1

EXTRACTION, PARTIAL PURIFICATION AND

CHARACTERIZATION OF CELLULASE FROM Aspergillus

fumigatus AND Aspergillus flavus IN SUBMERGED

FERMENTATION SYSTEM USING BREADFRUIT HULLS

Digitally Signed by: Content manager’s Name

DN : CN = Webmaster’s name

O = University of Nigeria, Nsukka

OU = Innovation Centre

Agboeze Irene E.

OSUAGWU, UCHECHUKWU O.

(PG/MSc/12/61930)

BIOLOGICAL SCIENCES

BIOCHEMISTRY

2

TITLE PAGE

EXTRACTION, PARTIAL PURIFICATION AND CHARACTERIZATION OF

CELLULASE FROM Aspergillus fumigatus AND Aspergillus flavus IN SUBMERGED

FERMENTATION SYSTEM USING BREADFRUIT HULLS AS CARBON SOURCE

A DISSERTATION SUBMITTED IN PARTIAL FULFILLMENT OF THE

REQUIREMENT FOR THE AWARD OF THE DEGREE OF MASTER OF SCIENCE

(M.Sc) IN ENZYMOLOGY AND PROTEIN CHEMISTRY, UNIVERSITY OF

NIGERIA, NSUKKA

BY

OSUAGWU, UCHECHUKWU O.

(PG/MSc/12/61930)

DEPARTMENT OF BIOCHEMISTRY

UNIVERSITY OF NIGERIA

NSUKKA

DECEMBER, 2014

3

CERTIFICATION

Osuagwu, Uchechukwu O., a postgraduate student with registration number

PG/M.Sc/12/61930 in the Department of Biochemistry has satisfactorily completed the

requirements for the research work for the degree of Master of Science (M.Sc.) in

Biochemistry. The work embodied in this report 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 DR S. O. O. EZE

(Supervisor) (Supervisor)

______________________________

PROF OFC NWODO

(Head of Department)

EXTERNAL EXAMINER

4

DEDICATION

This project is dedicated to my family members- Mr and Mrs Osuagwu, Nonye Osuagwu,

Uloma Ofole and Obioma Osuagwu.

5

ACKNOWLEDGEMENT

I wish to express my gratitude to my supervisors- Prof. F. C. Chilaka and Dr. S. O. O. Eze for

their guidance and support during the course of this work. I have benefitted immensely under

their tutelage both at undergraduate and postgraduate levels.

I am also thankful to the Department of Biochemistry for providing me with the necessary

equipment which were used to carry out this work. I am equally indebted to the Lecturers in

the Department who at some point indicated interest in the work and encouraged me to carry

on. They include: Prof. OFC Nwodo, Prof. L. U. S. Ezeanyika, Prof. P. N. Uzoegwu, Prof.

I.N.E. Onwurah, Prof. O.U. Njoku, Dr. V. N. Ogugua, Prof. E.O. Alumanah, Prof. H. A.

Onwubiko, Dr. O.C. Enechi, Dr. C. A. Anosike, Dr. C. S. Ubani, Dr. V. E. O. Ozougwu, Mr.

P. A. C. Egbuna, Mr. O. Ikwuagwu, Dr. O. U. Njoku and Mr. C. C. Okonkwo. I am

especially grateful to Dr. P. E. Joshua for his guidance and input to the success of this project

work. I also appreciate the external examiner for taking out his time to examine this project

I am grateful to my friends and colleagues who contributed in one way or the other to make

this a successful venture- Cliff Victor, Onos Iruoghene, Christopher Ugwu, Angela

Igboanugo, Onyedika Aruma and many other members of my class. I also thank Arinze Linus

for the necessary support and encouragement throughout the duration of work.

Finally, I owe immense gratitude to my parents- Mr & Mrs C. O. Osuagwu, and to my

siblings- Nonye, Uloma and Obioma for their patience and kindness during the duration of

the work. God bless you all.

6

ABSTRACT

Cellulase is a complex enzyme system commercially produced by filamentous fungi under

solid state or submerged cultivation. It has wide applications in textile, food and beverage

industry for effective saccharification process. In this study, Treculia africana (breadfruit)

hulls were used to induce cellulase production from Aspergillus flavus and Aspergillus

fumigatus grown under submerged fermentation conditions. Crude cellulase was harvested

after 5 days of growth with activities of 2.97 and 3.87 U/ml for enzymes produced by A.

flavus and A. fumigatus respectively. The protein concentrations for the crude enzymes were

found to be 4.03 and 4.17 mg/ml for A. flavus and A. fumigatus respectively. The enzymes

were then subjected to a two step purification process of ammonium sulphate precipitation

and gel filtration. Gel elution fractions were assayed for total cellulase activity. Two

prominent peaks indicating isoforms were observed. The isoforms were designated A and B

for enzymes of A. flavus and C and D for enzymes of A. fumigatus. These 4 fractions were

characterised separately. The pH optima of enzymes of A. flavus were 6.5 and 7.0

corresponding to the isoforms A and B with activities 3.31 and 3.53 U/ml respectively.

Enzymes from A. fumigatus had optimum pH of 5.0 for both isoforms C and D with

corresponding activities of 3.07 and 3.42 U/ml. The temperature optimum of enzymes of A.

flavus was 50⁰C with peak activities of 2.72 and 2.58 U/ml while enzymes of A. fumigatus

had maximum activities of 3.01 and 3.14 U/ml at a temperature of 55⁰C for both isoforms.

The Michaelis-menten constants Km, were 59.02, 47.67, 27.82 and 32 mg for isoforms A, B,

C and D respectively. Also, the maximum velocity, Vmax were 142.9, 166.7, 128.21 and

90.91 µmol/min for the isoforms A, B, C and D respectively. Using cellobiose as substrate,

Vmax values were 588.2, 476.2, 833.3 and 666.67 µmol/min for isoforms A, B, C and D

respectively. Km values of 7.7, 3.3, 11.1 and 9.1 mM were obtained for isoforms A, B, C and

D respectively.

7

TABLE OF CONTENT

Title Page - - - - - - - - - - i

Certification- - - - - - - - - - ii

Dedication - - - - - - - - - - iii

Acknowledgement- - - - - - - - - - iv

Abstract- - - - - - - - - - - v

Table of Content- - - - - - - - - - vi

List of Figures - - - - - - - - - - vii

List of Tables - - - - - - - - - viii

CHAPTER ONE: INTRODUCTION AND LITERATURE REVIEW

1.1 Cellulose - - - - - - - - - - 2

1.1.1 Structure of cellulose - - - - - - - - 2

1.1.1.1 Chemical structure - - - - - - - - 2

1.1.1.2 Crystalline structure - - - - - - - - 4

1.1.2 Cellulose biosynthesis - - - - - - - - 6

1.1.3 Sources of cellulose - - - - - - - - 6

1.2 Breadfruit (Treculia africana)- - - - - - - 7

1.3 Cellulose hydrolysis - - - - - - - - 8

.1.3.1 Acid hydrolysis - - - - - - - - - 8

1.3.2 Enzyme hydrolysis - - - - - - - - 8

1.4 Cellulases - - - - - - - - - - 9

1.4.1 Cellulase classification - - - - - - - 10

1.4.2 Types of cellulases - - - - - - - - 11

1.4.3 Endoglucanase (EC 3.2.1.4) - - - - - - - 11

1.4.4 Exoglucanase (EC 3.2.1.91) - - - - - - - 11

1.4.5 β-glucosidase (EC 3.2.1.21) - - - - - - - 11

1.5 Mechanism of action of cellulases - - - - - - 12

1.6 Molecular biology of cellulase - - - - - - - 13

1.7 Cellulase production - - - - - - - - 15

1.7.1 Cellulase producing microoganisms - - - - - - 15

1.7.1.2 Mesophilic microorganisms - - - - - - - 19

1.7.1.3 Thermophilic microorganisms - - - - - - 22

8

1.7.2 Aspergillus spp - - - - - - - - - 24

1.8 Fermentation methods - - - - - - - - 24

1.8.1 Solid-State fermentation (SSF) - - - - - - - 24

1.8.2 Submerged fermentation (SmF)/ Liquid fermentation (LF) - - - 24

1.9 Factors affecting cellulase enzyme production- - - - - - 25

1.9.1 Chemical Factors - - - - - - - - 25

1.9.1.1 Effect of carbon sources - - - - - - - 25

1.9.1.2 Effect of nitrogen sources - - - - - - - 27

1.9.1.3 Phosphorus sources - - - - - - - - 27

1.9.2 Physical factors - - - - - - - - - 27

1.9.2.1 pH - - - - - - - - - - 27

1.9.2.2 Temperature - - - - - - - - - 27

1.10 Applications of cellulases - - - - - - - 28

1.10.1 Cellulases in brewing and wine biotechnology - - - - 28

1.10.1.1 Beer brewing process - - - - - - - 28

1.10.1.2 Wine production - - - - - - - 29

1.10.2 Cellulases in pulp and paper biotechnology - - - - - 29

1.10.2.1 Biomechanical pulping - - - - - - - 30

1.10.2.2 Biodeinking - - - - - - - - - 30

1.10.3 Cellulases in textile and laundry biotechnology - - - - 31

1.10.3.1 Biostoning and biopolishing - - - - - - - 31

1.10.3.2 Laundry - - - - - - - - - 32

1.11 Aim of study and objectives - - - - - - - 33

1.11.1 Aim of study - - - - - - - - - 33

1.11.2 Specific objectives of the study - - - - - - 33

CHAPTER TWO: MATERIALS AND METHODS

2.1 Materials - - - - - - - - - 34

2.1.1 Reagents - - - - - - - - - - 34

2.1.2 Apparatus - - - - - - - - - 34

2.2 Method - - - - - - - - - - 35

2.2.1 Collection of breadfruit - - - - - - - - 35

2.2.2 Collection of microorganism - - - - - - - 35

9

2.2.3 Preparation of ground breadfruit hulls - - - - - - 35

2.2.4 Storage of pure fungal isolates - - - - - - - 36

2.2.5 The Fermentation broth - - - - - - - - 36

2.2.6 Inoculation of the broth - - - - - - - - 36

2.2.7 Harvesting of the fermented broth - - - - - - 37

2.2.8 Mass production of enzyme - - - - - - - 37

2.2.9 Procedure for protein determination - - - - - - 37

2.2.10 Cellulase assay - - - - - - - - 38

2.2.10.1 Glucosidase (Cellobiase) assay - - - - - - 38

2.2.10.2 Endoglucanase (CMCase) assay - - - - - - 38

2.2.10.3 Total cellulase (Filterpaperase) assay - - - - - 39

2.2.11 Partial purification of protein - - - - - - - 39

2.2.11.1 Determination of percentage ammonium

sulphate saturation suitable for cellulase precipitation - - - 39

2.2.11.2 Ammonium sulphate precipitation of cellulase - - - - 40

2.2.11.3 Gel filtration - - - - - - - - - 40

2.2.12 Studies on partially purified enzyme - - - - - - 40

2.2.12.1 Enzyme progress curve - - - - - - - 40

2.2.12.2 Effect of pH change on total cellulase activity - - - - 40

2.2.12.3 Effect of temperature change on total cellulase activity - - - 41

2.2.12.4 Determination of kinetic parameters - - - - - - 41

2.2.12.5 Further studies with partially purified enzyme - - - - 41

CHAPTER THREE: RESULTS

3.1 Incubation period (pilot studies) - - - - - - - 42

3.2 Studies on crude enzyme - - - - - - - - 43

3.2.1. Protein concentration of crude enzyme- - - - - - 43

3.2.2 Cellulase activity of crude enzyme - - - - - - 44

3.3 Ammonium sulphate precipitation profile of cellulases- - - - - 45

3.4 Gel filtration (elution profile of cellulase enzymes) - - - - 46

3.5 Summary of purification steps - - - - - - - 48

3.6 Changes in protein concentration of partially purified enzymes- - - 50

3.6.1 Changes in protein concentration of enzymes from A. flavus. - - - 50

10

3.6.2. Changes in protein concentration of enzymes from A. fumigates - - 51

3.7 Changes in total cellulase activity of partially purified enzymes - - - 52

3.7.1 Changes in total cellulase activity of

partially purified enzymes from A. flavus - - - - - 52

3.7.2 Changes in total cellulase Activity of

partially purified enzymes from A. fumigatus -- - - - - 53

3.8 Changes in specific activities of partially purified enzymes - - - 54

3.8.1 Changes in specific activities of partially purified enzymes from A. flavus - 54

3.8.2 Specific activities of partially purified enzymes from A. Fumigatus - - 55

3.9 Enzyme characterization - - - - - - - - 56

3.9.1 Enzyme progress curve - - - - - - - - 56

3.9.1.1 Enzyme progress curve of partially purified enzymes from A. flavus - 56

3.9.1.2 Enzyme progress curve of

partially purified enzymes from A. fumigatus - - - - 57

3.9.2 Effect of pH change on cellulase activity - - - - - 58

3.9.2.1 Effect of pH change on cellulase produced by A. flavus - - - 58

3.9.2.2 Effect of pH change on cellulase produced by A. fumigatus - - - 59

3.9.3 Effect of temperature change on cellulase activity - - - 60

3.9.3.1 Effect of temperature on cellulase produced by A. flavus - - - - 60

3.9.3.2 Effect of temperature on cellulase produced by A. fumigatus - - - 61

3.9.4 Determination of kinetic parameters (using filter paper as substrate) - - 62

3.9.4.1 Determination of kinetic parameters for enzymes of A. flavus - - - 62

3.9.4.2 Determination of kinetic parameters for enzyme of A. fumigates - - 64

3.9.5 Determination of kinetic parameters (using cellobiose as substrate) - - 66

3.9.5.1 Determination of kinetic parameters for enzymes of A. flavus - - 66

3.9.5.2 Determination of kinetic parameters for enzymes of A. fumigatus - - 68

CHAPTER FOUR: DISCUSSION

4.1 Discussion - - - - - - - - - 71

4.2 Conclusion - - - - - - - - - 76

4.3 Suggestions for further studies - - - - - - - 76

REFERENCES - - - - - - - - - 77

APPENDICES - - - - - - - - - 90

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LIST OF FIGURES

Fig. 1: Cellulose structure - - - - - - - - 4

Fig. 2: Crystalline forms of Cellulose I- - - - - - - 5

Fig. 3: Cellulose biosynthesis- - - - - - - - 7

Fig. 4: Breadfruit seed hulls - - - - - - - - 8

Fig. 5: Mechanism of cellulolysis - - - - - - - 13

Fig. 6: 3D structure of cellulase E4 from T. fusca- - - - - - 14

Fig. 7: A. flavus - - - - - - - - - 23

Fig. 8: A. fumigatus - - - - - - - - - 23

Fig. 9: Effect of incubation period on cellulase production - - - - 42

Fig. 10: Protein concentration of crude enzyme produced by the microoganisms- - 43

Fig. 11: Total cellulase, glucanase and cellobiase activities of crude enzyme - 44

Fig. 12: Ammonium sulphate precipitation profiles for celluluases

isolated from A. flavus and A. fumigatus.- - - - - - 45

Fig. 13a: Gel elution profile of proteins produced by A. flavus.- - - - 46

Fig. 13b: Gel elution profile of proteins produced by A. fumigatus - - - 47

Fig. 14: Changes in protein concentration after

partial purification of cellulases from A. flavus - - - - 50

Fig. 15: Changes in protein concentration after

partial purification of cellulases from A. fumigatus - - - - 51

Fig. 16: Changes in total cellulase activity after

partial purification of enzymes from A. flavus - - - - 52

Fig. 17: Changes in total cellulase activity

after partial purification of enzymes from A. fumigatus- - - - 53

Fig. 18: Changes in specific activity after partial

purification of cellulases from A. flavus.- - - - - - 54

Fig. 19: Changes in specific activity after partial

purification of cellulases from A. fumigatus- - - - - - 55

Fig. 20: Progress curve of cellulases isolated from A. flavus - - - - 56

Fig. 21: Progress curve of cellulases isolated from A. fumigatus - - - 57

Fig. 22: Effect of pH on cellulases produced by A. flavus- - - - - 58

Fig. 23: Effect of pH on cellulases produced by A. fumigatus- - - - 59

12

Fig. 24: Effect of temperature on cellulases produced by A. flavus - - - 60

Fig. 25: Effect of temperature on cellulases produced by A. fumigatus - - 61

Fig. 26: Lineweaver-Burke plot of cellulase A

from A. flavus using filter paper as substrate- - - - - 62

Fig. 27: Lineweaver-Burke plot of cellulase B

from A. flavus using filter paper as substrate-- - - - - 63

Fig. 28: Lineweaver-Burke plot of cellulase C

from A. fumigatus using filter paper as substrate- - - - - 64

Fig. 29: Lineweaver-Burke plot of cellulase D

from A. fumigatus using filter paper as substrate- - - - - 65

Fig. 30: Lineweaver-Burke plot of cellulase A

from A. flavus using cellobiose as substrate- - - - - - 66

Fig. 31: Lineweaver-Burke plot of cellulase B

from A. flavus cellobiose as substrate - - - - - - 67

Fig. 32: Lineweaver-Burke plot of cellulase C

from A. fumigatus cellobiose as substrate - - - - - 68

Fig. 33: Lineweaver-Burke plot of cellulase D

from A. fumigatus cellobiose as substrate - - - - - 69

LIST OF TABLES

Table 1: Lignocellulose composition of several agricultural wastes - - - 7

Table 2: Some mesophilic cellulolytic Bacteria- - - - - - 15

Table 3: Some mesophylic cellulolytic fungi - - - - - - 18

Table 4: Some (hyper) thermophilic cellulolytic Bacteria and Archaea - - 20

Table 5: Some (hyper) thermophilic cellulolytic Fungi- - - - - 21

Table 6: Summary of purification steps of cellulase from A. flavus- - - - 48

Table 7: Summary of purification steps of cellulase from A. fumigatus - - 49

Table 8: Cellulase characterization table- - - - - - - 70

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CHAPTER ONE

INTRODUCTION

Enzymes are the catalysts of biological processes. Like any other catalyst, an enzyme brings

the reaction catalyzed to its equilibrium position more quickly than would occur otherwise.

An enzyme cannot bring about a reaction with an unfavourable change in free energy unless

that reaction can be coupled to one whose free energy change is more favourable (Nelson and

Cox, 2000). The activities of enzymes have been recognized for thousands of years.

However, only recently have the properties of enzymes been understood properly (Wolfgang,

2007). Indeed, research on enzymes has now entered a new phase with the fusion of ideas

from protein chemistry, molecular biophysics, and molecular biology which have given rise

to applications in fields ranging from agriculture to industry (Wolfgang, 2007).

The enzyme industry as we know it today is the result of a rapid development seen primarily

over the past four decades and thanks to the evolution of modern biotechnology (Ole et al.,

2002). Enzymes found in nature have been used since ancient times in the production of food

products, such as cheese, sourdough, beer, wine and vinegar, and in the manufacture of

commodities such as leather, indigo and linen (Ole et al., 2002). All of these processes relied

on either enzymes produced by spontaneously growing microorganisms or enzymes present

in added preparations such as calves’ rumen or papaya fruit. The development of

fermentation processes during the later part of the last century, aimed specifically at the

production of enzymes by use of selected production strains, made it possible to manufacture

enzymes as purified, well-characterized preparations even on a large scale (Wolfgang, 2007)

Microbial cellulases have shown their potential application in various industries including

pulp and paper, textile, laundry, biofuel production, food and feed industry, brewing and

agriculture. Due to the complexity of the enzyme system and immense industrial potential,

cellulases have been a potential candidate for research by both academic and industrial

research groups (Shang, 2013). The growing concerns about depletion of crude oil and the

emissions of greenhouse gases have motivated the production of bioethanol from

lignocellulose, especially through enzymatic hydrolysis of lignocellulosic materials (Bayer et

al., 2004; Himmel et al., 1999)

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1.1 Cellulose

Cellulose is a linear polymer of β-D-glucose units linked through 1,4-β-linkages with a

degree of polymerization ranging from 2,000 to 25,000 (Kuhad et al., 1997). Cellulose chains

form numerous intra- and intermolecular hydrogen bonds, which account for the formation of

rigid, insoluble, crystalline microfibrils (Golan, 2011). Natural cellulose compounds are

structurally heterogeneous and have both amorphous and highly ordered crystalline regions

(Morana et al., 2011). The degree of crystallinity depends on the source of the cellulose and

the highly crystalline regions are more resistant to enzymatic hydrolysis (Morana et al.,

2011). Cellulosic materials are particularly attractive because of their relatively low cost and

abundant supply. As the most abundant polysaccharide in nature, cellulose decomposition

plays not only a key role in the carbon cycle of nature, but also provides a great potential for

a number of applications, most notably biofuel and chemical production (Lynd et al., 2012).

The central technological impediment to more widespread utilization of this important

resource is the general absence of low-cost technology for overcoming the recalcitrance of

cellulosic biomass.

1.1.1 Structure of Cellulose

1.1.1.1 Chemical Structure

Payen first used the term cellulose for this plant constituent which is the most widespread

organic compound on Earth (Payen 1938; Guo et al., 2008]). The total amount of this

polysaccharide on our planet has been estimated at 7 × 1011 tons (Coughlan, 1985) and

constitutes the most abundant and renewable polymer resource available today. Cellulose is

an insoluble crystalline substrate, flavourless, odourless, hydrophilic, insoluble in water and

in most organic solvents, chiral, and with a wide chemical variability (Coughlan, 1985). It is

a structural component of the cell wall of green plants accounting for almost 33% of the total

biomass. It is also biosynthesized in other living systems such as Bacteria and Algae.

Cellulose produced by plants usually exists within a matrix of other polymers primarily

hemicellulose, lignin, pectin and other substances, forming the so-called lignocellulosic

biomass, while microbial cellulose is quite pure, has a higher water content, and consists of

long chains (Jagtap and Rao, 2005) . It is a carbohydrate polymer with formula (C6H10O5)n ,

15

consisting of a linear chain of several hundred to over ten thousand 1,4-β-D-glucose units

linked through acetal functions between the equatorial -OH group of C4 and the C1 carbon

atom (Jagtap and Rao, 2005). The high stability of this conformation leads to a decreased

flexibility of the polymer, so this is usually described as a real tape.

There are two different types of intra- and one interchain hydrogen bonds in the structure, and

it has been considered that the intrachain hydrogen bonds determine the single-

chainconformation and the stiffness of cellulose, while the interchain hydrogen bond is

responsible for the sheetlike nature of cellulose (Watanabe and Tokuda, 2001; Klemm et al.,

2002; Klemm et al., 2005). The chains are arranged parallel to each other and form

elementary fibrils that have a diameter between 1.5 and 3.5 nm (microfibrils), the length of

the microfibrils is about of several hundred nm (Watanabe and Tokuda, 2001; Klemm et al.,

2002; Klemm et al., 2005) .

16

Fig. 1: Structure of Cellulose (Matheus et al., 2013)

1.1.1.2 Crystalline structure

The high degree of hydrogen bonds within and between cellulose chains can form a 3-D

lattice-like structure, while amorphous cellulose lacks this high degree of hydrogen bonds

and the structure is less ordered (Morana et al., 2011). The physical and chemical properties

of cellulose are defined by intermolecular interactions, cross-linking reactions, polymer

lengths, and distribution of functional groups on the repeating units and along the polymer

chains (Morana et al., 2011).

Initially, crystalline structure of native cellulose (cellulose I) has been studied by X-ray

diffraction and has been defined as monoclinic unit cells with two cellulose chains with a

twofold screw axis in a parallel orientation forming slight crystalline microfibrils (Gardner

and Blackwell, 2004; Klemm et al., 2005). Moreover, there are other types of crystal

17

structures: cellulose II, III, and IV (Gardner and Blackwell, 2004); the cellulose I, result the

less stable thermodynamically, while the cellulose II is the most stable structure (Klemm et

al., 2005). The cellulose I can turn into other forms using different treatments; for example,

by mercerization, using aqueous sodium hydroxide or dissolution followed by precipitation

and regeneration (formation of fiber and film) (O'Sullivan, 1997; Nishiyama et al., 2002).

However, additional information on the structure of noncrystalline random cellulose chain

segments are needed because it is very important for the accessibility and reactivity of the

polymer and the characteristics of cellulose fibers (Paakkari et al., 1989).

Fig. 2: Crystalline forms of Cellulose I (Matheus et al., 2013)

1.1.2 Biosynthesis of Cellulose

Cellulose is synthesized by a variety of living organisms, including plants, algae, bacteria,

and animals. It is the major component of plant cell walls with secondary cell walls having a

much higher content. The biosynthesis of cellulose essentially proceeds by the

polymerization of glucose residues using an activated substrate UDP-glucose (Saxena et al.,

200).

In plants, cellulose is synthesized on the plasma membrane by the enzyme cellulose synthase

that is present in the membrane. In the bacterium Acetobacter xylinum, the enzyme cellulose

synthase is present on the cytoplasmic membrane, and the cellulose is obtained

18

extracellularly. However, in other organisms, cellulose is found to be synthesized in other

regions of the cell. In the alga Pleurochrysis, cellulose scales are formed in the Golgi

apparatus and then deposited on the cell surface (Saxena et al, 2000).

The biosynthesis of Cellulose proceeds in at least two stages – polymerization and

crystallization. The first stage is catalyzed by the enzyme cellulose synthase, and the second

stage is dependent on the organization of the cellulose synthases possibly with other proteins

such that the glucan chains are assembled in a crystalline form (Saxena et al, 2000).

Fig. 3: Biosynthesis of Cellulose (Peter, 2008)

1.1.3 Sources of Cellulose

The plant cell wall is the major source of cellulose. Cellulose therefore abounds in

agricultural wastes of plant origin.

19

Table 1: Lignocellulose composition of several agricultural wastes

Lignocellulosic materials Cellulose (%) Hemicellulose(%) Lignin (%)

Hardwood 40-55 24-40 18-25

Softwood 45-50 25-35 25-35

Nut shell 25-30 25-30 30-40

Chestnut shell 27.4 10 44.6

Grape stalk 38 15 33

Corn stover 36.7 13.33 33

Wheat straw 30 50 15

Rice straw 32.1 24 18

Brewer‘s spent grain 16.8 28.4 27.8

Paper 85-99 0 0-15

Leaves 15-20 80-85 0

Cotton seeds hairs 80-95 5-20 0________

(Jorgensen et al., 2007)

1.2 Breadfruit (Treculia africana)

Treculia africana is a multipurpose tree species commonly known as African breadfruit. It

belongs to the family Moraceae and it grows in the forest zone, particularly the coastal

swamp zone (Agbogidi and Onomeregbor, 2008). African breadfruit is a traditionally

important edible fruit tree in Nigeria (Okafor, 1985) whose importance is due to the potential

use of its seeds, leaves, timber, roots and bark. It is increasingly becoming commercially

important in Southern Nigeria. Baiyeri and Mbah (2006) described African Breadfruit as an

important natural resource which contributes significantly to the income and dietary intake of

the poor. The seeds are used for cooking and are highly nutritious as pointed out by various

authors including; Okafor and Okolo (1974), Okafor (1990) and Onyekwelu and Fayose

(2007). However an important by-product of processing breadfruit seed is the seed hull (seed

coat or seed shell) and this may pose a risk to health as well as environment ( Atuanya et al.,

2012). These hulls are particularly rich in cellulose that could be harnessed for cellulase

production using microbes (Sonde and Odomelam, 2012)

20

Fig. 4: Breadfruit seed hulls (Atuanya et al., 2012)

1.3 Hydrolysis of Cellulose

Cellulose and can be hydrolyzed to sugars and microbially fermented into various products

such as ethanol or chemically converted into other products (Wyman, 1999). The primary

challenge is that the glucose in cellulose is joined by beta bonds in a crystalline structure that

is far more difficult to depolymerize than the alpha bonds in amorphous starch (Wyman,

1999). There are 2 broad based approaches to cellulose hydrolysis. These are acid based

hydrolysis and enzymatic hydrolysis.

1.3.1 Acid hydrolysis

When heated to high temperatures with dilute sulfuric acid, long cellulose chains break down

to shorter groups of molecules that release glucose that can degrade to hydroxymethyl

furfural (McParland et al., 1982). Generally, most cellulose is crystalline, and harsh

conditions (high temperatures, high acid concentrations) are needed to liberate glucose from

these tightly associated chains. Furthermore, yields increase with temperature and acid

concentration, reaching about 70% at 260 C (McParland et al., 1982). However, pyrolysis

21

and other side reactions become very important above about 220⁰C, and the amount of tars

and other difficult to handle by-products increases as the temperature is raised above these

levels (Brennan et al., 1986). In addition, controlling reaction times for maximum glucose

yields of only about 6 seconds at about 250ºC with 1% sulphuric acid presents severe

commercial challenges (Brennan et al., 1986).

1.3.2 Enzyme hydrolysis

Enzymatic hydrolysis has a potential to overcome many of the drawbacks of acid hydrolysis.

The conversion is carried out under mild conditions, thus greatly reducing the cost of

hydrolysis equipment. Sugars decomposition is avoided, thus eliminating this cause for loss

in yield (Hinz et al., 2009). Costly neutralization and purification equipment is unnecessary,

and disposal of waste streams from acid neutralization are eliminated. Balancing these

potential savings, extensive pre-treatment to breakdown to lignin and increase cellulase

accessibility is required to achieve good yields. The cost of high activity cellulolytic enzymes

is at very present very high (Wyman, 1999). Mutations and selection methods have been used

to develop and isolate Fusarium and Trichoderma strains of high cellulolytic activity

(Wyman, 1999).. In all, enzyme hydrolysis is a better alternative to industrial acid hydrolysis,

given the relatively low cost of enzyme hydrolysis and also higher yields of products

obtained via enzyme hydrolysis.

1.4 Cellulases

Cellulase refers to a class of enzymes produced chiefly by fungi, bacteria, and protozoans that

catalyze the cellulolysis (or hydrolysis) of cellulose (Golan, 2011). Cellulases catalyze the

hydrolysis of 1,4-β-D-glucosidic linkages in cellulose, and play a significant role in nature by

recycling this polysaccharide, which is the main component of the plant cell wall. Cellulases

work in synergy with other hydrolytic enzymes in order to obtain the full degradation of the

polysaccharide to soluble sugars, namely cellobiose and glucose, which are then assimilated

by the cell (Morana et al, 2011).

22

The enormous potential that cellulases have in biotechnology is the driving force for

continuous basic and applied research on these biocatalysts from fungi and bacteria.

Cellulases are found in many fields, such as animal feeding, brewing and wine, food, textile

and laundry, pulp and paper products. The growing interest in the conversion of

lignocellulosic biomass into fermentable sugars has generated an additional interest for

cellulases and their related enzymes (Morana et al, 2011).

1.4.1 Classification of Cellulase

Enzymes are designated according to their substrate specificity, based on the guidelines of the

international union of biochemistry and molecular biology (IUBMB). The cellulases are

grouped with many of the hemicellulases and other polysaccharidases as o-glycoside

hydrolases (EC 3.2.1.x) (Morana et al., 2011). Since the substrate specificity classification is

sometime little informative, because the complete range of substrates is only rarely

determined for individual enzymes, an alternative classification of glycoside hydrolases (GH)

into families based on amino acid sequence similarity has been suggested (Henrissat, 1991;

Henrissat and Bairoch, 1993; Henrissat and Bairoch, 1996). In addition, Henrissat et al.

(1998) have proposed a new type of nomenclature for glycoside hydrolases in which the first

three letters designate the preferred substrate, the number indicates the glycoside hydrolase

family, and the following capital letter indicates the order in which the enzymes were first

reported. For example, the enzymes CBHI, CBHII, and EGI of trichoderma reesei are

designated cel7a (CBHI), cel6a (CBHII), and cel6b (EGI).

Due to the great increase of identified glycoside hydrolases, Coutinho and Henrissat have

created an integrated database which is continuously updated (http://www.cazy.org/)

(Coutinho and Henrissat, 1999). At the 13 july 2010 update, glycoside hydrolases were

grouped into 118 families. In addition, 876 glycoside hydrolases have not yet assigned to a

family (glycoside hydrolase family ―non-classifiedǁ) because some of them display weak

similarity to established GH families, but they are too distant to allow a reliable assignment.

Cellulases are found in several different GH families (5, 6, 7, 8, 9, 12, 44, 45, 48, 51, 61, and

74), suggesting convergent evolution of different folds resulting in the same substrate

specificity (Morana et al., 2011). Gh family 9 contains cellulases from bacteria (aerobic and

anaerobic), fungi, plants and animals (protozoa and termites). Other families only group

23

hydrolases from a specific origin, as GH family 7 which contains only fungal hydrolases and

gh family 8 which contains only bacterial hydrolases. Cellulases from the same

microorganism can also be found in different families (e.g. The Clostridium thermocellum

cellulosome contains endoglucanases and exoglucanases from families 5, 8, 9, 44, and 48)

(Morana et al., 2011).

1.4.2 Types of cellulases

Cellulases, responsible for the hydrolytic cleavage of cellulose, are composed of a complex

mixture of enzymes with different specificities to hydrolyse glycosidic bonds. Cellulases can

be grouped into three major classes viz. Endoglucanase, exoglucanase and β-glucosidase.

1.4.2.1 Endoglucanase (EC 3.2.1.4)

Endoglucanases, often called carboxy methyl cellulases (CMCase), are proposed to initiate

random attack at multiple internal sites in the amorphous regions of the cellulose fiber to

open up sites for subsequent attack of cellobiohydrolases (Sunil et al., 2011). This action

results in a rapid decrease of the polymer length and in a gradual increase of reducing sugars

concentration (Morana et al., 2011)

1.4.2.2 Exoglucanase (EC 3.2.1.91)

Exoglucanase, better known as cellobiohydrolase, is the major component of the microbial

cellulase system accounting for 40-70% of the total cellulase proteins and can hydrolyse

highly crystalline cellulose. It removes mono-and dimers from the end of the glucose chain

(Sunil et al., 2011).

1.4.2.3 β-glucosidase (EC 3.2.1.21)

Β-glucosidase also known as cellobiase hydrolyses glucose dimers (cellobiose) and in some

cases cello-oligosaccharides to release glucose units (Sunil et al., 2011). These enzymatic

components act sequentially in a synergistic system to facilitate the breakdown of cellulose

and the subsequent biological conversion to β-glucose.

24

1.5 Mechanism of action of cellulases

Cellulolytic enzymes hydrolyze the 1,4-β-glycosidic bonds in cellulose, but they differ in

their specificities based on the macroscopic features of the substrate. They are progressive

enzymes when they interact with a single polysaccharide strand continuously, and non-

progressive types when they interact once and then, the polypeptidic chain disengages to

attack another polysaccharide strand (Morana et al., 2011) the enzymatic hydrolysis of

cellulose requires a carbohydrate binding module (CBM) that binds and arranges the catalytic

components on the surface of the substrate. Cellulases from fungi have a two-domain

structure with one catalytic domain, and one cellulose binding domain, that are connected by

a flexible linker. However, there are also cellulases that lack cellulose binding domain

(Morana et al., 2011)

The following are three types of reaction catalyzed by cellulases:

1) breakage of the non-covalent interactions present in the crystalline structure of

cellulose (endo-cellulase)

2) hydrolysis of the individual cellulose fibers to break it down into smaller sugars

(exo-cellulase)

3) hydrolysis of disaccharides and tetrasaccharides into glucose (beta-glucosidase).

25

Fig. 5: Mechanism of cellulolysis (Zhang et al., 2006).

1.6 Molecular biology of cellulase

Here, we shall briefly consider cellulase e4 from Thermomonospora fusca. T. Fusca is a

filamentous thermophilic soil bacterium and an important species degrading cellulose and

hemicelluloses in plant residues (Lykidis et al., 2007). Cellulase produced by T. Fusca is

unusual in that it has characteristics of both exo- and endo- cellulases (Sakon et al., 1997).

26

Fig. 6: The 3D structure of cellulase E4 from T. fusca ( Sakon et al., 1997)

The complete genome sequence shows that T. fusca has a single circular chromosome of

3,642,249 bp predicted to encode 3,117 proteins and 65 rna species with a coding density of

85% (Lykidis et al., 2007). Genome analysis reveals the existence of 29 putative glycoside

hydrolases in addition to the previously identified cellulases and xylanases. T. fusca has been

the source organism for isolating and studying multiple secreted cellulases and other

carbohydrate degrading enzymes (Hu and Wilson, 1988). Using classical biochemical

methods, six different cellulases have been identified. Four endocellulase genes (Hu and

Wilson, 1988) and two exocellulase (Irwin et al., 2000). In addition an intracellular β-

glucosidase and extracellular xyloglucanase have been cloned and characterized (Irwin et al.,

2000).

27

1.7 Production of Cellulase

Cellulases are well established in different industrial areas, and are currently the third largest

industrial enzyme worldwide, by dollar volume, mainly because of their use in cotton

processing and paper recycling, as detergent industry enzymes, and in juice extraction and

animal feeding additives as well (Nascimento and Coehlo, 2011). Here, we discuss cellulase

producing organisms, methods of fermentation and factors affecting enzyme production.

1.7.1 Cellulase Producing Microganisms

Microganisms involved in cellulase production will be grouped into two broad groups-

mesophilic microorganisms and thermophilic microorganisms.

1.7.1.1 Mesophilic microorganisms

Microorganisms growing best at moderate temperatures (between 10ºC and 45°C) are named

mesophiles. They represent the majority of microbial species on Earth, and their habitats

include the soil, the human body, the animals, etc. There are many mesophilic Bacteria and

Fungi that play a significant role in the carbon cycle on Earth, and there is increasing interest

in the enzymes from these microorganisms, since they have a key function in the conversion

of plant biomass into useful products (Morana et al., 2011).

28

Table 2. Some mesophilic cellulolytic Bacteria

___________________________________________________________________________ Microorganism Gram reaction Growth (°C) Growth conditions

Temperature

___________________________________________________________________________ Acetivibrio cellulolyticus - 37 Anaerobic

Bacillus megaterium + 30 Aerobic

Bacillus pumilus + 30 Aerobic

Bacteroides cellulosolvens - 35 Anaerobic

Butyrivibrio fibrisolvens + 37 Anaerobic

Cellulomonas fimi + 30 Aerobic

Cellulomonas fermentans + 30 Aerobic

Cellulomonas flavigena + 30 Aerobic

Cellulomonas gelida + 30 Aerobic

Cellulomonas iranes + 28 Aerobic

Cellulomonas persica + 28 Aerobic

Cellulomonas uda + 30 Aerobic

Cellvibrio mixtus - 20 Aerobic

Clostridium acetobutylicum + 37 Anaerobic

Clostridium cellulolyticum + 35-37 Anaerobic

Clostridium cellulofermentans - 40 Anaerobic

Clostridium cellulovorans - 37 Anaerobic

Clostridium herbivorans + 37 Anaerobic

Clostridium hungatei - 30 Anaerobic

Clostridium josui - 45 Anaerobic

Clostridium papyrosolvens - 25 Anaerobic

___________________________________________________________________________

(Morana et al., 2011)

Identification, purification and characterization of cellulases are continuously increasing and

always in progress, with incessant research and isolation of new microorganisms able to

produce novel cellulolytic activities. As an example, a bacterial strain, TR7-06(T), showing

high sequence similarity (98.5 %) to Cellulomonas uda DSM 20107(T), was isolated from

29

compost at a cattle farm near Daejeon, Republic of Korea. The isolated type strain of a novel

Cellulomonas species, named Cellulomonas composti sp. nov., possesses endoglucanase and

β-glucosidase activities (Kang et al., 2007). A microorganism capable of hydrolyzing rice

hull, one of the major cellulosic waste materials in Korea, was isolated from soil and

identified as Bacillus amyloliquefaciens DL-3 (Lee et al., 2008). Based on the characteristics

of this novel strain of Bacillus, Lee et al., (2008) aimed to develop an economical process for

production of cellulases by using cellulosic waste as inexpensive and widely distributed

carbon source. The new isolate produced an extracellular cellulase with an estimated

molecular mass of about 54.0 kDa. The deduced amino acid sequence of the cellulase from B.

amyloliquefaciens DL-3 showed high identity to cellulases from other Bacillus species, a

modular structure containing a catalytic domain of the GH family 5, and a cellulose-binding

module type 3 (CBM3). The purified enzyme was optimally active at 50°C and pH 8.0, and

showed broad thermal and pH stability ranging from 40 to 80°C and from 4.0 to 9.0,

respectively (Lee et al., 2008)

30

Table 3. Some mesophylic cellulolytic fungi

___________________________________________________________________________ Microorganism Growth Growth (°C) Growth conditions

Temperature

__________________________________________________________________________

Acremonium cellulolyticus 24 Aerobic

Anaeromyces mucronatus 37 Anaerobic

Aspergillus glaucus 30 Aerobic

Aspergillus niger 30 Aerobic

Aspergillus terreus 35 Aerobic

Caecomyces communis 37 Anaerobic

Ceratocystis paradoxa 20 Aerobic

Chrysosporium lucknowense 25-43 Aerobic

Cyllamyces aberensis 37 Anaerobic

Fusarium solani 25 Aerobic

Neocallimastix frontalis 37 Anaerobic

Neocallimastix patriciarum 37 Anaerobic

Orpinomyces sp. 37 Anaerobic

Penicillium funiculosum 24 Aerobic

Penicillium pinophilum 24 Aerobic

Phanerochaete chrysosporium 35 Aerobic

Piptoporus betulinus 25 Aerobic

Piromyces sp. 39 Anaerobic

Piromyces equi 39 Anaerobic

Pycnoporus cinnabarinus 24 Aerobic

Rhizopus oryzae 30 Aerobic

___________________________________________________________________________

(Morana et al., 2011)

Fungal cellulases are well-studied enzymes used in various industrial processes (Bhat, 2000).

A variety of aerobic and anaerobic Fungi are producers of cellulose-degrading enzymes. The

aerobic Fungi play a major role in the degradation of plant materials and are found on the

decomposing wood and plants, in the soil, and on the agricultural residues. The cellulase

31

systems of the aerobic Fungi Trichoderma reesei, T. koningii, Penicillium pinophilum,

Phanerochaete chrysosporium, Fusarium solani, Talaromyces emersonii, and Rhizopus

oryzae are well characterized (Bhat and Bhat, 1997). Much of the knowledge on enzymatic

depolymerization of cellulosic material has come from Trichoderma cellulase system. In

particular, the cellulase system of T. reesei (initially called T. viride) has been the focus of

research for 50 years (Reese and Mandels, 1971). A lot of work on cellulases has been

directed toward this fungus since it produces readily, and in large quantities, a complete set of

extracellular cellulases, and consequently, it has a high commercial value (Claeyssens et al.,

1998; Miettinen-Oinonen and Suominen 2002). In fact, T. reesei is capable of secreting more

than 30 g/L of protein into the extracellular medium (Conesa et al., 2001). It has been

reported that T. reesei possesses two CBH (cellobiohydrolase) genes, cbh1-2, and eight EG

(endoglucanase) genes, egl1-8, and that CBH I–II and EG I–VI are secreted proteins

(Foreman et al., 2003).

1.7.1.2 Thermophilic Microorganisms

The thermophilic microorganisms represent a unique group growing at temperatures that may

exceed 100°C. More precisely, thermophilic microorganisms thrive at temperatures from 65

to 85°C, and hyperthermophiles grow at temperatures of above 85°C (Morana et al., 2011).

Hyperthermophiles are microorganisms within the Archaea domain although some bacteria

are able to tolerate temperatures around 100°C. An extraordinary heat-tolerant

hyperthermophile is Methanopyrus kandleri, discovered on the wall of a black smoker from

the Gulf of California at a depth of 2000 m, at temperatures of 84-110°C. It can survive and

reproduce at 122°C (Takai et al., 2008). Thermophilic and hyperthermophilic

microorganisms have received considerable attention as sources of thermostable cellulolytic

enzymes, as the properties of these biocatalysts make them interesting candidates for

industrial applications. Running biotechnological processes at elevated temperatures has

many advantages. High temperature has a significant influence on the solubility of the

substrates (especially if viscous or polymers) and on the reaction rate. Moreover, problems of

microbial contamination can be avoided when a reaction is performed at elevated temperature

(Takai et al., 2008).

32

Table 4: Some (hyper) thermophilic cellulolytic Bacteria and Archaea

___________________________________________________________________________ Microorganism Gram reaction Growth Growth condition

Temperature (°C)

___________________________________________________________________________

Acidothermus cellulolyticus + 55 Aerobic

Alicyclobacillus acidocaldarius + 60 Aerobic

Anaerocellum thermophilum + 75 Anaerobic

Aquifex aeolicus - 85-95 Aerobic

Caldibacillus cellulovorans + 68 Aerobic

Caldicellulosiruptor saccharolyticus - 70 Anaerobic

Clostridium stercorarium + 65 Anaerobic

Clostridium thermocellum + 60 Anaerobic

Dictyoglomus thermophilus - 73 Anaerobic

Dictyoglomus turgidus - 72 Anaerobic

Pyrococcus abyssi - 96 Anaerobic

Pyrococcus furiosus - 98 Anaerobic

Pyrococcus horikoshii - 98 Anaerobic

(Morana et al., 2011)

Thermostable cellulases are of great biotechnological interest (Hongpattarakere, 2002). A

number of cellulolytic thermophilic Bacteria have been isolated, and many cellulose

degrading enzymes have been identified, characterized, cloned and expressed (Bergquist et

al., 1999). Conversely, screening of hyperthermophilic Bacteria for cellulose-degrading

enzymes has revealed that the presence of such enzymes is rather rare in this group. In

addition, among the Archaea, only the genus Pyrococcus and Sulfolobus have been found to

process thermoactive cellulases. Few aerobic thermophilic microorganisms have been

described to produce cellulases in comparison with the anaerobic ones. Acidothermus

cellulolyticus, isolated from 55-60°C acidic water and mud samples collected in Yellowstone

National Park, produces at least three thermostable endoglucanases (Mohagheghi, 1986). One

of them, E1 belonging to GH family 5, was crystallized, while properties and application of

33

the other enzymes are protected by patents (Sakon et al., 1996). The aerobic thermophilic

bacterium Rhodothermus marinus, isolated from a submarine hot spring at Reykjanes, NW

Iceland (Alfredsson et al., 1988), produces one higly thermostable cellulase (Cel12A) which

retains 50% activity after 3.5 h at 100°C (Hreggvidsson et al., 1996).

Table 5: Some (hyper) thermophilic cellulolytic Fungi

___________________________________________________________________________ Microorganism Growth Temperature (°C) Growth conditions

___________________________________________________________________________ Chaetomium thermophilum 45-55 Aerobic

Humicola grisea 45 Aerobic

Humicola insolens 40-50 Aerobic

Melanocarpus albomyces 45-55 Aerobic

(Morana et al., 2011)

Among the thermophilic Fungi, only a few number is described to be cellulase-producer The

thermophilic filamentous fungus Humicola sp. has been known to produce several cellulases,

and some of the genes have been cloned, sequenced and expressed (Takashima et al., 1996).

The cellulase system of the thermophilic fungus Humicola insolens possesses a battery of

enzymes that allows the efficient utilization of cellulose. This system, homologous to that of

T. reesei, contains five endoglucanases: EGI (Cel7B), EGII (Cel5), EGIII (Cel12), EGV

(Cel45A), and EGVI (Cel6B) in addition to two cellobiohydrolases: CBHI (Cel7A), and

CBHII (Cel6A) (Schulein, 1997).

The thermophilic fungus Chaetomium thermophile var. dissitum, was able to produce in the

culture medium all the enzymes involved in cellulose breakdown, namely endoglucanase

(41.0 kDa), exoglucanase (67.0 kDa) and β-glucosidase (Eriksen and Goksoyr, 1977). Lu et

al. (2002) reported that C. thermophile secreted in the culture medium a glycosylated

endocellulase with an apparent molecular weight of 67.8 kDa, as determined by SDS-PAGE.

The enzyme was optimally active at pH 4.0-4.5 and 60°C, and it retained 30% activity after

60 min at 70°C.

34

Melanocarpus albomyces, a rare true thermophilic Ascomycete capable of growing copiously

at 50°C, has been documented to produce high levels of endoglucanases under optimized

culture conditions (Jatinder et al., 2006). The endoglucanases from this fungus have been

recognized as potentially important in denim washing. In fact, the supernatant from M.

albomyces worked well in biostoning, with low backstaining. Three cellulases were identified

and purified to homogeneity, and two of them were endoglucanases with apparent molecular

masses of 20.0 kDa (Cel45A) and 50.0 kDa (Cel7A) (Miettinen-Oinonen et al., 2004).

The thermophilic fungus Thermoascus aurantiacus produces high levels of cellulase

components when grown on lignocellulosic carbon sources such as corncob and cereal straw

(Khandke et al., 1989). As these enzyme components are remarkably stable over a wide

range of pH and temperatures, they appear to have great commercial potential. A major

extracellular endoglucanase, with a molecular mass of 34.0 kDa, was purified and

characterized (Parry et al., 2002). It was optimally active at 70-80°C and pHs 4.0-4.4, and it

was stable at pH 5.2 and up to 60°C for 48 h. At 70°C and pH 5.2 the enzyme retained 40%

of the original activity after 48 h (Parry et al., 2002). The cellulase exhibited the highest

activity toward CMC; barley β-glucan and lichenan were also hydrolyzed, but the enzyme

was inactive on laminarin, confirming that it was an endoglucanase and was specific toward

β-1,4 linked polysaccharides (Parry et al., 2002).

1.7.2 Aspergillus spp

A large number of fungi have been reported in municipal solid waste. The Aspergillus sp.

was predominantly high among the other fungal species (Gautam et al., 2010). Most of the

work on fungal Cellulase is centered on the saccharification of cellulose by Aspergillus

(Milala et al., 2005) although Cellulase production on different carbon sources by Aspergillus

and other fungi have been reported (Ruijter and Visser, 1997).

Aspergillus is a genus consisting of several hundred mold species found in various climates

worldwide (Bennet, 2010). Aspergillus is a potential producer of cellulases (Mohammed et

al., 2005). The organisms are widespread in nature and are typically found in soil and

decaying organic matter, such as compost heaps, where they play an essential role in carbon

35

and nitrogen recycling (Bennet, 2010). The two species of aspergillus used for this work are

A. fumigatus and A. flavus.

Fig. 7: A. flavus (Winiati, 2013)

Fig. 8: A. fumigatus (Mirhendi, 2000)

36

1.8 Fermentation Methods

Fermentation is the technique of biological conversion of complex substrates into simple

compounds by various microorganisms such as bacteria and fungi. In the course of this

metabolic breakdown, they also release several additional compounds apart from the usual

products of fermentation, such as carbon dioxide and alcohol (Subramaniyam and Vimala,

2012). These additional compounds are called secondary metabolites. Secondary metabolites

range from several antibiotics to peptides, enzymes and growth factors (Machado et al.,

2004). Two broad methods will be considered for the production of cellulase enzyme. These

are solid state fermentation (SSF) and submerged fermentation (SmF) processes.

1.8.1 Solid-State Fermentation (SSF)

SSF utilizes solid substrates, like bran, bagasse, and paper pulp. The main advantage of using

these substrates is that nutrient-rich waste materials can be easily recycled as substrates

(Subramaniyam and Vimala, 2012). In this fermentation technique, the substrates are utilized

very slowly and steadily, so the same substrate can be used for long fermentation periods

(Subramaniyam and Vimala, 2012). Hence, this technique supports controlled release of

nutrients. SSF is best suited for fermentation techniques involving fungi and microorganisms

that require less moisture content. However, it cannot be used in fermentation processes

involving organisms that require high water activity (aw), such as bacteria. (Babu and

Satyanarayana, 1996).

1.8.2 Submerged Fermentation (SmF)/Liquid Fermentation (LF)

SmF utilizes free flowing liquid substrates, such as molasses and broths. The bioactive

compounds are secreted into the fermentation broth. The substrates are utilized quite rapidly;

Hence, they need to be constantly replaced/supplemented with nutrients (Subramaniyam and

Vimala, 2012). This fermentation technique is best suited for microorganisms such as bacteria

that require high moisture content. An additional advantage of this technique is that

purification of products is easier (Subramaniyam and Vimala, 2012). SmF is primarily used in

37

the production of secondary metabolites that need to be used in liquid form. Submerged

fermentation method for enzyme production is usually preferred since the enzyme will be

obtained in liquid form thus making for easy purification and characterization.

1.9 Factors Affecting Cellulase Enzyme Production

1.9.1 Chemical Factors

1.9.1.1 Effect of Carbon Sources

Since any cellulose biotechnological process is likely to base on crude enzymes, it is

important to increase their activities in the culture supernatants by selecting the best carbon

and nitrogen sources and optimizing their concentrations (Gomes et al., 2000). Cellulase

production is dependent on the nature of the carbon source used in the culture medium.

Various lignocellulose carbon sources have been tested for their ability to induce cellulase

production. Besides, the efficiency of enzyme production also depends on the bare chemical

composition of the raw material, accessibility of various components and their chemical and

physical associations. Wheat straw, rice straw and corn stover have been known as ideal

substrate for cellulose production (Panagiotou et al., 2003; Mishra and Nain, 2010). Several

investigations have indicated that cellulases are inducible enzymes, and different carbon

sources have been used to find their role in effecting the enzymatic levels. Cellobiose (2.95

mM) may act as an effective inducer of cellulases synthesis in Nectria catalinensis (Pardo

and Forchiassin, 1999). An increased rate of endoglucanase biosynthesis in Bacillus sp. was

reported in the presence of cellobiose or glucose (0.2%) in the culture medium (Paul and

Verma, 1990). Yeoh et al. (1986) had reported the inhibition of β-glucosidase activity at

higher concentrations of cellobiose to an extant of 80%; similarly, laminaribiose and glucose

also led to a 55–60% reduction in the enzymatic activity. Later, Shiang et al. (1991)

described a possible regulation mechanism of cellulose biosynthesis and proposed that sugar

alcohols, sugar analogues, xylose, glucose, sucrose, sorbose, cellobiose, methylglucoside etc.

at a particular concentration may induce a cellulose regulatory protein called cellulase

activator molecule (CAM). The level and yield of CAM get affected possibly due to substrate

concentration and some unknown factors imparted by moderators. Many different agro-

38

industrial wastes, synthetic or natural, have been examined as the carbon source for the

process. Among the cellulosic materials, sulfate pulp, printed papers, mixed waste paper,

wheat straw, paddy straw, sugarcane bagasse, jute stick, carboxymethylcellulose, corncobs,

groundnut shells, cotton, ball milled barley straw, delignified ball milled oat spelt xylan, larch

wood xylan, etc. have been used as the substrates

for cellulase production (Singh et al., 1991; Mishra and Nain, 2010). The observations

indicated that the production of cellulases increased with increase in substrate concentration

up to 12% during solid-state-fermentation using Aspergillus niger. Further increase in

substrate concentration decreased the production levels. This might have been due to

limitation of oxygen in the central biomass of the pellets, and exhaustion of nutrients other

than energy sources. Martins et al. (2008) and Steiner et al. (1993) also demonstrated that

carboxymethycellulose or cereal straw (1%, w/w) would be the best carbon source compared

to sawdust for CMCase and β-glucosidase production using Chaetomium globosum as the

producer organism. In contrast, 3% malt extract or water hyacinth was found optimum for

CMCase, FPase and β-glucosidase as observed with lactose as an additional carbon sources

(Mukhopadhyey and Nandi, 1999). However, the saccharification of alkali-treated bagasse at

higher substrate levels of 4% w/v was also reported (Singh et al., 1991). Interestingly, higher

concentrations (2.5–6.2% w/v) of carbon source were observed to be suitable for maximum

saccharification when cellobiose was supplemented in the medium containing delignified rice

straw, news print or other paper wastes as substrates ( Ju and Afolabi, 1999).

1.9.1.2 Effect of Nitrogen Sources

The effects of nitrogen sources on cellulase production are variable with respect to the fungi

used (Kachlishvili et al., 2006). Enzyme production is affected significantly under different

concentrations of nitrogen sources (Panagiotou et al., 2003). With different nitrogen sources,

enzyme activities are higher with organic nitrogen (Gao et al., 2008). Maximum cellulase

activity has been obtained with yeast extract (Gao et al., 2008), though other researchers

found that inorganic nitrogen sources produce an optimal result (Kalogeris et al., 2003). The

effect of different inorganic nitrogen sources such as ammonium sulfate, ammonium nitrate,

ammonium ferrous sulfate, ammonium chloride and sodium nitrate have been studied.

Among these, ammonium sulfate (0.5 g /L) led to maximum production of cellulases (Singh

et al., 1991). In contrast, Menon et al. (1994) observed a significant decrease in enzymatic

39

levels in the presence of ammonium salts as the nitrogen source. However, an increase in the

level of β-glucosidase was reported when corn steep liquor (0.8% v/v) was added into the

production medium. Corn steep liquor also resulted in 3-5 fold induction of endo- and

exoglucanase levels with synthetic cellulose, wheat straw and wheat bran as the substrates.

Enzyme production was sensitive to corn steep liquor (0.88 g/L), and production increased

significantly when mixed nitrogen sources (corn steep liquor and ammonium nitrate) were

supplied (Steiner et al., 1993). However, additional incorporation of nitrogen sources into

medium scale up the cost of the process (Sunil et al., 2011).

1.9.1.3 Phosphorus Sources

Phosphorus is an essential requirement for fungal growth and metabolism. It is an important

constituent of phospholipids involved in the formation of cell membranes. Besides its role in

linkage between the nucleotides forming the nucleic acid strands, it is involved in the

formation of numerous intermediates, enzymes and coenzymes essential in carbohydrate

metabolism, other oxidative reactions and intracellular processes (Singh et al., 1991).

Different phosphate sources such as potassium dihydrogen phosphate, tetra-sodium

pyrophosphate, sodium β-glycerophosphate and dipotassium hydrogen phosphate have been

evaluated for their effect on cellulases production (Garg and Neelkantan, 1982). It has been

widely known that potassium dihydrogen phosphate is the most favorable phosphorus source

for cellulase production (Sunil et al., 2011)

1.9.2 Physical Factors

1.9.2.1 pH

Different physical parameters influence the cellulose bioconversion, and pH is an important

factor affecting cellulase production (Pardo and Forchiassin, 1999). The effect of pH on

cellulase production has been analysed using Aspergillus niger, and found that pH 5.5 was

optimal for maximum cellulase production. On other side, the pH range of 5.5–6.5 was

optimal for β-glucosidase production from Penicillium rubrum (Menon et al., 1994). Eberhart

et al. (1977) has reported that production and release of cellulase from Neurospora crassa

depends on pH of the medium and maximum release occurs at pH 7.0, whereas the enzyme

40

remained accumulated in the cell at pH 7.5. Similarly, pH 7.0 is suitable for extracellular

production of cellulase from the Humicola fuscoatra (Rajendran et al., 1994). The adsorption

behavior of cellulases has been found to be affected by pH of the medium. Kim et al. (1988)

had reported maximum adsorption of cellulase from Aspergillus phoenicus at pH 4.8–5.5.

The pH range 4.6–5.0 has been found suitable for CMCase, filterpaperase (FPase) and β-

glucosidase production with Aspergillus ornatus and Trichoderma reesei (Mukhopadhyey

and Nandi, 1999).

1.9.2.2 Temperature

Temperature has a profound effect on lignocellulosic bioconversion. The temperature for

assaying cellulase activities is generally within 50–65 °C for a variety of microbial strains

(Menon et al., 1994; Steiner et al., 1993), whereas growth temperature of these microbial

strains was found to be in the 25–30 °C (Macris et al., 1989). Similarly Penicillium

purpurogenum, Pleurotus florida and Pleurotus cornucopiae show higher growth at 28 °C

but maximum cellulase activities at 50 °C (Steiner et al., 1993) and about 98, 59 and 76% of

the CMCase, FPase and β-glucosidase activities, respectively, retained after 48 h at 40 °C.

Researchers have shown that temperature influences the cellulose-cellulase adsorption

behaviour. A positive relationship between adsorption and saccharification of cellulosic

substrate was observed at temperature below 60 °C. The adsorption activities beyond 60 °C

decreased possibly because of the loss of enzyme configuration leading to denaturation of the

enzyme activity (Van-Wyk, 1997). Bronnenmeier and Staudenbauer (1988) reported that

extracellular as well as cell bound β-glulcosidase from Clostridium stercorarium required an

identical temperature of 65 °C for their activity. Further increase in temperature led to a sharp

decrease in the enzyme activity. Some of the thermophilic fungi having maximum growth at

or above 45–50 °C produce cellulase with wide temperature optima (50–78 °C) (Wojtczak et

al., 1987).

41

1.10 Applications of Cellulases

1.10.1 Cellulases in Brewing and Wine Biotechnology

The macerating enzymes, comprising cellulases, hemicellulases and pectinases, hydrolyze the

plant cell wall and, consequently, can be used in brewing and wine biotechnology to improve

the quality of finished products and avoid the use of chemicals. Enzyme preparations are used

in the brewing and distilling industries to decrease the viscosity of the mash and to improve

the overall efficiency of the process. In fact, cellulolytic and hemicellulolytic enzymes allow

the conversion of undigestible lignocellulosic biomass into fermentable sugars, with

consequent increase of alcohol yield.

1.10.1.1 Beer Brewing Process

Barley is the most common cereal used for the production of beer although wheat, corn, and

rice are also widely used. The main processes involved in beer production include milling to

reduce the size of the dry malt in order to increase the availability of the carbohydrates;

mashing where water is added to the malt; lautering where spent grains are removed from the

wort, boiling of the wort with flavouring hops, fermentation of the wort liquor, maturation,

conditioning, filtration and packaging of the final product. The high concentration of β-

glucan in the brewing process, resulting from unsuitable brewing process or low quality

barley, produces high viscosity of beer, formation of gelatinous precipitate, decrease of the

extract yield, and lower run-off of wort (Bamforth, 1994; Bhat, 2000; Guo et al., 2010). In

brewing process, cellulases are used during the mashing stage in order to hydrolyze excess β-

glucans and reduce the viscosity, thus improving the separation of the wort from the spent

grains. Oksanen et al. (1985) observed that the endoglucanase and the cellobiohydrolase from

the Trichoderma cellulase system produced a large reduction of the degree of polymerization

of the β-glucans, and wort viscosity. Moreover, the increased addition of enzymes used

resulted in improved filtering. A. niger, T. reesei, and P. funiculosum, which are generally

recognized as food grade microorganisms, are the major source of cellulases currently used in

the mashing step, as these enzymes provide technological benefit to beer manufacture

(Karboune et al., 2008).

42

1.10.1.2 Wine Production

Wine manufacture is a biotechnological process in which yeast cells and enzymes are

indispensable for ensuring a high quality product. The use of cellulases, hemicellulases and

pectinases during wine making, allows a better skin maceration, and superior color

extraction, particularly important in the production of red wine; in addition, it improves

clarification, filtration, and the overall quality and stability of the wine (Galante et al., 1998).

Pectinase preparations, used in wine making, were lately modified by addition of cellulases

and hemicellulases in small quantities to realize a more complete breakdown of the cells with

consequent fruit liquefaction in a moderately short time period (Plank and Zent, 1993). It has

also been demonstrated that the mixture of macerating enzymes worked better than pectinases

alone in grape processing (Haight and Gump, 1994).

1.10.2 Cellulases in Pulp and Paper Biotechnology

1.10.2.1 Biomechanical Pulping

Mechanical pulping process is electrical energy intensive and results in low paper strength.

Biomechanical pulping, defined as the enzymatic treatment of lignocellulosic materials

before the mechanical pulping step, has shown at least 30% savings in electrical energy

consumption, and significant improvements in paper strength properties. The potential of

enzymatic treatments has been assessed and the processes have proved successful (Gubitz et

al., 1998). Utilization of cellulases from fungal sources (T. reesei, Aspergillus sp.) (Buchert

et al., 1998; Suurnakki et al., 2000) saves 33% electrical energy and significantly improves

paper strength properties. A cellulase preparation produced by the ascomycete fungus

Chrysosporium lucknowense for using in the pulp and paper industry represents, at present,

an attractive alternative to the well-known cellulases from Fungi like Aspergillus sp. and T.

reesei for protein production on a commercial scale (Bukhtojarov et al., 2004; Hinz et al.,

2009).

43

1.10.2.2 Biodeinking

All over the world people give more attention to the environment and so, the recycle of waste

paper has to be considered also as a necessity for the protection of forest and economy. Paper

mill will gain profit from the utilization of recycled fiber, since it is profitable to decrease

pollution, cost, and investment. Conventional deinking technology with alkali is

characterized by a low efficiency on laser printed paper and is not considered

environmentally friendly. Consequently, researchers have concentrated their attention on new

deinking technologies (Moon and Nagarajan, 1998). The principle of enzymatic deinking is

based on the weakening of the connections between toner and fibers due to the enzyme attack

with separation of toner particles from fibers (Yingjuan et al., 2005; Shufang et al., 2005).

The enzymatic deinking allows us to avoid the use of alkali; moreover, using enzymes at

acidic pH it is possible to prevent the yellowing, modify the distribution of the ink particle

size, improve fiber brightness strength, pulp freeness and cleanliness, reduce fine particles

and reduce environmental pollution. Until 2000, the use of enzymes to perform biodeinking

was only investigated at the laboratory scale (Buchert et al., 1998; Bhat, 2000). Subsequently,

a mixture of cellulase, lipase, and amylase was employed in biodeinking process at industrial

level (Morbak and Zimmermann, 1998). The effect of combined deinking technology with

ultrasounds, UV irradiation and enzyme on laser printed paper was investigated. The results

confirmed that the dose of alkali can be reduced using biodeinking technology. Cellulases

from different microorganisms such as A. niger, T. reesei, Humicola insolens, Myceliophtora

fergusii, Chrysosporium lucknowense, Fusarium sp. were used for this purpose (Marques et

al., 2003).

1.10.3 Cellulases in Textile and Laundry Biotechnology

Since the early part of the last century, enzymes such as the cellulases have been used for a

wide range of applications in textile processing in replacement of the traditional methods.

44

1.10.3.1 Biostoning and Biopolishing

Jeans manufactured from denim are one of the world's most popular clothing items. In the

late 1970s and early 1980s, industrial laundries developed methods for producing faded jeans

by washing the garments with pumice stones, which partially removed the indigo dye

revealing the white interior of the yarn, which leads to the faded, worn and aged appearance.

This process was designated as ―stone-washing (Cavaco- Paulo, 1998). The use of 1-2 kg

stones per kg of jeans for 1 h during stone-washing met the market requirements, but caused

several problems including rapid consumption of washing machines, and unsafe working

conditions. As an alternative to the stone-washing, biostoning is by far the most economical

and environmental friendly way to treat denim. The cotton fabrics treated with the enzymes

loose the indigo, which later is easily removed by mechanical abrasion in the wash cycle

(Cavaco- Paulo, 1998; Yamada et al., 2005). The substitution of pumice stones by an

enzymatic treatment has many advantages: washing machines lower consumption and

elevated productivity, short treatment times and less intensive working conditions. Moreover,

it is possible to operate in a more safe environment because pumice powder is not produced,

and the process can be mechanized controlling, with the use of computer, the dosing devices

of liquid cellulase preparations (Bhat, 2000).

In the textile wet processing, the biopolishing is usually carried out with desizing, scouring,

bleaching, dyeing and finishing by utilization of cellulases. However, there are no clear

indications about the best cellulase mixture to use. (Miettinen-Oinonen and Suominen, 2002).

The use of these enzymes allow many improvements such as the removal of short fibers,

surface fuzziness smooth, polished appearance, more color uniformity and brigthness,

improved finishing, and fashionable effects. At last, due to increasing environmental

concerns and constraints being imposed on textile industry, cellulase treatment of cotton

fabrics is an environmentally friendly way of improving the property of the fabrics. In 2007,

Anish et al. (2006) isolated an endoglucanase from the alkalothermophilic bacterium

Thermomonospora sp. The enzyme which is used for denim biofinishing under alkaline

conditions, was effective in removing hairiness with negligible weight loss and imparting

softness to the fabric. Higher abrasive activity with lower back-staining was a preferred

property for denim biofinishing exhibited by the Thermomonospora endoglucanase.

45

1.10.3.2 Laundry

The most important reason to use enzymes in detergents is that they are biodegradable and a

very small quantity of these inexhaustible biocatalysts can replace very large quantity of

chemicals. Since detergents hold ionic and anionic surfactants, and bleaching agents

(oxidizing agents) that can partially or completely denature proteins, the enzymes for laundry

must be resistant to anionic surfactants and oxidizing agents. The accumulation of

microfibrils on the surface of the fabrics makes the fabrics look hairy and scatters incident

light, thereby lessening the brightness of the original colors. In detergent industry, cellulases

are used to remove microfibrils from the surface of cellulosic fabrics, enhancing color

brightness, hand feel and dirt removal from cotton garments that during repeated washings

can become fluffy and dull.

Other notable applications of cellulase are found in the treatment of wastes, production of

biofuels and also in the animal feed industry.

1.11 Aim and Objectives of study

1.11.1 Aim of study

This study is aimed at using microorganisms cultivated on agricultural waste to produce

cellulase enzyme with industrial potential.

1.11.2 Specific Objectives of the Study

This work is therefore designed to achieve the following specific objectives:

• Isolation of crude cellulase secreted by Aspergillus fungi.

• Determination of the protein content of the enzyme.

• To Assay for activity of the cellulase enzymes

• Partial purification of cellulase.

• Characterization of purified cellulase.

46

CHAPTER TWO

MATERIALS AND METHODS

2.1 Materials

2.1.1 Reagents

Chemicals/Regents Manufacturer

Bovine serum albumin (BSA) Sigma Chemical Company (USA)

Folin –Ciocalteau Sigma-Aldrich (USA

Ammonium sulphate British Drug House (BDH) Chem. Ltd (USA)

Tris HCL salt Merek specialist Private limited (Mumbai)

Sephadex G- 50 Sigma Chemicals Company Limited (USA)

All other chemicals used in this work were of analytical grade and were obtained from

reputable sources. Distilled water was used for all preparations of solutions and pH

measurements were made at room temperature using a pH meter.

2.1.2 Apparatus

Weighing balance: Ohaus Dial-O- Gram, Ohaus Cooperation, N.J. USA.

Water bath: Model DK.

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

Milling machine: Thomas Willey Laboratory Mill Model 4, Anthor H

(Thomas Company, Philadelphia, USA)

Autoclave: UDAY BURDON’s Patient Autoclave, India.

Incubator: B and T Trimline incubator.

Centrifuge: Finland Nigeria 80-2B.

Oven: Gallenkamp, England.

pH meter: Ecosan pH meter, Singapore.

47

Microscope: WESO microscope.

Glass wares: Pyrex, England

Uv/visible spectrophotometer: Jenway 6405, England

Visible spectrophotometer: Labscience 721, England

2.2 Methods

2.2.1 Collection of Breadfruit hulls

Breadfruit hulls were obtained from breadfruit processing centres at the Ogige market in

Nsukka town, Enugu State of Nigeria.

2.2.2 Collection of Microorganism

Aspergillus fumigatus and Aspergillus flavus strains were obtained from the post-graduate

laboratory of the Department of Biochemistry, University of Nigeria, Nsukka.

2.2.3 Preparation of Ground Breadfruit hulls

The hulls were sun dried for seven days and then ground into powder with the help of a

milling machine.

2.2.4 Storage of Pure Fungal Isolates

Fungal isolates were obtained from the Department of Microbiology. The pure fungal isolates

were maintained on potato dextrose agar (PDA) slopes or slants as stock cultures. PDA media

were prepared according to the manufacture’s description. In the procedure, 3.9g of PDA

powder was weighed and added to a small volume of distilled water and made up to 100 ml.

The medium was autoclaved at 121oC (15 psi) for 15 min. It was allowed to cool to 45

oC and

then poured into Petri dishes and allowed to gel. The plates were then incubated in a B & T

Trimline incubator at 37oC for 24hr to check for sterility.

48

2.2.5 The Fermentation Broth

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

containing 100 ml of sterile cultivation medium optimized for cellulase with 0.1% NH4NO3,

0.1% NH4 H2PO4, 0.1% MgS04.7H2O and 1% breadfruit hulls. The flask was stoppered with

aluminium foil and autoclaved at 121oC (15 psi) for 15 min. The experiment was performed

in duplicate for both fungal species.

2.2.6 Inoculation of the Broth

From the PDA slants, fresh plates were prepared and inoculated. Three days old cultures were

used to inoculate the flasks. In every 50 ml of the broth, two discs of the respective fungal

isolates were added using a cork borer of diameter 10 mm and then plugged properly. The

culture was incubated for 7 days at room temperature (30oC). This experiment was also

performed for both species of Aspergillus.

2.2.7 Harvesting of the Fermented Broth

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

separated by filtration. Each day, the filtrate was analyzed for cellulase activity till the 7th

day of fermentation.

2.2.8 Mass Production of Enzyme

After the 7 days pilot studies, the day of peak cellulase activity was chosen for mass

production of enzyme from the respective fungal isolates. Several 250 ml Erlenmeyer flasks

were used to produce 750 ml of the enzymes using the method described in sections 2.2.6 and

2.2.7. Harvesting was carried out on the respective peak days of enzyme activity.

49

2.2.9 Procedure for Protein Determination

Protein determination was done by Lowry’s method (1951). For protein standard curve, the

reaction mixture contained 0.0-1.0 ml of protein stock solution (2 mg/ml BSA) in test tubes

arranged in triplicates. The volume was made up to 1 ml with 0.05M sodium acetate buffer.

But for the test mixture, 0.1 ml of sample enzyme was mixed with 0.9ml of buffer. In either

case, 2ml of solution E was added and allowed to stand at room temperature for 10min. Then,

0.2ml of solution C (dilute Folin-Ciocalteau phenol reagent) was added with rapid mixing.

After standing for 30min, absorbance was read at 750nm using UV spectrophotometer.

Absorbance values were converted to protein concentration by extrapolation from the

standard curve.

2.2.10 Cellulase Assay

Cellulase activity was evaluated by assaying for the glycosidic activity of the enzyme. This

was achieved by measuring the release of reducing groups from filter paper, carboxymethyl

cellulose (CMC) and cellobiose using a modification of the 3, 5-dinitrosalicylic acid (DNS)

reagent assay method described by Miller (1959) as contained in Ghose (1997).

2.2.10.1 Cellobiase assay

The reaction mixture containing 0.2 ml 15 mM cellobiose in 0.05 M sodium acetate buffer of

pH 5.5 and 0.2 ml enzyme solution was incubated for 30 mins at 50 ºC. 1ml of DNS reagent

was added and the reaction was stopped by boiling the mixture in a boiling water bath for

10mins. The mixture volume was made up to 4 ml with 1 ml of Rochelle salt solution. The

reaction mixture was allowed to cool and then the absorbance read at 540 nm. One unit of

enzyme activity was defined as the amount of enzyme that catalyzed the release of one

micromole of glucose per minute

50

2.2.10.2 Endoglucanase (CMCase) assay

The reaction mixture containing 0.2 ml CMC (2%) in 0.05 M sodium acetate buffer pH 5.5

and 0.5 ml enzyme solution was incubated for 30 min at 50 ºC. Then, 1 ml of DNS reagent

was added and the reaction was stopped by boiling the mixture in a boiling water bath for 10

mins. The mixture was made up to 4 ml with 1 ml of Rochelle salt solution. The reaction

mixture was allowed to cool and then the absorbance read at 540 nm. One unit of enzyme

activity was defined as the amount of enzyme that catalyzes the release of one micromole of

glucose per minute

2.2.10.3 Total cellulase (Filterpaperase) assay

The reaction mixture containing 50 mg (1 cm x 6 cm) filter paper in 0.05 M sodium acetate

buffer of pH 5.5 and 0.5 ml enzyme solution was incubated for 1 hour at 50 ºC and 1 ml of

DNS reagent was added and the reaction was stopped by boiling the mixture in a boiling

water bath for 10 min. The volume was made up to 4 ml with Rochelle salt solution. The

reaction mixture was allowed to cool and then the absorbance read at 540 nm. One unit of

enzyme activity was defined as the amount of enzyme that catalyzed the release of one

micromole of glucose per minute.

2.2.11 Partial Purification of Protein

2.2.11.1 Determination of Percentage Ammonium Sulphate Saturation Suitable for

Cellulase Precipitation

Nine test tubes were used to form an ammonium sulphate precipitation profile. Cellulases

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

intervals of 10% in each test tube. The ammonium sulphate-crude enzyme solutions were

allowed to stand at cold temperature of 4oC for 30 hr till the supernatant could be gently

decanted off. The test tubes were centrifuged at 3500 rpm for 10 mins. Precipitates from the

individual percentage ammonium sulphate saturations were re-dissolved, respectively, in

equal volumes of 0.05 M acetate buffer pH 5.0. Cellulase activities of the precipitates were

51

assayed to determine the percentage ammonium sulphate saturation that precipitated enzyme

with maximum activity.

2.2.11.2 Ammonium Sulphate Precipitation of Cellulase

A known value, 750 ml of crude enzyme filtrate was used in the process. Also, 80% and 70%

ammonium sulphate saturation (for Aspergillus fumigatus and Aspergillus flavus,

respectively) were found suitable for mass precipitation of the enzymes from the fungal

isolates. Ammonium sulphate precipitation (at 80% and 70% saturation) was carried out by

dissolving gently 516 g and 436 g of the salt in the filtrates and stirring gently till the salt was

completely dissolved as seen in section 2.2.11.1. The precipitate was re-dissolved in 55 ml of

0.05 M acetate buffer pH 5.0 after centrifugation and then kept under cold condition for

further studies.

2.2.11.3 Gel filtration

The enzyme was introduced onto Sephadex G-25 packed column (2.6 × 61.50 cm) pre-

equilibrated with 0.05M Na-acetate buffer, pH 5.5. The protein was eluted with 0.05 M Na-

Acetate buffer, pH 5.5. The fractions with high cellulase activity were pooled together.

Cellulase activity from each of the eluted fraction were monitored at wave length of 540 nm

and protein absorbance read at 280 nm.

2.2.12 Studies on Partially Purified Enzyme

2.2.12.1 Enzyme Progress Curve

The enzyme was incubated with its substrate (filter paper) at 50 C and activity was assayed

at 5, 10 , 20, 30, 40, 50, 60, 70, 90 and 120 min respectively.

52

2.2.12.2 Effect of pH on Total Cellulase Activity

The optimum pH for enzyme activity was determined using 0.05 M sodium acetate buffer pH

4.0 - 5.5, phosphate buffer pH 6.0 - 7.5 and Tris-HCl buffer pH 8.0 – 9.0 at intervals of 0.5. A

known quantity, 0.2 ml of partially purified enzyme was dispersed into 0.8 ml of buffer of

different pHs into which 50 mg (1cm x 6cm) had been added to. Total Cellulase activity was

assayed as seen in section 2.2.10.3

2.2.12.3 Effect of Temperature on Total Cellulase Activity

The optimum temperature was determined by incubating the enzyme with 50mg filter paper

at 25-70oC, interval of 5

oC for 1 hour and at pH 5.5. The activity was then assayed using the

method described in 2.2.10.3

2.2.12.4 Determination of kinetic parameters

Kinetic parameters were determined using different concentrations of filter paper and

cellobiose. The Vmax and Km values of the enzymes were calculated using the Lineweaver-

Burke double reciprocal plot of 1/V against 1/[S].

53

CHAPTER 3

RESULTS

3.1 Incubation Period (Pilot Study)

Five (5) days of incubation produced the highest total cellulase activity at 2.97 and 3.87 U/ml

for enzymes produced by A. flavus and A. fumigatus respectively.

Fig. 9: Effect of incubation period on cellulase production

54

3.2 Studies on crude enzyme

3.2.1 Protein concentration of crude enzyme

Protein concentration of crude enzyme produced by A. flavus and A. fumigatus were found to

be 4.03 and 4.17 mg/ml respectively.

Fig. 10: Protein concentration of crude enzyme produced by the microoganisms

55

3.2.2 Cellulase activity of crude enzymes.

Cellulolytic activities of enzymes from the two fungi are shown below.

The total cellulase activity of 750 ml of crude enzyme produced by each microorganism was

found to be 3.04 and 4.16 U/ml for A. flavus and A. fumigatus respectively. Glucanase

activity was 2.86 and 4.84 U/ml for enzymes produced by A. flavus and A. fumigatus

respectively. Cellobiase activity was observed to be highest for the three enzyme assays. A.

flavus had an activity of 10.09 U/ml while A. fumigatus had an activity of 10.16 U/ml.

Fig. 11: Total cellulase, glucanase and cellobiase activities of crude enzyme

56

.3 Ammonium sulphate precipitation profile of cellulases

As in Fig. 12, 70% and 80% ammonium sulphate had highest cellulase activities at 3.55 and

3.3 U/ml for A. flavus and A. fumigatus respectively. Hence, 70% and 80% were chosen for

the precipitation of the enzymes from the two microorganisms.

Fig. 12: Ammonium sulphate precipitation profiles for celluluases isolated from A. flavus and

A. fumigatus.

57

3.4 Gel filtration (Elution profile of cellulase enzymes)

For cellulase isolated from A. flavus, two prominent peaks (A and B) were observed with

activities of 3.99 U/ml and 3.64 U/ml respectively as shown in Fig. 13a.

Fig. 13a: Gel elution profile of proteins produced by A. flavus.

58

For cellulases isolated from A. fumigatus, two prominent peaks C and D were also identified

with activities of 2.94 and 3.11 U/ml as show in Fig. 13b.

Fig. 13b: Gel elution profile of proteins produced by A. fumigatus

59

3.5 Summary of purification steps

Table 6: Summary of purification steps of cellulase from A. flavus

Purification step Volume

(ml)

Protein

conc.

(mg/ml)

Activity

(U/ml)

Spec.

Activity

(U/mg)

Total

Activity

(U)

Purification

fold

% Yield

Crude Enzyme

Filtrate

750 4.03 3.04 0.75 2280 1 100

% (NH4)2SO4

Precipitation

55 4.1 5.1 1.24 280.5 1.65 12.3

Gel filtrate A 20 2.75 3.99 1.43 59.8 1.91 2.62

Gel fitrate B 20 2.28 3.64 1.59 52.8 2.12 2.34

Table 6 shows the summary of the purification steps of cellulase from A. flavus. An initial

volume of 750 ml of crude enzyme extract yielded a protein concentration of 4.03 mg/ml

with an activity of 3.04 U/ml. and specific activity of 0.75 U/mg. Ammonium sulphate

precipitation re-dissolved in 55 ml of buffer yielded a protein concentration of 4.1 mg with

activity of 5.1 U/ml and specific activity of 1.24 U/mg. Gel filtrates of 20 ml each for the 2

isoforms of cellulase A and B yielded protein concentrations of 2.75 mg/ml and 2.28 mg/ml

for isoforms A and B respectively. The activities of the filtrates were 3.99 and 3.64 U/ml for

the corresponding isoforms. Specific activities were 1.43 and 1.59 U/ml respectively. The

overall percentage yield fell from 100% to 2.62 and 2.34% after gel filtration. Total activity

decreased from 2280 U to 59.8 and 52.8 U respectively after gel filtration.

60

Table 7: Summary of purification steps of cellulase from A. fumigatus

Purification step Volume

(ml)

Protein

conc.

(mg/ml)

Activity

(U/ml)

Spec.

Activity

(U/mg)

Total

Activity

(U)

Purification

fold

% Yield

Crude Enzyme

Filtrate

750 4.17 4.16 1.0 3120 1 100

% (NH4)2SO4

Precipitation

55 4.79 5.9 1.23 324.5 1.23 10.4

Gel filtrate A 20 2.16 2.94 1.36 58.8 1.36 1.88

Gel fitrate B 20 2.09 3.11 1.49 62.2 1.49 1.99

Table 7 shows the summary of the purification steps of cellulase from A. fumigatus. An initial

volume of 750 ml of crude enzyme extract yielded a protein concentration of 4.17 mg/ml

with an activity of 4.16 U/ml. and specific activity of 1.0 U/mg. Ammonium sulphate

precipitation re-dissolved in 55 ml of buffer yielded a protein concentration of 4.79 mg with

activity of 5.9 U/ml and specific activity of 1.23 U/mg. Gel filtrates of 20 ml each for the two

isoforms of cellulase A and B yielded protein concentrations of 2.16 mg/ml and 2.09 mg/ml

for isoforms A and B respectively. The activities of the filtrates were 2.94 and 3.11 U/ml for

the corresponding isoforms. Specific activities were 1.36 and 1.49 U/ml respectively. The

overall percentage yield fell from 100% to 1.88 and 1.99% after gel filtration. Total activity

reduced from 3120 U to 58.8 and 62.2 U respectively after gel filtration.

61

3.6 Changes in Protein Concentration of Partially Purified Enzymes

3.6.1 Changes in protein concentration of enzymes from a. flavus.

It was observed in Fig. 14 that the protein concentration increased from 4.03 to 4.1 mg/ml

after ammonium sulphate precipitation and decreased to 2.75 mg/ml and 2.28 mg/ml

(corresponding to isoforms A and B) after gel filtration.

Fig. 14: Changes in protein concentration after partial purification of cellulases from A. flavus

62

3.6.2 Changes in protein concentration of enzymes from A. fumigatus

As shown in Fig. 15, for A. fumigatus, the protein concentration increased from 4.17 to 4.79

mg/ml after ammonium sulphate precipitation and decreased to 2.16 and 2.09 mg/ml for

isoforms C and D respectively after gel filtration.

Fig. 15: Changes in protein concentration after partial purification of cellulases from A.

fumigatus

63

3.7 Changes in Total Cellulase Activity of Partially Purified Enzymes

3.7.1 Changes in total cellulase activity of partially purified enzymes from a. flavus

The activity of the enzyme increased 3.04 to 5.1 U/ml after ammonium sulphate precipitation

as shown in Fig 16. There was a reduction in activity after gel filtration to 3.99 and 3.64 U/ml

for the 2 isoforms A and B present.

Fig. 16: Changes in total cellulase activity after partial purification of enzymes from A. flavus

64

3.7.2 Changes in Total Cellulase Activity of Partially Purified Enzymes from A.

fumigatus

Enzyme activity increased from 4.16 to 5.9 U/ml after ammonium sulphate precipitation as

depicted in Fig. 17. However, it reduced to 2.94 U/ml and 3.11 U/ml (for isoforms C and D

respectively) after gel filtration.

Fig. 17: Changes in total cellulase activity after partial purification of enzymes from A.

fumigatus

65

3.8 Changes in Specific Activities of Partially Purified Enzymes

3.8.1 Changes in specific activities of partially purified enzymes from A. flavus

Specific activity of crude enzyme was found to be 0.75 U/mg as observed in Fig 18. This

value increased to 1.24 U/mg after ammonium sulphate precipitation. It further increased to

1.43 U/mg and 1.59 U/mg (corresponding to the two isoforms) after gel filtration.

Fig. 18: Changes in specific activity after partial purification of cellulases from A. flavus.

66

3.8.2 Specific activities of partially purified enzymes from A. fumigatus

Fig. 19 shows specific activity of 1.23 U/mg for crude enzyme after ammonium sulphate

precipitation. This value increased to 1.36 U/mg and 1.49 U/mg corresponding to the two

isoforms present after gel filtration.

Fig. 19: Changes in specific activity after partial purification of cellulases from A. fumigatus

67

3.9 Enzyme Characterization

3.9.1 Enzyme Progress Curve

3.9.1.1 Enzyme progress curve of partially purified enzymes from A. flavus

The A isoform exhibited maximum activity after 60 min of incubation as shown in Fig. 20

The B isoform, however, had maximum activity after 50 min of incubation.

Fig. 20: Progress curve of cellulases isolated from A. flavus

68

3.9.1.2 Enzyme progress curve of partially purified enzymes from A. fumigatus

Fig. 21 shows the progress curve of partially purified enzyme. For cellulase isolated from A.

fumigatus, the C isoform had highest activity after 70 min of incubation while the D isoform

had highest activity after 60 minutes of incubation.

Fig. 21: Progress curve of cellulases isolated from A. fumigatus

69

3.9.2 Effect of pH Change on Cellulase Activity

3.9.2.1 Effect of pH change on cellulase produced by A. flavus

Fig. 22 shows that the enzymes had optimum activities of 3.31 and 3.53 U/ml corresponding

to the two isoforms A and B of the enzymes at pHs of 6.5 and 7.0 respectively. Further

increase in pH led to a decline in enzyme activity.

Fig. 22: Effect of pH on cellulases produced by A. flavus.

70

3.9.2.2 Effect of pH Change on Cellulase Produced by A. fumigatus

For cellulase isolated from A. fumigatus, the enzyme exhibited highest activities of 3.07 U/ml

and 3.42 U/ml corresponding to the isoforms C and D at a pH of 5 for both forms as shown in

Fig. 23

Fig. 23: Effect of pH on cellulases produced by A. fumigatus

71

3.9.3 Effect of Temperature Change on Cellulase Activity.

3.9.3.1 Effect of temperature on cellulase produced by A. flavus

For cellulase isolated from A. flavus as shown in Fig. 24, maximum activity was observed at

a temperature of 50⁰C for both isoforms of the enzyme when assayed at pHs of 6.5 and 7.0

Fig. 24: Effect of temperature on cellulases produced by A. flavus

72

3.9.3.2 Effect of temperature on cellulase produced by A. fumigatus

Cellulase produced by A. fumigatus had optimum activity at 55 ⁰C for both isoforms when

assayed at a pH of 5.5 for both isoforms as shown in Fig. 25.

Fig. 25: Effect of temperature on cellulases produced by A. fumigatus

73

3.9.4 Determination of Kinetic Parameters (Using Filter paper as Substrate)

Kinetics parameter Vmax and Km determined from lineweaver burk plots (using filter paper as

substrate)

3.9.4.1 Determination of Kinetic parameters for enzymes of A. flavus

Isoform A of cellulase isolated from A. flavus had Vmax and Km values of 142.9 µmole/min

and 59.02 mg respectively as shown in Fig. 26.

Fig. 26: Lineweaver-Burke plot of cellulase A from A. flavus using filter paper as substrate

74

Isoform B of cellulase isolated from A. flavus had Vmax and Km values of 166.7 umole/min

and 47.67 mg respectively as shown in Fig. 27

Fig. 27: Lineweaver-Burke plot of cellulase B from A. flavus using filter paper as substrate

75

3.9.4.2 Determination of Kinetic parameters for enzymes of A. fumigatus

Isoform C of cellulase isolated from A. fumigatus had Vmax and Km values of 128.21

µmole/min and 27.82 mg respectively as shown in Fig. 28

Fig. 28: Lineweaver-Burke plot of cellulase C from A. fumigatus using filter paper as

substrate

76

Isoform D of cellulase isolated from A. fumigatus had Vmax and Km values of 90.91

µmole/min and 32 mg respectively as shown in Fig. 29

Fig. 29: Lineweaver-Burke plot of cellulase D from A. fumigatus using filter as substrate

77

3.9.5 Determination of Kinetic Parameters (Using Cellobiose as Substrate)

Kinetics parameter Vmax and Km determined from lineweaver burk plots (using cellobiose as

substrate)

3.9.5.1 Determination of Kinetic parameters for enzymes of A. flavus

Isoform A of cellulase isolated from A. flavus had Vmax and Km values of 588.2 µmole/min

and 7.7 mM as shown in Fig. 30

Fig. 30: Lineweaver-Burke plot of cellulase A from A. flavus using cellobiose as substrate

78

Isoform B of cellulase isolated from A. flavus had Vmax and Km values of 476.2 µmole/min

and 3.33 mM as shown in Fig. 31

Fig. 31: Lineweaver-Burke plot of cellulase B from A. flavus using cellobiose as substrate

79

3.9.5.2 Determination of Kinetic parameters for enzymes of A. fumigatus

Isoform C of cellulase isolated from A. fumigatus had Vmax and Km values of 833.3

µmole/min and 11.1 mM as shown in Fig. 32

Fig. 32: Lineweaver-Burke plot of cellulase C from A. fumigatus using cellobiose as substrate

80

Isoform D of cellulase isolated from A. fumigatus had Vmax and Km values of 666.67

µmole/min and 9.1 mM as shown in Fig. 33

Fig. 33: Lineweaver-Burke plot of cellulase D from A. fumigatus using cellobiose as substrate

81

Table 8: Characterization of Cellulase

Properties A.flavus (Isoform

A)

A.flavus (Isoform

B)

A.fumigatus (Isoform

C)

A.fumigatus (Isoform

D)

pH 6.5 7.0 5.0 5.0

Temperature (◦C) 50 50 55 55

Vmax (µmole/min)

(Filter paper)

142.9 166.7 128.21 90.91

Km (mg)

(Filter paper)

59.02 47.67 27.82 32

Vmax (µmole/min)

(Cellobiose)

588.2 476.2 833.3 666.67

Km (mM)

(Cellobiose)

7.7 3.33 11.1 9.1

82

CHAPTER 4

DISCUSSION

This study deals with the production, partial purification and characterization of cellulolytic

enzymes produced by two species of the Aspergillus genus- flavus and fumigatus. Studies

were undertaken to optimize the production of extracellular cellulases by these organisms.

The enzymes were then isolated and purified and their characteristics were studied.

The microbes when cultivated in the presence of breadfruit hulls as carbon source secreted

cellulolytic enzymes which exhibited highest activity after 5 days of incubation. Both species

of Aspergillus used in this work were grown over a 7 day period. The growth of fungi on a

medium is usually affected by several factors which include: nutrients, temperature, light,

aeration, pH, and water activity (Raim, 1998). Fungi generally exhibit four basic growth

stages: the spore, which is the dormant phase, followed by the spore germination or lag

phase, the growth or hyphae phase and the spore formation phase (Carlile et al., 2001). It is

expected that as the fungi grow, they secrete more proteins into the containing medium until

growth approaches the lag phase (Raim, 1998). Other researchers have previously reported

different days for obtaining maximum activity during incubation. Nwobodo and Okochi

(2011) reported maximum activity after 3 days of incubation using Aspergillus niger grown

on sawdust. Charitha and Kumar (2012) also observed highest activity after 7 days of

incubation using A. niger grown with waste paper. However, Adekunle et al (2012) reported

maximum enzyme activity at 6 day of growth when A. niger was grown on rice as carbon

source. Recently, Das et al (2013) reported maximum activity after 3 days growth when A.

fumigatus ABK9 was cultivated under submerged fermentation condition. Incubation time of

a fermentation experiment has a direct relationship with the production of extracellular

enzymes; however, microbial growth and enzyme production plateaus as the growth

approaches the lag or stationary phase. (Raim, 1998). The variation in optimum activities can

be attributed to the different environmental conditions which affect the growth of fungi (Das

et al., 2013). The findings in this work suggest that in order to obtain a commercial quantity

of cellulases (in an industrial setting) from A. flavus and A. fumigatus, especially under

submerged conditions using breadfruit hulls as carbon source, the crude enzymes should be

harvested on the 5th

day of growth. However, fresh pilot studies should be carried out when

83

other cellulosic substrates are used as carbon source or when other cellulase producing

microorganisms are used instead of those used in this work

Ammonium sulphate was used to salt out the proteins. Ammonium sulphate precipitation is

one of the most commonly used methods for protein purification from a solution. The

principle behind ammonium sulphate precipitation is the altering of the solubility of proteins

in the presence of a salt (Green and Hughes, 1955). In solution, proteins form hydrogen

bonds with water molecules through their exposed polar and ionic groups (Mitchinson and

Pain, 1986). When high concentrations of small, highly charged ions such as ammonium

sulphate are added, these groups compete with the proteins to bind to the water molecules.

This removes the water molecules from the protein and decreases its solubility, resulting in

precipitation (Green and Hughes, 1955; Harriette and Charles, 1913). Critical factors that

affect the concentration at which a particular protein will precipitate include: the number and

position of polar groups, molecular weight of the protein, pH of the solution, and temperature

at which the precipitation is performed. Protein concentration and activity were higher for the

precipitated enzymes when compared with that of the crude form with the greater activity

being attributed to the increased concentration of proteins in solution. The proteins were

purified by precipitating with ammonium sulphate at 70% and 80% for A. flavus and A.

fumigatus respectively as shown in the result. Charitha and Kumar (2012) reported 80%

saturation for enzymes of A. niger fermented with paper and timber sawmill industrial waste.

Bakare et al., (2005) reported maximum precipitation of proteins at 90% saturation for

proteins from mutants of Pseudomonas fluorescens. Shanmu et al., (2012) reported 80%

saturation for cellulases produced by bacteria that were grown on cow dung Guo et al.,

(2013) obtained 60% saturation for cellulase from a commercial enzyme preparation. It is

important to note that in all these precipitation experiments including the one performed in

this work, the supernatant had almost no activity and was therefore discarded. From the

results obtained from literature and from this work, cellulases tend to precipitate out at

increasing concentrations of ammonium sulphate from 60% to 90%.

On further subjection to gel filtration, two prominent peaks were observed for both cellulases

produced by both fungi indicating two forms or isoforms of the enzymes as seen in the

results. Multiplicity of cellulases produced by microbes is a general phenomenon (Marsden

and Gray, 1986; Brown and Gritzali, 1984). Both microbes showed at least two forms of

84

these enzymes. Heterogeneity of cellulases may be attributed to one or more of the following:

posttranslational modification (glycosylation, proteolysis, phosphorylation, acetylation etc.),

multiple gene expression (Agelos and Panagiotis, 1991)

After the purification, it was observed that the specific activity of the enzymes increased after

each step as can be deduced from the result. For enzymes from A. flavus, the initial specific

activity was 0.75 U/mg. After ammonium sulphate precipitation, this value increased to 1.24

U/mg and further to 1.43 U/mg and 1.59 U/mg for isoforms A and B respectively after gel

filtration. For enzymes of A. fumigatus on the other hand, the initial specific activity was 1.0

U/mg. After ammonium sulphate precipitation, this value increased to 1.23U/mg and further

to 1.36 U/mg and 1.49 U/mg for isoforms C and D respectively after gel filtration. Specific

activity is a measure of enzyme purity. This value increases as an enzyme preparation

becomes purer, since the amount of protein (mg) is typically less, but the rate of reaction

stays the same or increases due to reduced interference or removal of inhibitors (Nelson and

Cox, 2000).

The enzyme was characterized and effects of incubation time, pH and temperature on enzyme

activity were observed. Kinetic parameters were also studied. Cellulase isolated from A.

flavus had highest activities after 60 min and 50 min (corresponding to forms A and B) of

incubation as shown in the results. Those of A. fumigatus were observed to have highest

activity after 70 minutes and 60 minutes for forms C and D respectively of incubation as

shown in the results. The longer the enzyme was incubated with its substrate, the greater the

amount of product formed up to a point when there is a decline with time. This was probably

due to the combined effects of substrate utilization and product accumulation (Duggleby,

1986).

The effect of pH on the total cellulase activity of both enzymes produced by the two species

of Aspergillus was studied. A. flavus exhibited highest activities at pHs of 6.5 and 7.0

corresponding to the isoforms A and B identified after gel filtration. Enzymes produced by A.

fumigatus on the other hand had maximum activity at a pH of 5.5 for both forms of the

enzymes. On further increase in pH, activity fell probably due to changes in total net charge

of the enzymes. This effect of pH on charge distribution on the ionizable groups interrupts the

tertiary structure of the enzyme and thus causes its denaturation. Saraswati et al. (2012)

85

reported a pH of 7.0 for cellulase produced by bacillus isolated from cow dung. Sunita and

Sumit (2012) also reported an optimum pH of 7 for cellulase produced by Trichoderma viride

using sawdust and coir waste as carbon sources. Mawadza et al (2000) reported highest

activity at a pH range of 5-7 for bacillus sp CH43 strain and a pH optimum 5-6.5 for Bacillus

sp HR68 strain with the enzyme having highest activity at 6.5. The results are in line with the

observed pHs of 5.5, 6.5 and 7.0 reported in this work using 2 species of the Aspergillus

genus. The results obtained indicate that optimum pH is around neutral and slightly acidic

pH. Thus, cellulase enzyme obtained from these organisms will best be utilized for industrial

applications at neutral or slightly acidic pH.

The temperature optima of the total cellulase activity for both isoforms A and B secreted by

A. flavus was 50 ºC while the highest total cellulase activity was obtained at 55 ºC for both

forms C and D isolated from A. fumigatus. Enzyme-catalyzed reactions tend to be slower at

temperatures below the optimum. They then tend to go faster with increasing temperature but

only until a temperature optimum is reached. Above the optimum temperature, the kinetic

energy of the enzyme increases to the extent that the weak intermolecular attractions that

maintain the shape of proteins are broken and the enzyme molecule is disrupted - the enzyme

becomes denatured. Changing the shape of the enzyme results in less efficient binding of the

substrate (reactants) resulting in a significant decrease in enzyme activity (Duggleby, 1986).

Mawadza et al (2000) had earlier reported maximum activity at 45 ºC for total cellulase

activity of the enzymes produced by fusarium and penicilium. A temperature of 60 ºC was

also observed in same work by Mawadza et al (2000) to give maximum activity for enzymes

produced by Aspergillus sp. In a study carried out by Immanuel et al. (2006) the enzyme

produced by A. niger had less activity at 20 ºC but on further increase to 50 ºC, maximum

activity was obtained while maximum enzyme activity was recorded at 50 ºC for A.

fumigatus. Rahna et al. (2012) reported maximum activity at 50 ºC for total cellulase activity

of enzymes produced by pseudomonas sp using salvinia as substrate. These temperature

optima are relatively in agreement with those observed in this study. The implication of these

findings is that cellulolytic enzymes would best be utilized at 50 ºC for converting cellulosic

biomass into desirable products.

The effect of substrate concentration on the total cellulase activity of the enzymes was

studied. Thereafter, the kinetic constants were determined. With fixed enzyme concentration,

86

an increase in substrate concentration resulted in increase in enzyme activity until a saturation

point was reached at which further increases in substrate concentration did not result in

activity. This could be due to the formation of unreactive complexes formed between the

enzyme and substrate. Also to be considered is the fact that when substrate molecules are in

high concentration around the enzyme, they may bind to other sites other than the enzyme

active site or alternatively crowd the active site. Total cellulase activity, using filter paper as

substate, had maximum velocity of 142.9 and 166.7 µmol/min for both isoforms A and B of

the enzyme secreted by A. flavus while Km values of 59.02 and 47.67 mg were observed for

the corresponding isoforms. On the other hand, for enzymes produced by A. fumigatus, they

showed maximum velocity of 128.21 and 90.91 µmol/min for the 2 isoforms C and D

respectively. The Km values were 27.82 and 32 mg respectively. Using cellobiose as

substrate, Vmax values were 588.2, 476.2, 833.3 and 666.67 µmol/min for isoforms A, B, C

and D respectively. Km values 7.7, 3.3, 11.1 and 9.1 mM were obtained for isoforms A, B, C

and D respectively. The Km values serve to indicate the substrate concentration required to

achieve half the maximum initial reaction velocity. Km therefore measures the relative

affinity an enzyme has for its substrate. A smaller Km denotes a greater affinity that an

enzyme has for its substrate. From the results obtained, enzymes of A. fumigatus have a

greater affinity for cellulose when compared with those of A. flavus and are thus better

applied for an industrial saccharifying process, the reason being that the enzyme will act at a

more or less constant rate, regardless of variations in concentrations of substrate.

4.2 CONCLUSION

The results indicate possibility of using agricultural waste such as breadfruit hulls to induce

cellulase production in saprophytic microbial fungi, thus converting waste into wealth in the

form of enzymes of industrial importance. Most of the work done on the genus Aspergillus

have been on A. niger which is still currently the fungi of choice to produce these enzymes.

This work however provides options that could be considered for production of cellulases

using raw waste materials that could be easily obtained from the environment.

87

4.3 SUGGESTIONS FOR FURTHER RESEARCH

Based on the findings in this work, the following suggestions are made

1. Studies on the other physiological properties of the cellulase enzymes such as thermal

stability and pH stability should be conducted to understand their effects on the

enzyme activity.

2. Further purification of the enzymes using ion exchange chromatography and gel

electrophoresis should be conducted and Sodium Dodecyl Sulphate Polyacrilamide

Gel Electrophoresis (SDS PAGE) used to find out the molecular weight of these

enzymes.

3. X-ray crystallography and NMR should be used to elucidate the active sites of these

enzymes as well as their tertiary structures.

88

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101

APPENDICES

1.0 Preparation of Buffers

The standard buffers used in study were pH 4.0, pH 7.0 and pH 9.2. These buffers were used

to standardize the pH meter. The working buffers were prepared as thus: 0.05 M sodium

acetate and 0.05 M Tris-HCl buffers were prepared by dissolving 4.10 g sodium acetate salt

and 6.01 g Tris base, respectively in 1000ml of distilled water and stirred with a magnetic

stirrer till a homogenous solution was formed. The solutions were titrated againt acetic acid

and HCl, respectively till the required pHs were obtained. Also 0.05 M phosphate buffer was

prepared by dissolving 7.10g disodium hydrogen phosphate salt in 1000 ml stirred as for

sodium acetate and phosphate buffers and then titrated against the solution of its conjugate

acid, sodium dihydrogen phosphate till the required pHs were obtained.

1.1 Preparation of Dinitrosalicylic Acid (DNS) Reagent

A modification of DNS reagent method of Miller (1959) as contained in wang et al. (1997)

was used in the assay. The reagent contains 44 mM dinitrosalicylic acid, 4 mM sodium

sulphite, and 375 mM sodium hydroxide.

1.2 Preparation of Cellobiose

15 mM Cellobiose was prepared by dissolving 5.13 g cellobiose in distilled water and made

up to 1 litre

1.3 Preparation of Carboxymethyl Cellulose (CMC)

2% Carboxymethyl Cellulose was prepared by dissolving 2 g of carboxymethyl cellulose in

distilled water and made up to 100 ml.

102

1.4 Preparation of 50mM glucose

50 mM solution of glucose was prepared by weighing 0.9 g of industrial grade glucose and

dissolving in 100 ml of distilled water.

1.5 Calibration Curve for Glucose

A method described by Miller 1959 with little modifications by Wang et al., 1997 was used.

Ten test tubes were arranged in duplicate containing 0.0-1.0 ml of 50 mM glucose. Each tube

was made up to 1ml using 0.05 M sodium acetate buffer of pH 5.5. 1 ml DNS reagent was

added to each of the tubes and placed in boiling water bath for 10 mins. The tubes were then

removed and allowed to cool to room temperature. Na-K tartarate was added to the different

tubes to stabilize the colour, after which the absorbance was read at 540 nm. The

concentration of reducing sugar in each of them was calculated using the formula

“C1V1=C2V2” where:

C1= initial concentration of reducing sugar (mM)

V1= initial volume of the 50mM preparation measured into the tubes

C2= final concentration of reducing sugar (mM)

V2= final volume of the preparation measured in the tube

Using the value obtained from the table described above, the plot of optical density against

concentration was constructed.

1.4 Preparation of the Component Reagents For Protein Determination

Solution A: An alkaline sodium carbonate (Na2CO3) was prepared by dissolving 2 g of

Na2CO3 in 100ml of 0.1 M NaOH (0.4g of sodium hydroxide pellets were dissolved in 100

ml of distilled water).

Solution B: A copper tetraoxosulphate IV - sodium potassium tartarate solution was prepared

by dissolving 0.5 g of CuSO4 in 1 g of sodium potassium tatarate, all in 100 ml of distilled

water. It was prepared fresh by mixing stock solution, and so was done whenever required.

Solution C: Folin-Ciocalteau phenol reagent was made by diluting the commercial reagent

with water in a ratio of 1:1 on the day of use.

Solution D: Standard protein (Bovine Serum Albumin, BSA) solution.

103

Solution E: Freshly prepared alkaline solution was made by mixing 50 ml of solutions A and

1 ml of solution B.

1.6 Preparation of 2 mg/ml Bovine Serum Albumin (BSA) Standard Protein

0.2 g of BSA was dissolved in 100 ml of distilled water and then used as a protein stock

solution.

104

Protein Standard Curve, Using Bovine Serum Albumin (BSA)

105

Glucose Standard Curve

106