been the need for more thermophilic and thermostable α-amylase (Silva et al., 2018).

However, regulatory authorities often require that no detectable enzymatic activity remain in the

product. It becomes important that the stability of an enzyme is studied before its application

(Iftikhar et al., 2013).

Thermostable α-amylases have been isolated from Bacillus amyloliquefaciens, B. licheniformis

and B. subtilis. The enzymes obtained from B. licheniformis generally were stable than those from

other Bacillus species (Fossi et al., 2005; Bozic et al., 2011). Most of the α-amylases have been

reported to contain one or more intrinsic Ca2+ ions present near the active center formed by the

two domains, A and B (Bozic et al., 2011). Studies have shown that the Ca2+ is necessary for

enzyme folding and enzyme stability. Secondary calcium binding sites have also been reported,

which enhanced the thermostability of α-amylase (Swetha et al., 2006).

1.12.3 pH optima and stability

The pH optima of α-amylases vary from 2 to 12 and stable over a wide range of pH from 4 to 11

(Saranraj and Stella, 2013). This wide range of pH shows its adaptability to different

24

environmental conditions. The pH optimum of α-amylase could have a temperature dependency

as observed with Bacillus stearothermophilus. In some cases, pH optimum shows a calcium

dependency as in the case of Bacillus stearothermophilus (Gupta et al., 2003).

1.12.4 Effect of metal ions

α-Amylases are calcium metalloenzymes and on incubation with chelating agent such as EDTA,

loses activity (Tripathi et al., 2007; Kumari et al., 2010). There has been report of loss of α-

amylase activity from soybean after dialysis against 5 mM EDTA. This enzyme activity was

restored by dialysis against buffer containing calcium (Kumari et al., 2010). Substitution of

calcium by other divalent metal ions did not regain enzyme activity. Some studies have been

carried out on the ability of other ions to replace Ca2+ in restoring α-amylase activity after EDTA

inactivation. And results so far have shown that Sr2+, Mg2+ and Ba2+ were able to give a positive

result in few cases (Saranraj and Stella, 2013). In the presence of Ca2+, amylases are much more

thermostable than without it. α-Amylase from Aspergillus oryzae shows unusual inactivation in

the presence of Ca2+, but retains activity after EDTA treatment. There are also reports where Ca2+

did not have any effect on the enzyme (Rani et al., 2003). Zn2+ has varied effect on different

amylases. Zn2+ was reported to inhibit thermostable α-amylase from a thermophilic Bacillus sp

(Burhan et al., 2003).

Table 5: Concentration effect of metal ions on the activity of α-amylases (Swetha et al., 2006)

Metal ion Concentration(mM)/ effect Organism

Ca2+

K+

Na+

Co2+

Mg2+

Ba2+

Mn2+

Zn2+

5 promoting

5 promoting

10 promoting

5 promoting

5 inhibitory

5 promoting

5 promoting

10 promoting

1 promoting

1 inhibitory

10 promoting

Lipomyces starkeyi

Bacillus sp. ANT-6

L. manihotivorans LMG 18010

Bacillus sp. L1711

Bacillus sp. ANT-6

Bacillus sp. L1711

Vibrio sp.

L. manihotivorans LMG 18010

Bacillus sp. I-3

Bacillus KSM-K38

L. manihotivorans LMG 18010

25

Fe2+

Cu2+

Ni2+

Al3+

Fe3+

Hg+

5 inhibitory

1 promoting

20 inhibitory

1 promoting

10 inhibitory

5 promoting

10 inhibitory

5 inhibitory

1 inhibitory

1 inhibitory

Bacillus sp. ANT-6

Bacillus sp. I-3

Cryptococcus flavus

Bacillus sp. I-3

L. manihotivorans LMG 18010

B. halodurans LBK 34

L. manihotivorans LMG 18010

B. halodurans LBK 34

Bacillus sp. I-3

Thermobifida fusca NTU22

1.12.5 Effect of inhibitors

The inhibitory effect of metal chelators indicates the requirement of certain metal ions for α-

amylase activity. Many metal cations, especially heavy metal ions, sulphydryl group reagents, N-

bromosuccinimide, hydroxyl mercuribenzoic acid, iodoacetate, BSA, EDTA and EGTA have

shown inhibitory effect on α-amylases (Saranraj and Stella, 2013).

1.12.6 Molecular weight

The molecular weights of microbial amylases are usually 50 to 60 kDa as shown directly by

analysis of cloned amylase genes and deduced amino acid sequences (Rani et al., 2003).

Glycosylation of enzyme molecule accounts for higher molecular mass of α-amylase (Robert et

al., 2005). However, the lowest value of 10 kDa was reported for Bacillus caldolyticus and the

highest of 210 kDa for Chloroflexus aurantiacus (Saranraj and Stella, 2013).

1.12.7 Stabilization of α-amylase

Enzymes are mostly proteins with a labile nature. Inactivating agents such as temperature, pH,

chemicals, etc. impair the native conformation of an enzyme, thus affecting its catalytic activity.

The utility of an enzyme depends mainly on its operational and storage stability (Fagain, 2003).

Stability of enzymes can be achieved by screening intrinsically stable enzymes, adding stabilizing

agents, chemical modification, immobilization, protein engineering (Silva et al., 2018). Calcium

ion (Ca2+), among other stabilizing additives has been widely used to attain thermostability (Burhan

et al., 2003). It has also been documented that in some α-amylases, sodium ion (Na+), potassium

26

ion (K+), ammonium ion (NH4+) and bovine serum albumin have protective effect on α-amylase

(Swetha et al., 2006).

1.13 Applications of α-amylase

For some years now, a shift from the acid hydrolysis of starch to the use of starch-converting

enzymes in the production of maltodextrin, modified starches, or glucose and fructose syrups has

occured. Currently, these enzymes (Amylases) comprise about 30 % of the world’s enzyme

production (Tiwari et al., 2015).

The global demand of α-amylase for a wide variety of industrial application is significant and

growing. α-Amylases have extensive applications in starch based food and non-food industries

(Rajendra et al., 2016). A few important industrial applications of α-amylase are given below.

1.13.1 Production of fructose and glucose syrup

This process of production of fructose and glucose syrups involves three steps: gelatinization,

liquefaction, and saccharification (Ajita and Thirupathihalli, 2014). Gelatinization involves the

dissolving of starch granules in water to form a viscous starch suspension. Liquefaction of starch

is its partial hydrolysis into short chain dextrins by α-amylase resulting in reduction of the viscosity

of the starch suspension. Saccharification is the production of glucose and fructose syrup by further

hydrolysis. This is carried out by glucoamylase which acts as an exo-amylase by cleaving the α-

1,4 glycosidic linkages from the non-reducing terminal (Ajita and Thirupathihalli, 2014). The

action of pullulanase (debranching enzyme) along with glucoamylase yields high glucose syrup.

This high glucose syrup can then be converted into high fructose syrup by isomerization catalysed

by glucose isomerase. The fructose syrup obtained is used as a sweetener, especially in the

beverage industry (Ajita and Thirupathihalli, 2014).

Initially, the hydrolysis process was done at a high temperature using an acid which was followed

by the enzyme employed saccharification process. This had some drawbacks because the acidic

nature of the first process meant a corrosion resistant material must be used and the high

temperatures inactivated thermolabile enzymes if the hot starch hydrolysate passes into subsequent

step. Today, thermostable amylases can be employed for hydrolysis at a high temperature. Bacillus

Stearothermophilus, Bacillus amyloliquefaciens, Bacillus licheniformis and Pyrococcus furiosus

27

are few of the many microbial sources used to produce α-amylase that is used in starch conversion

(Ajita and Thirupathihalli, 2014).

1.13.2 Bakery industry

Improvement of quantity, aroma, taste and porosity of the product are the reasons for the use of α-

amylase in bakery industries (Iftikhar et al., 2013). α-Amylase causes the starch to hydrolyze into

small dextrins which can further be fermented by yeast. This increases the rate of fermentation.

Also the starch hydrolysis decreases the viscosity of the dough, thus improving its texture and

increasing loaf volume by rising of dough (Ajita and Thirupathihalli, 2014). After baking, there

could be changes in of baked products during storage. All undesirable changes like increase of

crumb firmness, loss of crispness of the crust, decrease in moisture content of the crumb and loss

of bread flavor together are called staling. α-Amylase has an anti-staling effect and it has been

used to improve the shelf life and softness retention of baked goods, but it must be used with

caution as slight overdose may result in gumminess of the bread (Gupta et al., 2008), which is

caused due to production of branched dextrins (Chi et al., 2009). In such cases pullulanase is used

in combination with α-amylase resulting in specific hydrolysis of compound (dextrin) responsible

for the gummy nature of amylase treated bread (Ajita and Thirupathihalli, 2014).

1.13.3 Detergent industry

Detergent industries are the primary consumers of enzymes including α-amylase. The use of

enzymes in detergents formulations enhances the detergents ability to remove tough stains and

making the detergent environmentally safe (Paula and Pérola, 2010). α-Amylases are used in the

formulation of enzymatic detergent, and 90 % of all liquid detergents contain enzymes (Hmidet et

al., 2009). These enzymes are used in detergents for laundry and automatic dishwashing to degrade

the residues of starchy foods such as potatoes, gravies, custard, chocolate, and other smaller water

soluble oligosaccharides (Paula and Pérola, 2010). For application in detergent making, the

oxidative stability of amylases is one of the most important criteria for their use because the

washing environment is very oxidizing. This limitation could be overcome by using α-amylase

from genetically modified organisms instead of natural selection of better strains (Mukherjee et

al., 2009). Scientists from Novozymes and Genencore International, two major suppliers of

detergent enzymes, have replaced the oxidant sensitive amino acid residue methionine at position

197 by leucine in B. licheniformis amylase which resulted in an amylase with improved resistance

28

against oxidative compounds (Gupta et al., 2008). α-amylases used in the detergent industry are

derived from Bacillus species or Aspergillus species (Paula and Pérola, 2010).

1.13.4 Desizing of textiles

Modern production processes in the textile industry usually lead to breaking of the warp thread.

To strengthen the thread, sizing agents are used which strengthen the thread by forming a layer on

it which can be removed after the fabric is woven. Starch is a preferred sizing agent as it is easily

available, cheaper and can be easily removed from the fabric. The layer of starch is subjected to

hydrolysis in the desizing process where α-amylase is employed to cleave starch particles

randomly into water soluble components that can be removed by washing. The enzyme acts

specifically on the starch molecules alone, leaving the fibers unaffected (Gupta et al., 2008; Bozic

et al., 2011).

1.13.5 Paper industry

α-Amylase has been used for the manufacturing and modification of starches for coated paper. It

improves the paper quality, protects against mechanical injury and increases the stiffness and

strength in paper. The conversion of raw starch into glucose and fructose by the action of α-

amylase is prerequisite for sizing and coating of the paper. So, α-amylase is widely used for some

paper sizing (Bozic et al., 2011).

1.13.6 Feed industry

It has been reported that by the use of α-amylase in feed industry, the body weight gain and feed

conversion ratio of farm animals have increased. α-Amylase readily hydrolyzes the starch

polymers into fructose and glucose, which increases the digestibility of carbohydrates (Sidkey et

al., 2011).

1.13.7 Bioethanol production

Ethanol is the most utilized liquid biofuel. For the ethanol production, starch is the most used

substrate due to its low price and easily available raw material in most regions of the world (Chi

et al., 2009). The bioconversion of starch into ethanol involves liquefaction and saccharification,

where starch is converted into sugar using an amylolytic microorganism or enzymes such as α-

29

amylase, followed by fermentation, where sugar is converted into ethanol using an ethanol

fermenting microorganism such as yeast Saccharomyces cerevisiae (Ajita and Thirupathihalli,

2014). Recently, protoplast fusion between the amylolytic yeast Saccharomyces fibuligera and S.

cerevisiae was performed to obtain a new yeast strain that can directly produce the biofuel from

starch, eliminating the need for a saccharification step (Ajita and Thirupathihalli, 2014).

1.14 Aim and Objectives

1.14.1 Aim

This study is aimed at the production, purification and characterization of α-amylase obtained from

Aspergillus nidulans in submerged fermentation system for the purpose of establishing its

biotechnological relevance.

1.14.2 Specific Objectives of the Study

The objectives of this study were to:

➢ Isolate fungi species from soil collected from laundry waste water disposal site.

➢ Screen the isolated species for production of α-amylase.

➢ Carry out morphological and molecular characterization of the fungal isolate.

➢ Purify the α-amylase produced through ammonium sulphate precipitation, dialysis and gel

filtration (Sephadex G-100).

➢ Characterize the partially purified α-amylase.

➢ Carry out the pH and thermostability assessment of the enzyme.

30

CHAPTER TWO

MATERIALS AND METHODS

2.1 Materials

2.1.1 Reagents/Chemicals

Below is a list of the chemicals used in the study and their sources

Bovine serum albumin (BSA) Bio Rad Laboratories (India)

Copper II tetraoxosulphate VI (CuSO4) Merck (Germany)

Folin-Ciocalteau phenol reagent Sigma-Aldrich (USA)

Sodium dihydrogen phosphate (NaH2PO-4) Merck (Germany)

Sodium acetate Vickers Laboratories Limited, (London)

Sodium Hydroxide Merck (Germany)

Sodium potassium tartarate (Rochelle salt) Merck (Germany)

Sodium trioxocarbonate IV (Na2CO3) May and Baker Limited (England)

Tris-hydrochloric acid (Tris-HCl) Merck (Germany)

3, 5-dinitrosalicylic acid (DNSA) Sigma-Aldrich (USA)

All other chemicals used in this work were of analytical grade unless otherwise stated.

2.1.2 Apparatus

Glass wares: Pyrex, England

Centrifuge: Finland Nigeria 80-2B.

pH meter: Ecosan pH meter, Singapore.

Sensitive weighing balance: B2404-5 mettler Toledo, Switzerland.

Autoclave: Uday Burdon’s Patent Autoclave, India.

Water bath: Model DK. 23

Weighing balance: Ohaus Dial-O-Gram,Ohaus Cooperation, and

N.J. USA.

UV/Visible spectrophotometer: Jenway 6405, England.

31

Magnetic stirrer: AM-3250B Surgi Friend Medicals, England.

Milling machine: Thomas Willey laboratory Mill Model 4,

Anthor H (Thomas Company, Philadelphia,

USA).

2.1.3 Collection of plant material

The cassava tubers were obtained from Ogige main market, in Nsukka, Enugu State of Nigeria.

2.1.4 soil samples

The soil samples used for the experiment were collected from laundry waste water disposing site

at Odim, Nsukka, Enugu State. The organisms present there were thought to have unique properties

to enable them survive in the presence of those detergent materials. Therefore, the enzymes from

these organisms are most likely to retain activity in conditions applicable in industries.

2.2 Methods

2.2 1 Processing of cassava starch

Cassava starch was processed using the method described by Corbishley and Miller, (1984).

Freshly harvested cassava tubers were peeled, washed, and grated. The grated cassava (1000g)

was soaked in 4 L of distilled water for one hour after which it was sieved (3 times) with muslin

cloth. This was allowed to stand for 4 hours and the supernatant decanted. The isolated wet starch

was sun dried and packaged in plastic air tight container, labelled and kept in a cool, dry place.

This was used as carbon source for the fermentation process as well as substrate for the extracted

enzyme.

2.2.2 Isolation of α-amylase producing fungi

The following steps were taken to obtain an amylase producing fungi;

2.2.2.1 Serial dilution agar plate technique for isolation of fungi

Isolation of fungi from the soil was done by a modified serial dilution agar plate technique. Potato

dextrose agar (PDA) supplemented with streptopenicillin as antibacterial agent was used for the

isolation of soil molds. Ten grams (10 g) of soil (finely pulverized and air dried) was suspended

in 90 mL sterilized distilled water (blank no. 1) and shaken vigorously on a magnetic stirrer for 20

32

min to obtain uniform suspension of soil molds. Then 10 mL of suspension was transferred, while

in motion, from the stock suspension (no. 1), into 90mL sterile water blank (no. 2) with sterile

pipette under aseptic conditions to make 1:100 (10-2) dilution and shaken well for about five

minutes. Further dilutions from (10-3) to (10-7) were made by pipetting 10 mL suspension into

sterile water blanks number 3, 4, 5, 6 and 7 from water blank number 2, 3, 4, 5 and 6, respectively.

Finally, 1 mL aliquots of the suspension of final five dilutions i.e. (10-3) to (10-7) were added to

labeled and sterilized Petri plates. Approximately, 25 mL cooled molten (45 °C to 50 °C) PDA

supplemented with streptopenicillin, was added to each Petri plate and mixed gently by rotation.

After solidification of the medium, the inoculated Petri plates were incubated in an inverted

position at 25 °C for 3 to 7 days and observed for the appearance of mold colonies produced on

each plate of different dilutions. The purification of soil fungi was done by needle inoculation and

disc transfer methods on PDA plates. The inoculated PDA plates were incubated at 25 °C for 5

days and observed for purity and pure cultures were subcultured on PDA slants. The inoculated

slants were observed for the growth of pure cultures and maintained at 4 °C in a refrigerator.

2.2.2.2 Storage of pure fungal isolates

Pure fungal isolates were maintained on potato dextrose agar (PDA) slopes or slants as stock

cultures. Potato dextrose agar (Lab M Limited, UK) media were prepared according to the

manufacturer’s description by weighing 3.9g of PDA powder into small volume of distilled water

contained in a conical flask and made up to 100ml. The medium was autoclaved at 121 °C (15 psi)

for 15min. It was allowed to cool to about 45 °C and then poured into Petri dishes and allowed to

gel. The plates were then incubated in a B and T Trimline incubator at 37oC for 24hr to check for

sterility.

2.2.2.3 Examination of isolated fungi

Three days old pure cultures were examined under a light microscope (WESO) at X400

magnification. The colour, texture, nature of mycelia or spores and growth patterns were also

observed.

33

2.2.2.4 Fungal identification

This was carried out according to the method described by Martin et al. (2004). Three days old

pure cultures were 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 a light microscope at X400 magnification. Identification was

carried out by relating features and the micrographs to “Atlas of mycology” by Barnett and Hunter

(1972).

2.2.3 Molecular characterization of fungal isolates

2.2.3.1 DNA extraction from the microorganism

DNA was extracted from one week old cultures of pure fungal isolates as described by Murray

and Thompson (1980). Mycelium was scraped from the surface of cultures and transferred into

Eppendorf tubes followed by the addition of pre-warmed (60-65 °C) Cetyltrimethylammonium

bromide (CTAB) extraction buffer and then pulverized using a Fast Prep machine. Proteins were

removed from the supernatant with repeated phenol/chloroform extraction while nucleic acids

were precipitated from the aqueous layer using 96 % ice-cold iso-propanol (0.6 v/v) and 3 M

sodium acetate (0.1 v/v, pH 5.2). The resultant nucleic acid solution was digested with 0.01 mg/μL

RNAse to remove the RNA. Finally, DNA quantification will be measured and quantified using

NANODROP (ND-1000) spectrophotometer.

2.2.3.2 PCR amplification of the ITS region

The ITS region (ITS-1, 5.8S, and ITS-2) of the ribosomal RNA operon were amplified with primer

pairs ITS1 (5′-TCCGTAGGTGAACCTGCGG-3′) and ITS4 (5′GCTGCGTTCTTCATC-

GATGC-3′). To the hot start PCR premix tubes, 16 µl of Nuclease free water, 2 µl of Primer (1 µl

of forward primer and 1 µl of Reverse primer), and 2 µl of DNA extract were added and

microcentrifuge (Model LX-100) to mix for 30 sec. This was followed by loading on the thermo

cycler (PCR machine) and the PCR conditions preset. PCR products were analyzed by 2 % (w/v)

agarose gel electrophoresis and visualized by staining with ethidium bromide and UV illumination.

Product size was estimated by comparison with a 100 bp-ladder DNA marker.

34

2.2.3.3 DNA sequencing

The PCR products were purified using PCR purification kit and sequenced in both directions using

the same set of primers as for the PCR reactions.

2.2.3.4 BLAST analysis of DNA sequences

The ITS sequences obtained were compared with those in GenBank using the BLASTn (Basic

Local Alignment Search Tool for nucleotides) search algorithm. GenBank sequences that showed

high similarity and coverage (from 95 % upwards) with the sequences from the fungal isolates

were downloaded and included in this analysis for comparative purposes. DNA sequences were

aligned using Multiple Sequence Alignment based on Fast Fourier Transform (MAFFT) version 5

or any other suitable software (Katoh et al., 2005).

2.2.3.5 Phylogenetic analysis

Phylogenetic analysis of the ITS sequence data was conducted using Molecular Evolutionary

Genetics Analysis (MEGA) version 7 (Tamura et al., 2016).

2.2.4 Fermentation experiments

2.2.4.1 Preparation of fermentation broth

Submerged fermentation (SmF) technique was employed using a 250ml Erlenmeyer flask

containing 100ml of sterile cultivation medium optimized for amylase with 0.3 % (NH4)2SO3, 0.6

% KH2PO4, 0.1 % MgSO4.7H2O, 0.01 % FeSO4 and 1 % starch. The flask was stoppered with

aluminum foil and autoclaved at 121 oC (15 psi) for 15min.

2.2.4.2 Inoculation of broth

From the PDA slants, fresh plates were prepared as described above and inoculated. Three days

old cultures were used to inoculate the flasks. In every sterile flask, three discs of the respective

fungal isolates were added using a cork borer of diameter 10mm. The culture was incubated for

10 days at room temperature (25 °C).

35

2.2.4.3 Harvesting of fermented broth

At each day of harvest, flasks were selected from the respective groups and mycelia biomass

separated by filtration. Each day, the filtrate was analyzed for α-amylase activity and extracellular

protein concentration till the tenth day of fermentation.

2.2.4.4 Mass production of enzyme

After the ten-day pilot submerged fermentation (SmF) study, the day of peak amylase activity was

chosen for mass production of enzyme. A known number of 250ml Erlenmeyer flasks were used

to produce 2 L of the enzyme using the method described in sections 2.2.4.1 and 2.2.4.2 above.

Isolation was carried out on the peak day of enzyme activity.

2.2.5 Assays for α-amylase activity using dinitrosalicyclic acid (DNSA) reagent

α-Amylase activity was determined by the dinitrosalicyclic acid (DNSA) method as described by

Bernfeld, (1951). The amylase activity was assayed by incubating 0.5 ml of the enzyme with 0.5

ml of 1 % w/v starch dissolved in 50 mM sodium acetate buffer of pH 5.5, at 50 °C for 50 min.

One mil (1 ml) of 3, 5- dinitrosalicyclic acid (DNSA), was added to stop the reaction, followed by

boiling for 10 min. Sodium potassium tartarate (1 ml) was added to stabilize the colour and the

mixture was then allowed to cool. The same procedure used for the test solutions was used for the

blank except that the blank had no enzyme solution. The released reducing equivalent (maltose)

was monitored spectrophotometrically (using UV- VIS (JENWAY 6405)) at a wavelength of 540

nm against a blank. One unit of α-amylase activity was expressed as the amount of enzyme that

releases one micromole (μmol) of the reducing equivalent (maltose) per minute under assay

conditions. The enzyme activity was estimated by the amount of reducing equivalent (maltose)

released during the hydrolysis of the starch.

2.2.6 Effect of pH on α-amylase production

To determine optimal pH for α-amylase production, fungus cultures were cultivated in a 150 mL

flask containing 50 mL of medium optimized with different pH, ranging from 3.0 to 8.0. The pH

of the medium was adjusted by using 1 N HCl or 1 N NaOH. The flasks were kept in stationary

stage at 28 °C for 6 days of cultivation.

36

2.2.7 Determination of protein content

Protein content of the enzyme was determined by the method of Lowry et al. (1951), using bovine

serum albumin (BSA) as standard as outlined in Appendix 1

2.2.7.1 Procedure for protein determination

For protein standard curve, the reaction mixture contained 0.0 - 1.0 ml of protein stock solution

(2mg/ml BSA) in test tubes arranged in duplicates. The volume was made up to 1 ml with distilled

water. But for the test mixture, 0.5 ml of the enzyme was mixed with 0.5 ml of buffer. In either

case, 5 ml of solution E (solution E is described in appendix 1) was added to each tube and allowed

to stand at room temperature for 10 min. Then 0.5 ml of solution C (dilute Folin-Ciocalteau phenol

reagent) was added with rapid mixing. After standing for 30 min, absorbance was read at 750 nm

using UV spectrophotometer. Absorbance values were converted to protein concentrations by

extrapolation from the protein standard curve.

2.2.8 Purification of the enzyme

2.2.8.1 Determination of percentage ammonium sulphate saturation suitable for ɑ-amylase

precipitation

Nine test tubes were used to form an ammonium sulphate precipitation profile. ɑ-Amylase was

precipitated with gentle stirring at 20-100 % saturation of solid ammonium sulphate. The

ammonium sulphate-crude enzyme solutions were allowed to stand at 4oC for 30 hours till the

supernatant could be gently decanted off. The test tubes were centrifuged at 3500 revolutions per

minutes (rpm) for 10 minutes. Precipitates from the individual percentage ammonium sulphalte

saturations were redissolved in equal volumes of 50 mM sodium acetate buffer, pH 5.5. ɑ-Amylase

activity of the precipitates were assayed to determine the percentage ammonium sulphate

saturation that precipitated the enzyme with maximum activity.

2.2.8.2 Mass ammonium sulphate precipitation of ɑ-amylase

500 mL of crude enzyme were used in this process. From the studies in section above, forty percent

(40 %) ammonium sulphate saturation was found suitable for the precipitation of the enzyme.

Ammonium sulphate precipitation (at 40 % saturation) was carried out by dissolving gently 113 g

37

of the ammonium sulphate salt in 500 ml of the enzyme and stirring gently till the salt was

completely dissolved. The precipitates were redissolved in 100ml of 50 mM acetate buffer, pH 5.5

after centrifugation at 3500 rpm for 10 min and then kept under cold condition for further studies.

2.2.8.3 Dialysis

Dialysis bags preserved in 90 % ethanol were rinsed several times with distilled water and then

with buffer till they were free of the ethanol. Sodium acetate (0.01M and 0.05M) buffer pH 5.5

was used for enzyme dialysis. Dialysis was carried out for 12 hours with continuous stirring and

buffer changed firstly at 3 h interval for 0.01 M concentration and then 6 h interval for 0.05 M

concentration with a view to removing low molecular weight substances and other ions that may

interfere with enzyme activity. After, the dialyzed enzyme was also assayed for α-amylase activity

according to Bernfeld, (1951) and protein concentration by Lowry et al. (1951) while the

remaining sample was stored under freezing condition.

2.2.8.4 Gel filtration chromatography

A known weight of the powdered sephadex G-100 was swollen at room temperature for three days

and thereafter, made up to 120 ml volume, and packed into a column (75 by 2.0 cm) and

equilibrated with 50 mM sodium acetate buffer, pH 5.5, so as to equilibrate the pH of the packed

gel to that of the enzyme. A volume of 10ml of the dialysed enzyme was passed through gel

filtration to further purify the enzyme. The enzyme was applied to the column and eluted at a rate

of 5ml per 20 min. A volume of 5 ml of 50 fractions were collected and ɑ-amylase activity assayed

for in each fraction. Protein content was also monitored at 280nm, by using UV/Visible

spectrophotometer. The active fractions were pooled and stored under freezing condition.

2.2.9 Studies on purified enzyme

2.2.9.1 Effect of pH on α-amylase activity

The optimum pH for ɑ-amylase activity was determined using 50 mM sodium acetate buffer (pH

3.5 - 5.5), 50 mM phosphate buffer (pH 6.0 - 7.5) and 50 mM Tris-HCl buffer (pH 8.0 - 10.0), at

intervals of pH 0.5. In the assay, 0.5ml of enzyme was incubated with 0.5ml of 1 % starch in 50

mM of appropriate buffer for 50 min. ɑ-Amylase activity at these various pH was then monitored

as described earlier in section 2.2.5.

38

2.2.9.2 Effect of temperature on α-amylase activity

The optimum temperature was determined by incubating the enzyme with starch solution (1 %) at

30-90oC and the enzyme activity was assayed as described in section 2.2.5.

2.2.9.3 Effect of metal ions on α-amylase activity

A volume of metal ions (Ca2+, Mg2+, Mn2+, Fe2+, and Co2+) of varied concentrations (20, 30, 40,

50mM) were incubated with 0.5ml of enzyme solution and 0.5ml of starch solution (1%) at

optimum conditions of pH and temperature obtained in sections 2.2.9.1 and 2.2.9.2 above. The

reactions were carried out in triplicates and the enzyme activity was calculated as the average of

the independent repetitive sets of experiments.

2.2.9.4 Effect of substrate concentration on α-amylase activity

The effect of substrate concentration on the activity of amylase was determined by incubating the

enzyme with 10, 20, 30, 40, 50, 60, 70, 80, 90 and 100 mg/ml of starch solution using optimum

conditions of pH and temperature. The maximum velocity, Vmax and Michaelis constant, KM values

of the enzyme were determined using the Lineweaver-Burk plot of initial velocity data.

2.2.10 pH stability

The stability of the partially purified (dialyzed) enzyme was examined by measuring the residual

activity of the enzyme after being incubated for 2hours at pH ranging from 3.0 to 10.0 with 1.0

interval and at room temperature. The effect of pH on the activity of α-amylase was determined

by incubating 0.5ml enzyme solution (1:9 dilution) and different pH buffers, ranging from 3 -10

(1:1 dilution) over a period of two hours. At interval of 30mins, an aliquot was withdrawn and the

residual enzyme activity was then measured as described in the α-amylase assay section.

2.2.11 Thermal inactivation studies

The kinetic and thermodynamic parameters for the thermal inactivation of α-amylase from

Aspergilus nidullans were determined based on isothermal inactivation experiments for varying

period of time at varying temperatures (40º, 50º, 60º, 70º and 80º) in a temperature controlled water

bath. The activity of the partially purified (dialysis) enzyme solution was initially determined (1:9

dilution) without preheating which represents A0. Then the enzyme solution was placed in a pre-

warmed tube at the specified temperature (40, 50, 60, 70 and 80 °C), and aliquots were withdrawn

39

using a micropipette at 30 min time intervals for the incubation period of 120 min. Afterwards, the

samples were immediately cooled in ice water to stop the thermal inactivation process (so that

enzyme’s secondary and tertiary structures become stable after the heat treatment and any

reversible unfolding reverts back to the native forms (Griffin et al., 1984). The residual enzyme

activity was then measured as described in section 2.2.5 which represents At. The stability of the

enzyme was expressed as percentage residual enzyme activity. The incubation was carried out in

screw capped tubes to prevent change of volume of the sample and hence, the enzyme

concentration due to evaporation.

2.2.11.1 Determination of percentage residual activity

The % residual activities of the enzyme fractions were calculated using the formula below:

𝐴𝑡/𝐴0×100 %

2.2.11.2 Denaturation constants (kd) of the enzyme fraction

The kd of the ɑ-amylase preparations was estimated as follows. The calculated values of the %

residual activities were tabulated and kd was determined from the slope of plots of Iog % residual

activity at various temperatures against time (t) from the equations below:

𝐴𝑡/𝐴0= exp(−𝑘𝑑𝑡)

log(𝐴𝑡/𝐴0) = −𝑘𝑑𝑡 first other kinetics.

2.2.11.3 Half-life (t1/2) of the enzyme solution

The half-life (t1/2) was obtained from the equation,

t1/2 = ln 2/kd

2.2.11.4 Activation energy of denaturation Ea

The Ea was obtained from the Arrhenius equation:

𝐼𝑛𝑘d=𝐴𝑒−𝐸𝑎𝑅/𝑇.

where kd is the first order rate constant of thermal inactivation of the enzyme activity, obtained

through the slopes of the plots described above.

40

Espachs-Barroso et al. (2003), observed that in food processing, it is common to characterize first-

order reactions in terms of the thermal death concepts (D and Z-values). It is important to reduce

the number of deleterious enzymes in foods/food products to ensure proper safety. This is most

often done by thermal processing. Time-temperature measurements of residual enzyme activity

reduction are determined by thermal process parameters such as D- and Z- values.

2.2.12 Thermodynamics parameters

The change in enthalpy (ΔH0, KJmol-1), which is the amount of energy required to bring the

enzyme to the activated state for the subsequent denaturation at a given temperature, Gibbs free

energy (ΔG0, KJmol-1) of activation of the thermal denaturation and the entropy of activation of

the denaturation (ΔS0, KJmol-1 K-1) were calculated through the following equations,

Where Ea is the activation energy for denaturation, T is the corresponding absolute temperature

(K), R is the gas constant (8.314 Jmol-1K-1), Kh is the plancks constant (11.04 x 10-36 Jmin-1), Kb

is the Boltzmann constant (1.38x10-23 JK-1) and K (which is, kd) is the deactivation rate constant

(min-1) (Siddiqui et al., 1997; Riaz et al., 2007).

.

41

CHAPTER THREE

RESULTS

3.1 Microorganisms

3.1.1 Selection of amylolytic fungi

Three distinct isolates were obtained from the soil samples as shown in Figure 6. The isolates

showed positive result for amylase production.

3.1.2 Macroscopic and microscopic examination of fungal isolates

Two pure cultures; brown in colour, grayish near apex with smooth walled surface and bi-seriate

vesicle serration were identified as Aspergillus species. One of these two pure cultures displayed

irregular conidia surface and was suspected to be A. niger while the other with smoother surface

was suspected to be A. nidulans. The culture with bright conidial pigments was identified as

Trichoderma specie.

3.2 Production of α-amylase and study on crude

3.2.1 Effect of incubation period on α-amylase production

After a ten-day pilot study, the day of maximum α-amylase production from the isolated fungal

species, Aspergillus nidulans, Aspergillus niger and Trichoderma harzianum were day 6, 8 and 6,

respectively with activities of 124.125, 81.851 and 81.559 µmolmin-1 as shown in Figure 7. The

organism suspected to be Aspergillus nidulans which had highest activity was selected for

molecular characterization and used for further studies.

3.3 Molecular characterization of selected fungal isolate

The result of the selected organism’s DNA extraction and amplification is shown in Figure 8.

Comparing the electrophoretic detection of DNA amplicons on agarose gel with DNA markers

revealed that the ITS region of the organism is between 500 – 600 bp. The result of the DNA

sequencing is still expected.

42

3.2.3 Effect of medium pH on α-amylase production

As shown in Figure 9, α-Amylase production increased within slightly acidic pH medium with its

peak at pH 5.0 (140.707 μmol/min). Enzyme production reduced as pH increased from neutral to

the alkaline region.

3.2.4 Protein concentration of crude enzyme

The crude enzyme produced by Aspergillus nidulans had a protein concentration of 0.392mg/ml

as shown in Table 6.

3.2.5 α-Amylase activity of crude enzyme

As shown in Table 6, α-amylase activity of the crude enzyme was 145.6 μmol/min.

3.3 Purification of crude α-amylase

A decrease in the volume of enzyme obtained and total activity was observed with each

purification step. While a decrease in protein content was observed after ammonium sulphate

precipitation. The summary of the changes in activity and protein content associated with each

purification step is given in Table 6.

3.3.1 Ammonium sulphate precipitation profile of α-amylase

As shown in Figure 10, highest protein precipitation was observed at 40 % saturation with

Ammonium sulphate.

3.3.2 Gel Filtration of α-amylase

Figures 11 shows the elution profile for α-amylase harvested on day 6 equilibrated with 50 mM

sodium acetate buffer, pH 5.5. Fractions 13 - 25 showed α-amylase activity.

3.4 Characterization of α-amylase

3.4.1 Effect of pH on α-amylase activity

The optimum pH value for α-amylase was found to be 5.0 with an activity of 93.877 μmol/min, as

shown in Figure 12.

43

3.4.2 Effect of temperature on α-amylase activity

The optimum temperature for α-amylase was observed to be 60 °C, with activity of 136.734

μmolmin-1, after which a steady decline in activity was observed as shown in Figure 13.

3.4.3 Effect of metal ions on α-amylase activity

As shown in Figure 14, the activity of α-amylase was enhanced by the presence of Ca2+ and Mn2+

while Co2+, Fe2+ and Mg2+ had inhibitory effects on the enzyme.

3.4.4 Effect of substrate concentration on α-amylase activity

The study on the effect of substrate concentration revealed an increase in α-amylase activity with

increasing substrate concentration up to 100 mg/ml after which no further increase was observed.

3.4.5 Determination of kinetic parameters for substrate concentration

The Michealis constant, KM and maximum velocity, Vmax obtained for α-amylase from the

Lineweaver-Burk plot of initial velocity at different substrate concentrations were found to be

18.28 mg/ml and 144.927 µmol/min, respectively.

3.4.6 Effect of pH stability on α-amylase activity

The enzyme was observed to maintain more than 50 % of its activity for 120 min from pH 4.0 to

7.0. At pH 8.0 and 9.0, the enzyme maintained more than 50 % of its activity for 60 min but less

than 50 % for 120 min. At pH 3.0 and pH 10.0, the enzyme had 76 and 58 % decrease in activity,

respectively (Figure 17).

3.4.7 Thermal denaturation study of the enzyme

3.4.7.1 Percentage residual activities of the enzyme

The enzyme was observed to be thermally stable at temperature range of 40 °C – 50 °C for 120 min

with residual activity of 89 and 84 %, respectively. At 60 °C, 74 % activity was maintained for 60

min, which was followed by a decrease in residual activity. At 70 °C and 80 °C, the enzyme had

86 and 88 % decrease in activity respectively after 120 min of incubation (Figure 18).

3.4.7.2 Calculation of denaturation rate constant (kd) of the enzyme

As shown in Table 7, the denaturation rate constant (kd) increased with temperature increase.

44

3.4.7.3 Calculation of half-life (t1/2) of the enzyme

The half-life of the enzyme decreased as temperature increased (Table 7).

3.4.7.4 Calculation of D-values of the enzyme

The decimal reduction time (D-values) had a temperature dependent decrease as shown in Table

7.

3.4.7.5 Calculation of Z-values of the enzyme

The Z-value obtained from the thermal denaturation study of the enzyme was 31.75 °C.

3.4.7.6 Calculation of activation energy of inactivation of the enzyme

The activation energy for inactivation (Ea (inactivation)) of the enzyme was calculated to be

66.793 KJ/mol (Table 7).

3.5 Thermodynamics parameters of the enzyme

As shown in Table 7, enthalpy (ΔH0) was calculated and observed to decrease with increasing

temperature. Entropy (ΔS0) values obtained were positive and Gibb’s free energy (ΔG0) values

were also positive.

45

Figure 6: Pure cultures of A. niger, A. nidulans and Trichoderma harzianum

46

Figure 7: Effect of incubation period on α-amylase production by A.nidulans, A. niger and

Trichoderma sp

0

20

40

60

80

100

120

140

0 1 2 3 4 5 6 7 8 9 10

α-A

myla

se A

ctiv

ity

(µm

ol/

min

)

Day(s)

47

Figure 8: Agarose gel electrophoretogram of amplified ITS region organism of interest

(Aspergillus nidulans). The column labeled M shows the migration of 100 bp ladder DNA marker.

The column labeled 9 shows the position and migration of the organism of interest. The last column

is the control to which nothing was added. The absence of a band on this column signifies good

result

48

Figure 9: Effect of pH of medium on α-amylase production of Aspergillus nidulans

105

110

115

120

125

130

135

140

145

3 4 5 6 7 8

α-A

myla

se A

ctiv

ity

(µm

ol/

min

)

pH

49

Figure 10: Ammonium sulphate precipitation profile for α-amylase activity from A. nidulans

0

20

40

60

80

100

120

140

20 30 40 50 60 70 80 90 100

α-A

myla

se A

ctiv

ity

(µm

ol/

min

)

Ammomium sulphate saturation (%)

50

Figure 11: Gel filtration elution profile for α-amylase from A. nidulans

0

0.02

0.04

0.06

0.08

0.1

0.12

0.14

0.16

0.18

0

20

40

60

80

100

120

1 3 5 7 9 1113151719212325272931333537394143454749

Pro

tein

con

cen

trati

on

(mg/m

l)

α-A

myla

se A

ctiv

ity

(µm

ol/

min

)

Tube number

activity

protein

51

Table 6: The purification table of α-amylase from Aspergillus nidulans

Enzyme

Volume

(ml)

Protein

(mg/ml)

Total

protein

(mg/ml)

Activity

(µmol/min)

Total

activity

(U)

Specific

Activity

(U/mg)

Purification

fold

%

yield

Crude 1000 0.392 392 145.6

145600 371.43 1 100

(NH4)2SO4 500 0.075

37.5 43.88

21940 585.07 1.575 15.068

Dialysis

Gel

Filtration

30

10

0.178

0.0364

5.34

0.364

137.68

81.04

4130.4

810.4

773.48

2226.37

2.082

5.994

18.82

19.62

52

Figure 12: Effect of pH on purified α-amylase activity from Aspergillus nidulans

0

10

20

30

40

50

60

70

80

90

100

3.5 4 4.5 5 5.5 6 6.5 7 7.5 8 8.5 9 9.5 10

α-A

myla

se A

ctiv

ity (

µm

ol/

min

)

pH

53

Figure 13: Effect of temperature on purified α-amylase activity from Aspergillus nidulans

0

20

40

60

80

100

120

140

160

30 40 50 60 70 80 90

α-A

myla

se A

ctiv

ity

(µm

ol/

min

)

Temperature (0C)

54

Figure 14: Effect of some divalent metal ions on α-amylase activity from Aspergillus nidulans

0

10

20

30

40

50

60

70

80

CaCl2 CoCl2 MnCl2 MgCl2 FeCl2 control

α-A

myla

se A

ctiv

ity

(µm

ol/

min

)

20mM

30mM

40mM

50mM

55

Figure 15: Effect of substrate concentration on α-amylase activity from Aspergillus nidulans

0

5

10

15

20

25

30

35

40

45

50

0 20 40 60 80 100 120

α-A

myla

se A

ctiv

ity

(µm

ol/

min

)

Substrate concentration (mg/ml)

56

Figure 16: Lineweaver-Burk plot of α-amylase from A. nidulans

-0.03

-0.02

-0.01

0

0.01

0.02

0.03

0.04

0.05

0.06

0.07

-0.1 -0.05 0 0.05 0.1 0.15

1/a

ctiv

ity (

µm

ol/

min

)-1

1/S (mg/ml)-1

57

Figure 17: Percentage residual activity of α-amylase from A. nidulans at pH 3.0 to 10.0 for 120

min

0

20

40

60

80

100

120

0 30 60 90 120

% R

esid

ual A

ctiv

ity

Time (min)

pH 3

pH 4

pH 5

pH 6

pH 7

pH 8

pH 9

pH 10

58

Figure 18: Percentage residual activity of α-amylase from A. nidulans at pH 5.0 and temperatures

40 to 80 °C for 120 min

0

20

40

60

80

100

120

0 30 60 90 120

% R

esid

ual A

ctiv

ity

Time (min)

40°C

50°C

60°C

70°C

80°C

59

Figure 19: Log % Residual activity of α-amylase at temperature of 40 to 80٥C

0

0.5

1

1.5

2

2.5

0 30 60 90 120

log %

Res

idu

al A

ctiv

ity

Time (min)

40°C

50°C

60°C

70°C

80°C

60

Figure 20: A graph of log of D-value against temperature (°C) for the determination of the Z-

value

0

0.5

1

1.5

2

2.5

0 10 20 30 40 50 60 70 80 90

log D

Temperature (°C)

61

Figure 21: Arrhenius plot of ln kd against 1/T (K) for the determination of the Activation energy

of thermal deactivation (Ea)

-5

-4.5

-4

-3.5

-3

-2.5

-2

-1.5

-1

-0.5

0

0.0028 0.00285 0.0029 0.00295 0.003 0.00305 0.0031 0.00315 0.0032 0.00325

lnkd

1/T(K)

62

Table 7: Kinetic parameters for thermal inactivation of α-amylase

Temp

(°C)

kd

(min)-1

(t1/2)

(min)

D-value

(min)

40 0.0138 50.22806 166.854

50 0.0166 41.75585 138.7099

60 0.0819 8.463336 28.11459

70 0.1474 4.702491 15.62134

80 0.1735 3.995085 13.27138

Ea 66.793 KJ/mol

Z-value 31.75 (°C)

63

Table 8: Thermodynamic parameters for thermal inactivation of α-amylase

Enthalpy

(KJ/mol)

Free energy

(KJ/mol)

Entropy

(KJ/mol)

64.190718 54.51741323 0.035433351

64.107578 55.84754879 0.030256517

64.024438 53.24209743 0.039495753

63.941298 53.24952975 0.039163986

63.858158 54.40787421 0.034616424

64

CHAPTER FOUR

DISCUSSION

Alpha amylase (α-Amylase) is an enzyme with diverse applications and constitutes a class of

industrial enzymes of approximately 25 % of the world’s enzyme market (Rajendra et al., 2016).

α-Amylase has been applied in food industries, textile, paper, detergent, and pharmaceutical

industries (Saranraj and Stella, 2013). This has created a need for the production of very effective

and industrially stable, starch hydrolyzing α-amylase. Bacillus species and Aspergillus species

have been the most exploited microbial sources of α-amylase. Fungi species were chosen for this

work as a result of its GRAS (generally regarded as safe) status (Iftikhar et al., 2013) and tolerance

to acidity which reduces bacterial contamination (Rajendra et al., 2016). Fungi are also known for

their high secretory ability of industrially useful enzymes (Hans, 2014).

In this present study, three fungi isolate from the soil obtained from laundry waste water disposal

site were identified using their cultural characteristics such as the colony colour, texture, margins

and the presence of exudates. The taxonomic descriptions of Aspergillus niger, A. nidulans and

Trichoderma harzianum by Klich, (2002), matched the colony morphology of the isolates in this

study. Macroscopic and microscopic characteristics of fungi compared to Klich, (2002) taxonomic

description was also used by Zulkifli, and Zakaria, (2017) to identify fungi from corn grains and

Matome et al. (2017) for isolates from maize and soil. The findings of Abdul et al. (2015) who

morphological and molecular identified the fungus isolated from tropical bed bugs to be

Trichoderma harzianum by the description of Klich, (2002), emphasizes the need for such basic

identification approaches for the rapid screening of isolates in places where the availability of

advanced technologies for identification is still a challenge.

A ten-day submerged fermentation carried out on these organisms using starch as carbon source

showed that A. nidulans had maximum production (124.125 µmolmin-1) on the 6th day while A.

niger and Trichoderma harzianum maximally produced α-amylase on the 8th (81.851 µmolmin-1)

and 6th (81.559 µmolmin-1) day, respectively. The peak days were followed by a gradual decrease

till the tenth day. Therefore, A. nidulans with better enzyme production was chosen for further

studies. Behailu and Abebe, (2018) reported day 3 as the day of maximum extracellular α-amylase

production when 6 discs of 8mm size inoculum of culture of A. niger were used on submerged

65

fermentation, in contrast to the 3 discs of 8mm size inoculum of the three cultures used in this

study. Jiby et al. (2016) reported a maximal extracellular α-amylase secretion after 7 days of

submerged fermentation with 3 discs of 8mm size culture of A. niger. Inoculum size or

concentration was reported by Jiby et al. (2016) to affect the level of enzyme secretion. Other

factors such as pH, temperature, moisture content and nutrient type also affect the extracellular

secretion of enzyme and maximum day of enzyme production (Raul et al., 2014). The reduction

of α-amylase production beyond the day of maximum production may be due to depletion of

nutrient or build-up of toxic substances.

The result of molecular characterization of the organism of interest revealed an amplicon band of

the ITS region to be approximately 600 bp. This result shows that the DNA extraction and PCR

amplification process of the ITS region was correctly performed as fungal ITS region is

approximately 600 bp. Similar to the result of this study, Abdul et al. (2015) reported that the PCR

amplification of ITS region of their four fungal isolates obtained from bed bug, yielded a single

fragment of an approximately 500 and 600 bp. Although morphological identification of organisms

such as fungi is still relied upon heavily, it is now widely accepted that morphological and colony

appearance alone is insufficient to accurately identify the species. This is due to some of the

challenges of morphological method of identification such as inability of conidia to form in some

cases and the issue of confounding identification (Huzefa et al., 2017).

The pH optimization study carried out on production media of Aspergillus nidulans showed a peak

activity of 140.706 µmol/min at pH 5.0, followed by that of pH 6.0 (138.739 µmol/min). At neutral

pH and alkaline pH, the activity of the enzyme was observed to decrease. Similarly, Murali et al.

(2011) reported peak activity of α-amylase from A. niger at pH 5.0 using submerged fermentation.

pH optimum of 4.0 was recorded for α-amylase production on submerged fermentation using an

Aspergillus specie (Ominyi, 2013). The result of the pH optimization study suggests that A.

nidulans can be applied in processes that allow whole cell utilization such as in bioethanol

production with Saccharomyces cerevisiae that share similar pH optimum for growth.

Amalkadhimghadban, (2016) reported a maximum enzyme production with a media of pH 7.0 by

Baccillus Licheniformis. This differences in optimal pH for α-amylase production by Aspergillus

and Bacillus species are in accordance with the report by Jiby et al. (2016) that most Aspergillus

efficiently produce enzymes under slightly acidic pH while bacteria secretes enzymes more in

66

neutral and slightly alkaline pH. This could be because of the ability of Aspergillus to produce

efficiently, organic acids such as citric acid and kojic acid that are harnessed for use in food

industry and itaconic acid that is used in the polymer industry (Polizeli et al., 2016).

40 % ammonium sulphate saturation was found suitable to precipitate protein with the highest

enzyme activity. Lawal et al. (2014) reported 60 % ammonium sulphate saturation for α-amylase

from Aspergillus niger. Sidkey et al. (2011) and Varalakshmi et al. (2013) reported that 40-60 %

saturation gave the highest enzyme activity of α-amylase isolated from A. flavus and A. oryzae,

respectively. These results are in line with that obtained from the present study. Generally, proteins

become more soluble in aqueous solution when salts such as ammonium sulphate is added. Low

salt concentration stabilizes the various charged groups on a protein molecule, thus enhancing the

solubility of proteins in aqueous solution (Krisna and Sandra, 2014). This phenomenon is known

as salting in. However, with increasing salt concentration, the water molecules become more

attracted to the salt allowing protein-protein interaction which leads to aggregation and subsequent

precipitation or salting out. Therefore, the result of the present study in which 40 % ammonium

sulphate saturation gave the highest activity suggest that α-amylase from A. nidulans would be

relatively hydrophobic (having more hydrophobic amino acid residues).

There was an increase in specific activity after ammonium sulphate precipitation and dialysis

which suggests that more of the proteins of interest were being purified. The further increase in

enzyme activity after dialysis shows that ammonium sulphate salts which may interfere with the

enzyme activity were removed.

The gel filtration carried out on dialyzed enzyme using sephadex G 100 showed increased specific

activity (2226.37 U/mg) with a purification fold of 5.99. This observation indicates that the

purification processes chosen were ideal for α-amylase purification.

The purified enzyme was characterized based on pH, temperature, and changes in substrate

concentration on the activity. From the study of the effect of pH on enzyme activity, as pH

increased from 3.5 to 5.0, α-amylase activity was observed to increase. Further pH increase beyond

pH 5.0 reduced the activity of the enzyme. Therefore, optimal pH was found to be 5.0. This

observation followed the statement of Hans, (2014) that enzyme activity under the influence of

varying pH will increase from acidic region up to an optimal point, beyond which activity

67

decreases. This behavior of enzymes in varying pH conditions is attributed to the state of

protonation of the functional groups of amino acid that participate in catalysis. Protonation and

deprotonation of functional groups could have a positive or a negative effect on catalysis. At

extreme pH regions, the 3-D structure of the enzyme may be irreversibly denatured. The

observation of activity decrease beyond pH 5.0 may be due to preference of A. nidulans to lower

pH for its growth and metabolism. Also, many amino acids in a protein molecule carry a charge

because they contain basic and acidic groups that are usually situated on their surface when in

aqueous solution. Changes in pH will affect these charges according to their acid dissociation

constants and bring about a change in the 3-D structure (denaturation) of the enzyme which causes

reduction or loss of activity (Hans, 2014). Changes in pH may as well affect the shape or charge

properties of the substrate so that either the substrate cannot bind to the active site or it cannot

undergo catalysis. Wang et al. (2018) in their examination of eight α-amylase gene family (denoted

as Amy A, C, D, E, F, G, H and M) from Aspergillus niger, cloned and expressed in Pichia pastoria

noted an optimal pH of 4.5 for Amy A, C, E and F, whereas, Amy G and M displayed maximal

activity at pH 5.0. Amy D and H had optimum pH of 6.0. This observation is comparable with the

result obtained in the present study. The temperature study followed a similar pattern with an

increase in enzyme activity as temperature was raised from 30 °C to 60 °C. Further increase in

temperature beyond 60 °C reduced the activity of the enzyme, making 60 °C the optimum. This

indicates that α-amylase from A. nidulans would be suitable for a wide range of industrial

processes such as in detergent and food industries. Wang et al. (2018) recorded that AmyA, AmyD

and AmyF exhibited maximal activity levels at 30 °C. The optimum temperature for AmyC,

AmyE, AmyH and AmyM was 40 °C, whereas AmyG showed optimum temperature at 60 °C.

Dey and Banerjee, (2015) reported an optimal pH and temperature of 5.5 and 50 °C, respectively

for α-amylase from A. oryzae in a liquid cultivation media. This shows that α-amylase from A.

nidulans can compare favorably with those from mostly used sources, A. niger and A. oryzae in

the industries processes.

The present study investigated the effect of some divalent metal ions (Co2+, Cu2+, Mg2+, Fe2+, Ca2+

and Mn2+) on the α-amylase activity from A. nidulans. The observed result showed a calcium

dependent increase in the α-amylase activity from A. nidulans. This suggests that the α-amylase

from A. nidulans is a Ca2+ dependent type. Ca2+ enables intramolecular cross linking of disulfide

bridge (Wang et al., 2018) and this favors the stable conformation of the enzyme. Of the eight

68

recombinant α-amylases studied by Wang et al., (2018), six α-amylases (AmyA, C, E, F, G, and

H) showed slight Ca2+ independent increase in the activity of the enzymes. In this study, Mn2+ also

was observed to increase the activity of α-amylase from A. nidulans. This is consistent with the

results of Almeida et al. (2017) and Wang et al. (2018). Wang et al. (2018) recorded increase in

activity of most α-amylase in the presence of Mn2+. This suggests that Mn2+ may have a stabilizing

effect on the groups on the active site of the enzyme. Other metal ions Co2+, Cu2+, Mg2+ and Fe2+

showed inhibitory effect on α-amylase activity from A. nidulans. This result again is similar to that

observed by Wang et al. (2018). From their result, Co2+, Cu2+, Mg2+ and Fe2+ inhibited the eight

recombinant α-amylase from A. niger. However, Mukherjee et al. (2018) observed inhibitory

effect on α-amylase activity from A. niger in presence of Ca2+, Mg2+, Zn2+, Mn2+, Cu2+, Fe3+ at 5

mM concentration. Studies have shown that the effect of divalent metal ions on the activity of α-

amylase can vary with microbial sources and even among close relatives (Dey and Banerjee, 2015).

This could be attributed to the existence of isoforms. Though isozymes catalyze the same reaction,

they differ in amino acid sequence which brings about differential display in some properties.

On the effect of substrate concentration, the Lineweaver-Burk plot was used to calculate the

Michaelis constant, KM and the maximum velocity, Vmax values. The Vmax and KM were calculated

to be 144.927 μmol/min and 18.28 mg/ml, respectively. Mukherjee et al. (2018) reported a KM and

Vmax values of 1.4mg/ml and 0.992 μmol/min, respectively. Sheoran and Dhankhar, (2016) in their

study on A. flavus observed a KM and Vmax values of 1.43 mg/ml and 250 mol/min/mg, respectively.

The KM of an enzyme reveals the concentration of substrate required to achieve half maximal

activity of the enzyme. The KM value of an enzyme also shows the strength of binding of the

substrate to the enzyme. Substrates that bind more tightly to the enzyme active site will have a low

KM whereas, loosely bound substrates will have a high KM. On the other hand, maximal rate, Vmax

(indicator of enzyme efficiency) shows the number of substrate molecules catalyzed per second.

Enzymes are protein molecules by nature and therefore are made up of folded polypeptide chains.

Its primary, secondary and tertiary structures are determined by the order of their amino acid. The

functional or native state of an enzyme (tertiary structure) is based on molecular geometry and

intramolecular chemical interactions of groups informed by the amino acid sequence (Silva et al.,

2018). In the presence of a denaturing agent (pH, temperature, metal salts) and upon storage,

denaturation of enzyme sets in. Denaturation refers to the unfolding of the 3-D structure of an

69

enzyme to a disordered polypeptide state. Renaturation may be achieved if the denaturation agent

is relieved (Hans, 2014). As enzymes are required to function in conditions different from their

physiological conditions in industrial processes, their stability becomes crucial to increase their

economic value and industrial application.

The pH stability studies showed that the enzyme was most stable at pH 5.0, retaining 99 % and 90

% of its activity after 60 and 120 min, respectively. This result is consistent with the assertion of

Hans, (2014) who stated that an enzyme is fairly stable at its own optimal pH and such pH is best

for its testing (assaying), storage and application. The enzyme was found to be stable between the

pH range of 4.0 to 6.0, with about 10 % or less loss of activity after 60 min. Mahmood et al. (2018)

reported a similar stable pH of 4.0 to 6.0 for α-amylase from A. niger. This result suggests that α-

amylase from A. nidulans can be applied in food industry, especially in baking because most

doughs have pH values between 5.0 and 6.0. α-Amylase application in baking industry generates

sugar for yeast fermentation, improve moisture retention, decrease staling and have a crumb-

softening effect on the final product. The increased sugar helps to improve flavor and enhance

crust colour (Ajita and Thirupathihalli, 2014). At pH 7.0 the α-amylase percentage residual

activities after 60 min and 120 min of incubation were 79 % and 72 %, respectively. The enzyme

still retained more than 50 % of its activity after 60 min of pre-incubation at pH 8.0 and 9.0. At

pH 3.0 and 10.0, the enzyme had 76 % and 58 % decrease in activity and a further 90 % and 88 %

decrease, respectively after 120 min. Although the optimum pH for α-amylase in this study was

5.0, it exhibited relative stability within the pH range of 4.0 to 9.0. This result is similar to the

report of Sethi et al. (2016) on α-amylase from A. terreus with maximum activity observed at pH

5.0 but fairly stable from pH 3.0 to 10.0. This result suggests that the enzyme produced could be

applied in a wide range of industrial processes that operate within the observed stable pH.

High temperatures distorts the functional configuration of enzymes leading to their inactivation.

Obtaining a thermostable enzyme continues to be a major challenge for enzyme application in

some industrial processes. Therefore, in the present study, the thermal stability of α-amylase from

A. nidulans was investigated for the purpose of finding its suitable industrial application. The

enzyme was observed to be thermally stable at temperature range of 40 °C – 50 °C maintaining

about 96 % and 90 % of initial activity for 60 min and about 89 % and 84 % of initial activity after

120 min. Mahmood et al. (2018) reported a maximal residual activity of 96 % at 40 °C with the

70

crude α-amylase from A. niger after 70 min of incubation. Working on recombinant α-amylases

from A. niger, Wang et al. (2018) reported that AmyA, D, F and AmyM were stable below 40 °C

whereas AmyC and AmyE retained more than 60 % of activity after heat treatment between 40 to

60 °C for 60 min.

In the present work, at 60 °C (the optimal temperature), α-amylase maintained 74 % activity for

60 min, followed by a decrease in residual activity. This result suggests that α-amylase from A.

nidulans could be applied in detergent making industry that operates at about 60 °C and would

reduce energy consumption in industries that operates at temperature lower than 60 °C. At

temperature beyond 60 °C the enzyme was observed to be readily denatured. According to Frédéric

et al. (2016), there was 100% retention of initial activity at 80 °C by α-amylase produced from

Lactobacillus fermentum after 30 min of incubation. In baked products, α-amylase used usually

continue to be minimally active which retards staling, but there is the possibility of the crumb

becoming gummy if the enzyme activity is appreciable. Therefore, the result obtained in this study

suggest that α-amylase from A. nidulans could be a better alternative to the more thermostable

enzyme from bacterial in baking industry.

Several kinetic indicators (inactivation rate constant (kd), half-life (t1/2) and decimal reduction time

(D-value)), and thermodynamic indicators (activation energy of denaturation Ea, enthalpy of

denaturation ΔH0, free energy of denaturation ΔG0 and entropy associated with thermal

denaturation ΔS0 have been used to investigate enzyme thermostability (Kouamé et al. 2017). In

the present study, the inactivation rate constant kd increased with temperature increase and half-

life (t1/2) decreased with increasing temperature, indicating more thermal denaturation of the

enzyme with increase in temperature. At the optimum temperature of 60 °C, a half-life of 8.46 min

was observed. This is similar to the value of 8.5 min half-life observed with ɑ-amylase from

sorghum bicolor at 60 °C (Kumar, 2008). The D-value was equally found to decrease with

temperature increase. D-value is the time at a particular temperature for the initial enzyme activity

to decrease by 90 %. Therefore, a high D-value indicates that the enzyme is thermally stable. The

Z-value which is the temperature increase needed to further reduce the D-value by ten folds was

calculated to be 31.75 °C. A high Z-value is an indicator of enzyme thermal stability (Kouamé et

al., 2017). This result suggests that relatively low temperature increase will be needed to inactivate

the enzyme if applied in food industry. The activation energy for denaturation (Ea) was calculated

71

to be 66.79 Kjmol-1. Comparing free and immobilized α-amylase from A. awamori, Karam et al.

(2017) found the immobilized enzyme to have Ea value of 16.7 Kjmol-1, approximately two folds

the value of the free enzyme 8.71 Kjmol-1. High Ea value means higher energy requirement for

denaturation. At the optimum temperature of 60 °C, the enthalpy of denaturation was observed to

be 64.024 Kjmol-1. Karam et al. (2017) reported enthalpy (ΔH0) values of 14.0Kjmol-1 and

6.02Kjmol-1 at 50 °C for the immobilized and free α-amylase from A. awamori. Also, a high ΔH0

value is an indication of relatively high energy requirement for denaturation. In this study, ΔH0

was found to decrease with temperature increase which means that less energy is required to

denature the enzyme at high temperature. The positive value of ΔH0 shows that the denaturation

reaction is endothermic Kouamé et al. (2017). Another thermodynamic parameter that gives

insight to the thermostability of an enzyme is the free energy of denaturation (ΔG0). A negative

ΔG0 value or a low positive value would reveal spontaneity in the thermal denaturation of the

enzyme (Karam et al., 2017). In this study, 53.25 Kjmol-1 was observed as the free energy of

denaturation (ΔG0) at the optimum temperature of 60 °C. Karam et al. (2017) in their study,

observed a higher free energy of denaturation value with immobilized α-amylase from A. awamori

compared to the free enzyme with values of 94.96 and 87.96 Kjmol-1. Entropy associated with

thermal denaturation (ΔS0) will normally increase with denaturation of the enzyme molecule. A

relatively low ΔS0 value indicates a more ordered or stable state while a high or positive value

reveals increased randomness or disorder. In the present work, ΔS0 was found to be positive which

suggests increase in disorder with increasing temperature. At the temperature optimum (60 °C),

ΔS0 was observed to be 0.039 Kjmol-1. Again, comparing immobilized and free α-amylase at their

optimum temperature (50 °C), Karam et al. (2017) recorded ΔS0 values of -253.7 and -249.7

Kjolmol-1, respectively. The result of the present study suggests that if applied in an industrial

process that require inactivation of the enzyme in the final product, low amount of energy will be

invested to achieve that compared to a more ordered enzyme (one with very low entropy value) at

high temperature.

4.1 Conclusion

Results obtained from this study have shown that A. nidulans is a good source for the production

of α-amylase for biotechnological applications. This is reflected by its hypersecretory ability

relative to A. niger and Trichoderma harzianum. The pH and temperature optimum of 5.0 and 60

°C together with the evidence of a wide range of pH and temperature stability shows feasibility for

72

application of this enzyme in detergent industry, textile desizing and paper industry. The thermal

sensitivity of the enzyme beyond 60 °C as revealed by the thermodynamic study is ideal for its

application in food industry where thermal inactivation of the enzyme is essential in the final

product. α-Amylase from A. nidulans being calcium dependent raises hopes of obtaining an

improved stability in the presence of this metal ion for application at temperature higher than that

observed in this study.

4.2 Suggestions for further studies

To further improve the stability and catalytic properties of α-amylase from A. nidulans;

1. Further research should be carried out on the effect of metal ions on the thermostability of the

enzyme.

2. Immobilization on the enzyme should also be investigated.

3. Since enzymes are not naturally designed for industrial but physiological functions, genetic

manipulation of the organism, A. nidulans could be used to produce α-amylase with desired

specificity for a particular industrial application.

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APPENDIX

1.0 PREPARATION OF REAGENTS

1.1 Preparation of Buffers

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The buffers used in this study were of pH 5.0. These buffers were used to assay for enzyme activity

and used during the enzyme purification. The working buffers were prepared by this procedure.

Sodium acetate buffer of 50 mM and Phosphate buffer of 50 mM were prepared by dissolving

1.640g sodium acetate salt and 2.84g of disodium hydrogen phosphate salt in separate 1000ml of

distilled water and stirred with a magnetic stirrer till a homogenous solution was formed. The

solutions were titrated against acetic acid and sodium di-hydrogen phosphate, respectively till the

required pH was obtained.

1.2 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 tartarate, 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

distilled water in a ratio of 1:1 on the day of use.

Solution D: Standard protein (Bovine Serum Albumin, BSA) solution/ Enzyme

Solution E: Freshly prepared alkaline solution was made by mixing 50ml of solutions A and 1ml

of solution B.

1.3 Preparation of 2mg/ml Bovine Serum Albumin (BSA) Standard Protein

An amount of 0.2g of BSA was dissolved in 100ml of distilled water and then used as a protein

stock solution. Ten tubes, labelled A-J, were used. Each of the tubes was in triplicates.

1.4Preparation of Dinitrosalicyclic Acid (DNS) Reagent

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A modification of DNS reagent was used in the assay. The reagent contains 2.5g dinitrosalicyclic

acid, 1.25g sodium sulphite, and 3.2g sodium hydroxide.

1.5Preparation of Rochelle’s Salt

This reagent contains 75g of sodium potassium tartarate in 125ml of distilled water.

1.6 Preparation of 5mM Glucose Standard

0.9 g of D- Glucose (molecular weight 180 g/mole) was dissolved in 1000ml of distilled water a

homogenous stock solution was obtained. From the stock, it was serially diluted as shown below.

Ten test tubes were used and labelled from A-J. Each test tube was in triplicate, after the addition

of the Rochelle’s salt, the mixtures were read spectrophotometrically at 540 nm.

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Figure 22: Protein Standard Curve

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1

Ab

sorb

an

ce a

t 750n

m

Protein concentration in mg/ml

86

Figure 23: Glucose Standard Curve

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

0 1 2 3 4 5 6

Ab

sorb

ance

at

54

0n

m

Concentration (mM)

5mM Glucose standard curve