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OKOYE, IFEANYI GODWIN
(PG/M.Sc/08/48333)
PG/M. Sc/09/51723
PRODUCTION AND PARTIAL CHARACTERIZATION OF CELLULASES FROM ASPERGILLUS
FUMIGATUS USING TWO DISTINCT PARTS OF CORN COB AS CARBON SOURCES
BIOLOGICAL SCIENCES
A THESIS SUBMITTED TO THE DEPARTMENT OF BIOCHEMISTRY, FACULTY OF
BIOLOGICAL SCIENCES, UNIVERSITY OF NIGERIA NSUKKA
Webmaster
Digitally Signed by Webmaster’s Name
DN : CN = Webmaster’s name O= University of Nigeria, Nsukka
OU = Innovation Centre
MAY, 2012
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PRODUCTION AND PARTIAL CHARACTERIZATION OF CELLULASES FROM ASPERGILLUS
FUMIGATUS USING TWO DISTINCT PARTS OF CORN COB AS CARBON SOURCES
A DISSERTATION SUBMITTED IN PARTIAL FULFILMENT OF THE
REQUIREMENTS FOR THE AWARD OF THE DEGREE OF MASTER OF
SCIENCE (M.Sc) IN BIOCHEMISTRY (INDUSTRIAL BIOCHEMISTRY
AND BIOTECHNOLOGY), UNIVERSITY OF NIGERIA, NSUKKA
BY
OKOYE, IFEANYI GODWIN
(PG/M.Sc/08/48333)
DEPARTMENT OF BIOCHEMISTRY
UNIVERSITY OF NIGERIA, NSUKKA
MAY, 2012
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CERTIFICATION
Okoye, Ifeanyi Godwin, a postgraduate student of the Department of Biochemistry
with Registration Number; PG/M.Sc/08/48333, has satisfactorily completed the
requirements of research work for the degree of Master of Science (M.Sc) in Industrial
Biochemistry and Biotechnology. The work embodied in this dissertation is original
and has not been submitted in part or full for any other diploma or degree of this or any
other university.
………………………… …………………………
Prof. F. C. Chilaka Prof. L. U. S. Ezeanyika
(Supervisor) (Head of Department)
…………………………………
External Examiner
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DEDICATION
This work is dedicated to God whose glory it is to conceal things but who also honours
dedicated scientists who do His bidding, by enabling them search out and unravel the
concealed things. And to my elder brother, Mr. Christopher I. Okoye, who bore the
burden of my sponsorship in this programme.
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ACKNOWLEDGEMENT
I am deeply grateful to God who saw me through starting from the genesis to the
completion of this work. He not only gave me strength in times of discouragement but also bathed
me with favour and mercy as this programme lasted. He provided sponsors for me and gave me
one of the best supervisors in the Department of Biochemistry. I am indebted to my supervisor,
Prof. F.C. Chilaka, an emblem of erudition blended with thoroughness and liberality. You believe
in training not only the head but also the hands. I have learnt from you why true scientists have to
do it over and over. Hence, the saying comes true that it is not how long that matters, but how
well. Thank you for teaching me the difference between true happiness and what people call
achievement. Mr. D.A. Yoosu provided me with corn cobs from the National Cereals Research
Institute, Yandev Benue, State. Thank you. My gratitude is extended to Dr. C.U. Anyanwu in
whose laboratory I carried out the microbiological aspect of this work. You gave me an unlimited
access to your laboratory in response to my supervisor’s request. To Mr. Baturh Yarkwan, my
friend and senior colleague in biotechnology research, I give a million thanks. Just one night with
you in Dr. Anyanwu’s laboratory made a great difference. It was there you painstakingly taught
me the nitty-gritty of microbial isolation and preservation. Your immense contribution to the
entire work is well appreciated. Worthy of mention is the contribution of Dr. E.A. Eze. He helped
in partial identification of the isolated fungi and his unassuming nature is inspiring. May God
bless him.
My special thanks go to all the lecturers in the Department of Biochemistry especially those
who played direct roles in my training. From their wealth of knowledge I have benefited
immensely. Of special note are the following: the Head, Department of Biochemistry, Prof. L.U.S.
Ezeanyika whose pragmatic, principled and humble leadership style has helped in the timely
completion of this academic work. Prof. I.N.E. Onwurah whose love for righteous living has
encouraged me in no small measure. May God bless Prof. O. Obidoa, Prof. O.F.C. Nwodo, Prof.
P.N. Uzoegwu, Prof. O.U. Njoku, Prof. E.O Alumanah, Dr. V.N. Ogugua, Dr. B.C. Nwanguma,
Dr. S.O.O. Eze, Dr. H.A. Onwubiko, Dr. Parker E. Joshua and Dr C.S. Ubani whose
contributions at one time or the other especially during post-graduate seminars helped in fine-
tuning this work. Dr. Parker E. Joshua and Dr C. S. Ubani deserve my special appreciation
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because they have been particularly helpful in very many ways I cannot fully express. You have a
sure reward.
The Chief Technologist, Mrs M.N. Nwachukwu encouraged me a lot by giving me every
available equipment needed for my work. Mr. Jude Chime is also worthy of appreciation. He was
very co-operative as regards my use of the instrument room.
My very close friend and co-researcher, Valentine Ogwuche has made great impact on my
life. My meetng you was divinely orchestrated. Ben Onah, Nonso Nsude, Aghogho A., Nchedo
O., Ejike Orji and Fortunatus Ezebuo are not only close to me but are also genuine friends that
mean well. I have learnt a great deal from all of you including my senior colleague, Obinna Oje.
My beloved room mate, Kalu Chukwudi, my esteemed course mates and my numerous Christian
brethren in GSF have cared for me in very many ways. May God reward you all.
My success story would not be complete without the roles played by those of my own
flesh and blood. My mother, Mrs L.C. Okoye was always there for me, showering me with
motherly love and care. Mum, you are a rare gift! My uncle and his wife, Chief Pharm. & Mrs
Tochukwu Maduagwuna were really very supportive in many ways. May you not lose your
reward for the kindness you have shown to me. My elder brother, Christopher I. Okoye bore the
burden of my sponsorship in this programme. It’s amazing how you shared your monthly salary
with me for more than two years. I am deeply grateful. Aunty Rose showed great concern for my
welfare as this work lasted. Mr and Mrs Ernest Mokezie showed me great kindness. You are
implicated for God’s blessings. For my siblings Mrs Ebele Nwankwo, Uka, Oby and Emmy, I
pray the good Lord think on you for your understanding and bless you along with all others who
have contributed to my success in one way or the other.
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ABSTRACT
Cellulases were produced from isolated fungus Aspergillus fumigatus using two distinct parts of
corn cob as carbon sources. The corn cobs used were collected from National Cereals Research
Institute, Yandev, Benue State; Agric Farms, UNN and Ogige Market, Nsukka. These plant
materials were sun-dried for three days and then broken to separate the hard outer part from the
soft inner part (the pulp). Each part was milled separately and the outer part was labeled CC-Outer
while the pulp, CC-Inner. Aspergillus fumigatus isolated from sewage water was adapted to each
of the two distinct parts of corn cob used as the sole carbon source. Assays conducted on
submerged fermentation filtrate from a pilot study showed that the organism’s production of
cellulase was highest on the 3rd
and 4th
days using CC-Inner and CC-Outer respectively as carbon
sources. Three litres of crude cellulases were produced from the organism using each of the two
carbon sources. The cellulase from CC-Inner was labeled CC-INNER while the one from CC-
Outer was labeled CC-OUTER. The two crude cellulases were partially purified by subjecting
them to 50% ammonium sulphate precipitation followed by dialysis. The partially purified
cellulases were then characterized with respect to pH, temperature and thermostability. While the
optimum pH of CC-INNER was 6.0, CC-OUTER was a neutral cellulase pH 7.0. The optimum
temperature of CC-INNER was 550C whereas that of CC-OUTER was 50
0C. The stability of the
enzymes from 350C to 70
0C within one hour decreased as indicated by reduced enzyme activites.
At 700C, CC-OUTER had lost 45.88% of its original activity while activity loss of 58.14% was
obtained for CC-INNER. The total cellulase activity and total protein of crude CC-OUTER were
14.861Units and 0.112mg/ml respectively while the respective values for the dialysed enzyme
were 45.833Units and 0.426mg/ml. For the crude CC-INNER, the total cellulase activity and total
protein were 26.111Units and 0.164mg/ml whereas the dialysed enzyme gave 92.917Units and
1.063mg/ml respectively. The study has shown that corn cob could serve as cheap carbon source
for fungal cellulase production especially, the pulp.
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TABLE OF CONTENTS
Title…………………………………………………………………………….………. i
Certification……………………………………………………………..…………….………...ii
Dedication……………………………………………………………........................iii
Acknowledgement……………………………………………………………………...iv
Abstract………………………………………………………………………………vi
Table of contents…………………………………………….………………..….….vii
List of tables.…………………………………………………………………….…..xii
List of figures..………………………………………………………………….…..xiii
List of plates………………………………………………………………………....xv
List of abbreviations..…..…………………………………………………………...xvi
CHAPTER ONE: INTRODUCTION
1.1 Lignocellulose …………………………………..……………………………....2
1.1.1 Lignin……………………………………………………………………..………………4
1.1.2 Hemicellulose………………………………………………………..………....5
1.1.3 Cellulose ……………………………………………………………..……......6
1.2 Degradation of lignocellulose…….…………………………………...........…....8
1.2.1 The chemical method of lignocellulosic biomass conversion………...….…....9
1.2.2 Thermochemical method of lignocellulosic biomass conversion ……….........9
1.2.2.1 Direct combustion …………………………………………………..….…..10
1.2.2.2 Gasification …………………………………………………………..…….10
1.2.2.3 Pyrolysis…………………………………………………………….......…..10
1.2.2.4 Liquefaction ….………………………………………………...…..………11
1.3 Biochemical method of lignocellulosic biomass conversion ………....……….11
1.3.1 Lignin – degrading enzymes………………………………………..…….......12
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1.3.2 Hemicellulose-degrading enzymes ………………………………..………....13
1.3.3 Cellulose-degrading enzymes….....………………………………………......15
1.3.3.1 Endoglucanases (Egl)…….……………………………………………...…16
1.3.3.2 Exoglucanases………….………………………………………………......18
1.3.3.3 β-Glucosidase …………….……………………………………………..…19
1.4 Production of cellulase..…………………….……………………………….....20
1.4.1 Factors affecting cellulase production …...……………………………….....21
1.4.1.1 pH ……………………………….…….……………………………….......21
1.4.1.2 Temperature…………………………….…………….…………………....22
1.4.1.3 Duration of incubation …...…………….……………….………………....23
1.4.1.4 Effect of metal ions on cellulase production….………...………………....24
1.4.1.5 Carbon source………………………………….……………………….......25
1.5 Structure of cellulase…..………………………….…………………………....26
1.5.1 Catalytic domain (CD)….……………..……………………………………..27
1.5.2 Linker…………………….……………………………………………….....27
1.5.3 The cellulose binding domain (CBD)…..……………….………………......28
1.6 Mechanism of action of cellulase...….………..………….……..…………......29
1.6.1 Synergism in cellulose hydrolysis.…….…..……….…………………..…....30
1.7 General features of Aspergillus fumigatus…...………..…………………....….32
1.7.1 Scientific classification…….………………………….……………………..33
1.8 Rationale of the research.….....……………...……………………………...…34
1.9 Objectives of the research…………………………………………………......34
CHAPTER TWO : MATERIALS AND METHODS
2.1 MATERIALS………………………………………………………………….35
2.1.1 Plant material….…………...………………………………………………...35
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2.1.2 Cellulolytic microorganism………………………………………………….35
2.1.3 Chemicals/Reagents..………………………………………………………………..35
2.1.4 Equipment/Apparatus.……………………………………………………….36
2.2 METHODS…………………………………………………………………....37
2.2.1 Processing of corn cobs ………..………………………………………....…37
2.2.2 Isolation of cellulolytic microorganisms……....………………………….....37
2.2.2.1 Collection of soil samples……………………………………………….....37
2.2.2.2 Determination of soil pH…….………….………………………………....37
2.2.2.3 Preparation of soil extract enrichment ..…..……….………………….......37
2.2.2.4 Preparation of mineral salt medium for isolation…..………….……….....38
2.2.2.5 Inoculation of the first isolation broth….…..…………………………......38
2.2.2.6 Sub-culturing into another isolation broth…........………….……………..38
2.2.2.7 Sub-culturing on solid media (SDA) ……………………………………..39
2.2.2.8 Inoculation of plates………..……….………………………………..........39
2.2.2.9 Storage on SDA slants ……………………………………………………39
2.2.2.10 Macroscopic photographs of the fungal isolates……..………....……....39
2.2.2.11 Culturing of the isolates on corn cob media ……….……………………39
2.2.3 Method of protein determination…...……………………………………….40
2.2.3.1 Protein calibration curve …..…………………………………………......40
2.2.4 Method of glucose determination…...………………………………...….....41
2.2.4.1 Glucose calibration curve …...…………...…………………………….…41
2.2.5. Sub-merged fermentation experiments……………………………...……..41
2.2.5.1 The fermentation broth …………..…………………………………...….41
2.2.5.2 Inoculation of the broth…..………………………………..………...…...42
2.2.5.3 Harvesting the fermented broth………………………………………………42
2.2.6 Cellulase assay …..……………………………………………………………..42
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2.2.6.1 Preparation of filter paper strips …..……...………………………………….42
2.2.6.2 Cellulase assay mixture……………...……………………………………….42
2.2.7 Protein assay mixture…………..……….………………………...…………….43
2.2.8 Partial purification of the crude enzyme filtrate…….………………………….43
2.2.8.1 Ammonium sulphate saturation and centrifugation……………………...….43
2.2.8.2 Dialysis…..…………………………………………………….…..………….43
2.2.8.3 Studies on partially purified cellulase samples..………..…………………….44
2.2.8.3.1 Effect of pH on enzyme activity…..………..………………………………44
2.2.8.3.2 Effect of temperature on enzyme activity….....…………………………….44
2.2.8.3.3 Heat stability study….………………….………………………………......44
2.2.9 Fungal identification….……………………………………………………...…45
CHAPTER THREE: RESULTS
3.1 Pilot study for determination of day of peak enzyme production…..…..………..46
3.2 Studies on mass-produced crude cellulases……………………………...….……50
3.2.1 Extracellular protein secretion…...……….…………………………………….50
3.2.2 Cellulase activities of the crude CC-OUTER and CC-INNER………..………51
3.3 Studies on partially purified cellulases………….………………………………..52
3.3.1a Purification for CC-OUTER……………….………………………………....52
3.3.1b Purification for CC-INNER……….……………………………………….…53
3.3.2 pH profile of partially purified cellulases……...…………………………….…54
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3.3.2a The pH profile of partially purified CC-INNER……………………………..54
3.3.2b The pH profile of partially purified CC-OUTER…………………………….55
3.3.3 Optimum temperature…............……..…………………………………………56
3.3.3a Optimum temperature for CC-INNER………………………………………..56
3.3.3b Optimum temperature for CC-OUTER……………………………………….57
3.3.4 Heat stability studies……...….…………………………………………………58
3.3.4a Heat stability study on CC-INNER…………………………………………...58
3.3.4b Heat stability study on CC-OUTER…………………………………………..59
3.3.5 Percentage loss in the activities of CC-INNER and CC-OUTER
from 35-700C…………………………………………………………………..60
3.4 Macroscopic photographs of isolated fungi…………..…..…………….…....…..62
CHAPTER FOUR: DISCUSSION AND CONCLUSION
4.1 DISCUSSION………………………………………………………………...…66
4.2 Conclusion...…………………….……………………………………………….71
4.3 Recommendations…..………….……..……………………………………...….71
References.…………………………………………………………………………..73
Appendices….…………………………………………………………………….…87
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LIST OF TABLES
Table 1: Lignocellulose contents of common agricultural residues and wastes..…..3
Table 2: Optimum pH for cellulase production by micro-organisms.……………...22
Table 3: Temperature optima for cellulase production in microorganisms………...23
Table 4: Optimal duration of incubation for microbial cellulase production……...24
Table 5: Purification for CC-OUTER……….…………….…….………………....52
Table 6: Purification for CC-INNER ……….……………….…………….............53
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LIST OF FIGURES
Fig. 1: Structure of lignin…..…………………………... …………………...…….......5
Fig. 2: Structure of hemicellulose showing some bonds between the sugars……....….6
Fig. 3: Chemical structure cellulose…..………………………………………………..8
Fig. 4: Schematic representation of sequential stages in cellulolysis……………..…..20
Fig. 5: Schematic representation of TrCBHI catalytic domain….……………………28
Fig. 6: Model describing the degradation of cellulose by cellulases…..……………...31
Fig. 7: Pilot study for determination of day of maximum enzyme production using
CC-Inner as carbon source (Total activity)……………….…….…………..…..46
Fig. 8: Pilot study for determination of day of maximum enzyme production using
CC-Outer as carbon source (Total activity)…………………….…………..…..47
Fig. 9: Pilot study for determining the day of maximum enzyme production using
CC-Inner as carbon source (Specific activity)………………………………….48
Fig. 10: Pilot study for determining the day of maximum enzyme production using
CC-Outer as carbon source (Specific activity)…………………………………49
Fig. 11: Comparison of extracellular protein secreted by the organism using CC-
Outer and CC-Inner as substrates………………………………………………50
Fig. 12: Comparison of total cellulase activities of crude CC-OUTER and
CC-INNER……………………………………………………………………..51
Fig. 13: Optimum pH of the partially purified CC-INNER………………………..…54
Fig. 14: Optimum pH of the partially purified CC-OUTER………………………….55
Fig. 15: Optimum temperature of the partially purified CC-INNER…………………56
Fig. 16: Optimum temperature of the partially purified CC-OUTER………………...57
Fig. 17: Heat stability of the partially purified CC-INNER…………………………..58
Fig. 18: Heat stability of the partially purified CC-OUTER………………………….59
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Fig. 19: Percentage loss in CC-INNER activity under varying
temperatures (35-700C)…………………………………………………………60
Fig. 20: Percentage loss in CC-OUTERER activity under varying
temperatures (35-700C)…………………………………………………………61
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TABLE OF PLATES
Plate 1: Aspergillus niger grown on Sabouraud Dextrose Agar……………......……………….62
Plate 2: Aspergillus spp grown on Sabouraud Dextrose Agar…………………………………..63
Plate 3: Aspergillus fumigatus grown on Sabouraud Dextrose Agar……………….……………64
Plate 4: Rhizopus spp grown on Sabouraud Dextrose Agar…………….....………………….….65
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LIST OF ABBREVIATIONS
CBD Cellulose Binding Domain
Cbh Cellobiohydrolase gene
CBH I Cellobiohydrolase I
CBH II Cellobiohydrolase II
CD Catalytic Domain
Cel 6A Cellulase gene 6A
Cex Clostridium exoglucanase
Cen Clostridium endoglucanase
CMCase Carboxymethylcellulase
DNS 3,5-Dinitrosalicylate
EG Exoglucanase
Egl Endoglucanase
FPase Filterpaperase
FPU Filter Paper Units
UI International Units
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CHAPTER ONE
INTRODUCTION
Common knowledge has it that till date, sugar-based industries continue to depend mainly
on carbohydrate-rich raw materials such as cereals, sugar-cane and tuber crops for their industrial
productions. These foodstuff-turned raw materials are not even enough to satisfy the dietary needs
of our hunger-stricken world. Their use in the production of sugar-based chemicals depletes our
food reserves and aggravates the already existing problems of hunger and starvation. Apart from
depriving the teeming population of food, industrial products become expensive due to high cost
of these foodstuff-turned raw materials. One major product of such industries is ethyl alcohol
which has very many uses. Ethanol is used as solvent, extractant and antifreeze. It is also used as
substrate for the synthesis of many other solvents of dyes, pharmaceuticals, lubricants, detergents,
pesticides, plasticizers, explosives and resins, and for the manufacture of synthetic fibres (Sasson,
1984).
In 1985, Brazil launched a programme of blending 20% ethanol (produced from sugar
cane and cassava) with petrol and thus saved about 40% of its petrol consumption (Dubey, 2008).
Before that time, USA had commercialized the same mixture as ‘gasohol’ thereby increasing
ethanol production in the country even from cereals. Yet our environment is littered with
agricultural wastes (corn cob, coco-nut coir, orange bagasse, sugar cane bagasse, etc), which
being lignocellulosic in nature, contain varying amounts of cellulose that can be harnessed
through simple technologies to produce cheaper ethanol. The production of ethanol from sugars or
starch impacts negatively on the economics of the process making ethanol more expensive
compared with fossil fuels (Howard et al., 2003). Plant cell walls are the most abundant renewal
source of fermentable sugars on earth (Himmel et al., 1999; Saleem et al., 2008) and are the major
reservoir of fixed carbon in nature (Yang et al., 2007).
In March, 2007, the US government awarded $385 million in grants aimed at jump-
starting ethanol production from non-traditional sources like wood chips, switchgrass and citrus
peel (Lammers, 2007). Cellulosic ethanol is a type of biofuel produced from lignocelluloses, a
structural material that comprises much of the mass of plants. In comparison to gasoline, ethanol
burns cleaner with a greater efficiency, thus putting less carbon dioxide and overall pollution in
the air. Moreso, only low levels of smog are produced from combustion (Demain et al., 2005).
According to the U.S. Department of Energy, ethanol from cellulose reduces green house gas
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emission by 90 percent, when compared to gasoline and in comparison to corn-based ethanol
which decreases emissions by 10 to 20 percent (Montenegro, 2006). Carbon dioxide gas emissions
are shown to be 85% lower than those from gasoline. Cellulosic ethanol contributes little to the
greenhouse effect and has a five times better net energy balance than corn-based ethanol (Demain
et al., 2005). When used as fuel, cellulosic ethanol releases less sulphur, carbon monoxide,
particulates, and greenhouse gases (Weeks, 2006).
The major components of all plant cell walls are cellulose, hemicelluloses and lignin.
Cellulose is the most abundant component. Large amounts of lignocellulosic “waste” are
generated through forestry and agricultural practices, paper-pulp industries and they pose an
environmental pollution problem. Sadly, much of the lignocellulosic waste is often disposed of by
biomass burning, which is not restricted to developing countries, but is considered a global
phenomenon (Levine, 1996).
One of such lignocellulosic wastes used to enrich the dustbins or refuse dumps or worse
still, burnt especially in developing economies is the corn cob. When corn is harvested, the kernel
or grains are used but the cob is thrown away as useless. It is therefore the focus of this research to
see how the cellulose in corn cob can be used as a cheap raw material for cellulase production.
Moreso, no work has been done on the comparative yield of cellulase from the pulp of corn cob
and the hard outer part.
1.1 Lignocellulose
Lignocellulose is the major structural component of woody plants and non woody plants
such as grass and represents a major source of renewable organic matter. Lignocelluloses consist
of lignin, hemicelluloses and cellulose. The chemical properties of the components of
lignocellulosics make them a substrate of enormous biotechnological value (Malherbe and Cloete,
2003). Table 1 below as compiled by Betts et al.(1991); Sun and Cheng (2002) shows the typical
compositions of the three components in various lignocellulosic materials.
Table 1 : Lignocellulose contents of common agricultural residues and wastes
Lignocellulosic materials Cellulose (%) Hemicelluloses (%) Lignin (%)
Hardwood stems 40-55 24-40 18-25
Softwood stems 45-50 25-35 25-35
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Nut shells 25-30 25-30 30-40
Corn cobs 45 35 15
Paper 85-89 0 0-15
Wheat straw 30 50 15
Rice straw 32.1 24 18
Sorted refuse 60 20 20
Leaves 15-20 80-85 0
Cotton seed hairs 80-95 5-20 0
Newspaper 40-55 25-40 18-30
Waste paper from chemical Pulps 60-70 10-20 5-10
Primary waste water Solids 8-15 Not available 24-29
Fresh bagasse 33.4 30 18.9
Swine waste 6 28 Not available
Solid cattle manure 1.6-4.7 1.4-3.3 2.7-5.7
Coastal Bermuda grass 25 35.7 6.4
Switch grass 45 31.4 12.0
S32 rye grass (early leaf) 21.3 15.8 2.7
S32 rye grass (seed setting) 26.7 25.7 7.3
Orchard grass (medium maturity ) 32 40 4.7
Grasses 25-40 25-50 10-30
Source: Betts et al. (1991); Sun and Cheng (2002).
1.1.1 Lignin
The word lignin originates from the Latin ‘lignum’ which means wood (Sarkanen and
Ludwig, 1971). It is the chemical substance that confers rigidity and recalcitrance to microbial
attack in trees and plants. In lignocellulosic materials, lignin is the most abundant non-
polysaccharide. It is a high molecular weight material composed of phenylpropene units.
The three units which comprise the lignin complex network, are guaiacyl (from the
precursor coniferyl alcohol), springyl (from the precursor sinapyl alcohol) and p-hydroxyphenyl
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(from the precursor p-coumaryl alcohol). Hardwood lignin with a higher content has been found
easier to extract by alkaline extraction as compared to softwood lignin (Ramos et al., 1992).
An interesting difference between the synthesis of cellulose and hemicelluloses and that of
lignin polymer is that lignin is synthesized by radical coupling of randomly organized monomeric
units. In contrast, cellulose and hemicelluloses are synthesized enzymatically in a structured
manner (Nimz, 1974). Lignin is phenolic in nature; it is very stable and difficult to isolate. It
occurs between the cells and the cell walls. It is deposited during lignification of the plant tissue
and gets intimately associated within the cell walls with cellulose and hemicelluloses and impacts
the plant an excellent strength and rigidity (Dubey, 2008). In other words, in lignocellulose
structure lignin is linked to both hemicelluloses and cellulose forming a physical seal around them
such that an impenetrable barrier exists preventing the penetration of solutions and enzymes.
Fig. 1: Structure of lignin
Source: http://www.xplora.org/downloads/Knoppix/ESPER
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1.1.2 Hemicellulose
Hemicelluloses, the second most common polysaccharides in nature, represent about 20 to
35% of lignocellulosic biomass (Koukiekolo et al., 2005). Hemicellulose macromolecules are
often polymers of pentoses (xylose and arabinose), hexoses (mostly mannose) and a number of
sugar acids (Howard et al., 2003). The proportion of these constituents differ from plant to plant
and the degree of polymerization that brings about the formation of the polymer does not exceed
50. The polymer has branched chains and occurs as amorphous mass around the cellulose strands,
insoluble in water but easily solubilized in alkali.
Xylans are the most abundant hemicelluloses. Xylans of many plant materials are
heteropolysaccharides with homopolymeric backbone chains of 1,4-linked -D-xylopyranose
units. Besides xylose, xylans may contain arabinose, glucuronic acid, or its 4-0-methyl ether and
acetic, ferulic, and p-coumaric acids. The frequencies and composition of branches depend on the
source of xylan (Aspinall, 1980).
Fig. 2: Structure of hemicellulose showing some of the bonds between the
sugars ( -xylose-β(1,4)- mannose- β(1,4)- glucose-α(1,3)- galactose )
Source: http://wiki.lamk.fi/display/rebwp7/Bioethanol+production
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Complete hydrolysis of hemicelluloses requires the interaction of a number of hemicellulosic
enzymes that have the ability to cleave main chains and side chains (Koukiekolo et al., 2005).
1.1.3 Cellulose
Cellulose is the most abundant polymer in the biosphere with its estimated synthesis rate
of 1010
tonnes per year (Schlesinger, 1991; Singh and Hayashi, 1995; Lynd et al., 2002).
According to Hornby (2010), it is a natural substance that forms the cell walls of all plants and
trees and is used in making plastics, paper, etc. Cellulose-rich plant biomass is one of the
foreseeable and sustained source of fuel, animal feed and feedstock for chemical synthesis (Bhat,
2000).
Cellulose is an unbranched polymer of -1,4 linked anhydrous glucose units constituting
40-60% of cell wall materials of plant (Thieman and Palladino, 2009). It can have a degree of
polymerization (DP) of up to 15,000 D-glucose units. D-glucose molecules exist in different
forms in aqueous solution. In solution, the cyclic hemiacetal forms will exist in a ratio of and
anomers, thermodynamic phenomenon called mutarotation. Upon polymerization of glucose units,
0-glycosidic bonds are formed, whereby the glycosidic carbon C-1 is linked through an oxygen
atom to a carbon atom C-4 on another monosaccharide residue, a reaction that involves the loss of
a molecule of water.
Depending on whether the configuration at the anomeric C-1 carbon of a residue is or ,
resulting glycosidic bond will be either an -1, 4-glycosidic bond or a -1, 4-glycosidic bond. The
polymerization of many -D-glucose units produces the polymer known as starch, whereas the
polymerization of -anomers of D-glucose yields cellulose. In a cellulose chain every monomeric
glucose unit is rotated 1800
relative to one another thus making cellobiose the repeating unit. In
aqueous solution, the hemiacetals at the reducing end of cellulose chain exist in equilibrium with
relatively small, yet significant amounts of non-cyclic aldehydes. These non-cyclic aldehydes can
undergo oxidation and so are detectable through the use of different oxidizing reagents.
Starch, a polymer linked in -1, 4-glycosidic bonds, tend to coil into helical structures. In
contrast, cellulose, linked in -1, 4-glycosidic bonds adopts long, linear chains. The linear
arrangement of cellulose presents a uniform distribution of –OH groups that can ‘zip’ the chain
together by intra-and interchain hydrogen bonding. Zipping many cellulose chains together
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produces a crystalline lattice in the cell wall. The chains are arranged in microfibrils of about 30
to 100 molecules lying side-by-side. Typically, 36 cellulose chains with the same direction, but
with different starting and ending points, assemble into one microfibril with a diameter of 5-
15nm. The microfibril is greatly stabilized through intermolecular hydrogen bonds (Gardner and
Blackwell, 1974). These intra and intermolecular interactions in a microfibril most probably make
cellulose resistant to degradation.
Four polymorphs of crystalline cellulose are known: cellulose I, cellulose II, cellulose III
and cellulose IV. Native cellulose always exists as cellulose I, although cellulose II is the more
thermodynamically stable form. In cellulose I, the parallel glucan chains all run in the same
direction (Chanzy and Henrissat, 1985), whereas the glucan chains in cellulose II are probably
arranged in an antiparallel fashion. If cellulose I is dissolved and recrystallized, it is converted to
cellulose II. Cellulose III is obtained by treatment of cellulose II with alkali solution and vacuum
drying. The swollen cellulose crystals hold ions and water molecules within the lattice that
influence the packing (Porro et al., 2007). Cellulose IV is obtained by heating cellulose III in
glycerol. Both cellulose III and cellulose IV revert to the parent polymorph (I or II) on
hydrothermal treatment. All native celluloses are a composite mixture of two crystalline
allormorphs, designated I and I. The main component in higher plants is I and in bacteria,
tunicates and algae, the dominating component is I (Sturcova et al., 2004).
Fig. 3: Chemical structure of cellulose.
Source: http://www.scientificpsychic.com/fitness/carbohydrates2.html
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1.2 Degradation of lignocellulose
Lignocellulose materials vary in their proportions of cellulose, hemicelluloses and lignin.
Of the three components, lignin is the most recalcitrant to degradation whereas cellulose, because
of its highly ordered crystalline structure, is more resistant to hydrolysis than hemicelluloses.
Alkaline (Chahal, 1992) and acid (Nguyen, 1993; Grethlein and Converse, 1991) hydrolysis
methods have been used to degrade lignocelluloses.
Weak acids tend to remove lignin but result in poor hydrolysis of cellulose whereas strong
acid treatment occurs under relatively extreme corrosive conditions of high temperature and pH
which necessitate the use of expensive equipment. Also, unspecific side reactions occur which
yield non-specific by-products other than glucose, promote glucose degradation and therefore
reduce its yield. Some of the unspecific products can be deleterious to subsequent fermentation
unless removed. There are also environmental concerns associated with the disposal of spent acid
and alkali (Howard et al., 2003).
There are technologies currently used to convert lignocelluloses to ethanol and other
chemical products. These technologies include chemical, thermochemical and biochemical
processes. Among all these processes or methods, biochemical method is probably the safest since
it employs nature’s chiral molecules and does not cause environmental pollution.
1.2.1 The chemical method of lignocellulosic biomass conversion
This basically entails the breakdown of lignocellulosic materials using either dilute or
concentrated acids, organic solvents e.g. organosolv, ammonia, etc. Dilute acid with no enzymes
added has been used for the complete hydrolysis of lignocelllosic material on a commercial scale
since the beginning of the 1930s (the Scholler or Madison percolation process (Jones and Semrau,
1984; Parisi, 1989). The most generally used acid is H2SO4 due to lower price and fewer problems
with corrosion compared to HCl. However, glucose yield seldom exceeds 55-60% of the
theoretical yield using this method (Kadam et al.,2000; Lee et al., 1999; Parisi, 1989). The major
problem here is that high temperatures are needed, and at such temperatures, pentoses from
hemicelluloses and hexoses are rapidly degraded to by-products which inhibit further yield from
fermentation since it is usually a method of pretreatment. There is also a need for advanced
reactor design as well as negative impact on the environment. In the case of organosolv there is
high cost of recovery of organic solvent.
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1.2.2 Thermochemical method of lignocellulosic biomass conversion
This encompasses different non-biological routes of biomass conversion into energy, and
include: (a) direct combustion, (b) gasification, (c) pyrolysis and (d) liquefaction.
1.2.2.1 Direct Combustion
Biomass from plants (wood, agricultural wastes) or animal (cow dung) origin are directly
burnt for cooking and other purposes. In recent years, “hogfuel” production technology has been
developed which is being utilized for generation of electricity (Dubey, 2008). Nowadays
municipal agricultural and light industrial wastes are used for conversion into energy by direct
burning in refuse fired energy systems (Ghosh and Bisaria, 1981). Hog fuel combustion
technology has been developed recently in the USA. This fuel which is a mixture of wood and
bark waste burnt directly, is produced in large sized boilers made up of steel. Boilers are designed
time to time to develop a good control system of combustion (Dubey, 2008).
1.2.2.2 Gasification
This is a process of thermal degradation of carbonaceous material under controlled amount
of air or pure oxygen, and high temperature up to around 1,0000C. As a result of gasification, high
amount of gases is produced. This gasification of biomass is done in a gasifier designed in various
ways (Dubey, 2008). The gasification process does not rely on chemical decomposition of
cellulose chain (cellulolysis). Instead of breaking the cellulose into sugar molecules, the carbon in
the raw material is converted into synthesis gas, using what amounts to partial combustion. The
carbon monoxide, carbon dioxide and hydrogen may then be fed into a special kind of fermenter.
Instead of sugar fermentation with yeast, this process uses a microorganism named Clostridium
ljungdahlii. This microorganism will ingest (eat) carbon monoxide, carbon dioxide and hydrogen
and produce ethanol and water. Alternatively, the synthesis gas from gasification may be fed to a
catalytic reactor where the gas is used to produce ethanol and other higher alcohols through
thermochemical process.
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1.2.2.3 Pyrolysis
Pyrolysis is defined as the destructive distillation or decomposition of organic matter, for
example, solid residues, wastes (saw dust, wood chips, wood pieces) in an oxygen deficient
atmosphere or in the absence of oxygen at high temperature (200-5000C or rarely 900
0C).
Products of pyrolysis are gases, organic liquids and chars, depending on the pyrolysis process and
temperature of reaction. The condensable liquids separate into aqueous (pyroligneous acid), oil
and tar fraction (if the substrate is wood). The composition of gas is carbon monoxide (28-33%),
methane (3-18%), higher hydrocarbons (1-3%). During pyrolysis, hydrogen content of gas
increases with increasing temperature (Jahn, 1982).
Pyrolysis has been employed to produce charcoal for the last few decades. Charcoal is a
smokeless and low sulphur fuel used mostly for cooking purpose. Apart from wood, other
wastes/residues used in pyrolysis are cotton, sugar-cane bagasse, groundnut shell, etc.
1.2.2.4 Liquefaction
Liquefaction involves the production of oils for energy from wood or agriculture and
carbon residues by reacting them with carbon monoxide and water/steam at high pressure
(4,000ib/in2) and temperature (350-400
0C) in the presence of catalysts. By this method about 40-
50% oil can be obtained from wood. This oil serves as good source of fuel (Jahn, 1982).
1.3 Biochemical method of lignocellulosic biomass conversion
Biochemical method of lignocelluloses conversion otherwise called bioconversion converts
lignocellulosic biomass into energy, fertilizer, food and chemicals through biological agency. The
biological agents are the micro-organisms such as bacteria, actinomycetes, fungi and algae. These
organisms secrete nature’s chiral catalysts known as enzymes with which they breakdown the
biopolymers.
There have been several attempts to pretreat lignocelluloses using various chemicals and
temperatures. The objectives of the pretreatment are to increase the enzymatic accessibility of the
cellulose as well as the hemicelluloses. The first step in the pretreatment is a mechanical reduction
of the size of the lignocelluloses while the second is a physical and/or chemical method. However,
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because of certain disadvantages of some of the pretreatment methods, enzymes are best used for
lignocellulosic biomass conversion. The lignin complex can be degraded enzymatically to the
phenylpropene monomers instead of using synthetic chemicals. Microorganisms degrading lignin
have been widely studied, such as white rot fungi and the enzyme system from Phanerochaete
chrysosporium (Shoemaker and Leisola, 1990). To get reduced carbon compounds required for
energy production, microorganisms produce cellulolytic enzymes to degrade the insoluble
cellulose into soluble oligomers which can either be directly taken up and metabolized or
degraded to glucose before taken up.
Cellulolytic enzymes can be divided into two distinct groups:
(a) Enzymes produced by aerobic microorganisms
(b) Enzymes produced by anaerobic microorganisms
While the former are secreted as individual cellulolytic enzymes, the latter are released as
multi-enzyme complexes called cellulosomes. Cellulolytic enzymes of fungi are usually secreted
into the external milieu as extracellular enzymes.
There are however, three classes of enzymes that degrade lignocelluloses, each class
degrading one of the three components of the biomass. These enzymes are lignases,
hemicellulases and cellulases. The lignases work on lignin content while hemicellulases hydrolyse
the constituent hemicelluloses. The cellulose which remains is finally saccharified by cellulolytic
enzymes known as cellulases.
1.3.1 Lignin – degrading enzymes
The white rot fungi belonging to the basidiomycetes are the most efficient and extensive
lignin degraders (Akin et al., 1995; Gold and Alic, 1993) with P. chrysosporium being the best
studied lignin-degrading fungus producing copious amounts of a unique set of lignocellulytic
enzymes. It is a good host for the production of lignin degrading enzymes or even in the case of
direct application to lignocelluloses bioconversion processes. Less known white-rot fungi such as
Daedalea flavida, Phlebia fascicularia, P. floridensis and P. radiate have been found to
selectively degrade lignin in wheat straw and hold out prospects for bioconversion biotechnology
where the aim is just to remove the lignin leaving the other components almost intact (Arora et al.,
2002). Less prolific lignin degraders among bacteria such as those belonging to the genera
Cellulomonas, Pseudomonas and the actinomycetes Thermomonospora and Microbispora and
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bacteria with surface-bound cellulase-complexes such as Clostridium thermocellum and
Ruminococcus are beginning to receive attention as representing a gene pool with possible unique
lignocellulolytic genes that could be used in lignocellulase engineering (Vicuna, 1988; McCarthy,
1987; Miller (Jr) et al., 1996; Shen et al., 1995; Eveleigh, 1987). Two families of lignolytic
enzymes are widely considered to play a key role in the enzymatic degradation: phenol oxidase
(laccase) and peroxidase (Mn P) (Krause et al., 2003; Malherbe and Cloete, 2003). Other enzymes
whose roles have not been fully elucidated include peroxide-producing enzymes: glyoxal oxidase
(Kersten and Kirk, 1987), glucose oxidase (Kelly and Reddy, 1986), veratryl alcohol oxidases
(Barbonnais and Paice, 1988), methanol oxidase (Nishida and Eriksson, 1987) and oxidoreductase
(Bao and Renganathan, 1991).
1.3.2 Hemicellulose-degrading enzymes
Hemicellulose is a collective term referring to those polysaccharides soluble in alkali.
They are associated with cellulose of the plant cell wall. They would include non-cellulose -D-
glucans, pectic substances like polygalacturonans. They also include several
heteropolysaccharides such as those consisting mainly of galactose (arabinogalactans), mannose
(galactoglucomannans and glucomannans) and xylose (arabinoglucuronoxylans and
glucuronoxylans). However, only the heteropolysaccharides with a much lower degree of
polymerization (100-200 units) as compared to that of cellulose (10,000-14,000 units), are
referred to as hemicelluloses. According to Howard et al., (2003), the principal sugar components
of these hemicelluloses polysaccharides are: D-xylose, D-mannose, D-glucose, D-galactose, L-
arabinose, D-glucuronic acid, 4-0-methyl-D-glucuronic acid, D-galacturonic acid, and to a lesser
extent, L-rhamnose, L-fucose and various O-methylated sugars.
Hemicellulases like most other enzymes which hydrolyse plant cell polysaccharides are
multidomain proteins (Henrissat and Davies, 2000; Prates et al., 2001). These proteins generally
contain structurally discrete catalytic and non-catalytic modules. Depending on the amino acid or
nucleic acid sequence of their catalytic modules, hemicellulases are either glucoside hydrolases
(GHs) which hydrolyse glycosidic bonds or carbohydrate esterases (Ces), which hydrolyse ester
linkages of acetate or ferulic acid side groups and according to their primary sequence homology
they have been grouped into various families.
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Exo--1, 4-xylosidase (EC: 3.2.1.37) acts on -1, 4-xylooligomers xylobiose, endo--1, 4-
xylanase acts on -1, 4-xylan. Exo--1, 4-mannosidase (EC: 3.2.1.25) acts on -1, 4-
mannooligomers mannobiose. Endo--1, 4-mannanase (EC: 3.2.1.78) acts on -1, 4-mannan. In
the same vein, endo--1, 5-arabinase (EC: 3.2.1.99) acts on -1, 5-arabinan while -L-
arabinofuranosidase (EC: 3.2.1.55) acts on -arabinofuranosyl (12) or (13) xyloogligomers
-1, 5-arabinan. -Glucuronidase (EC: 3.2.1.39) acts on 4-0-methyl--glucuronic acid (12)
xylooligomers. Furthermore -Galactosidase (EC: 3.2.1.22) works on -galactopyranose (16)
mannooligomers. The last two enzymes among the major glycoside hydrolases are
endogalactanase (EC: 3.2.1.89) and -glucosidase (EC: 3.2.1.21) which act on -1,4-galactan and
-glucopyranose(14) mannopyranose respectively.
On the other hand, there are three enzymes of the major hemicellulase classification which
are esterases. These hemicellulolytic esterases include feruloyl esterase which hydrolyses the ester
linkage between the arabinose substitutions and ferulic acid, and acetyl esterases which hydrolyse
the acetyl substitutions on xylose moieties. Feruloyl esterases aid the release of hemicelluloses
from lignin and renders the free polysaccharide product more amenable to degradation by the
other hemicellulases (Prates et al., 2001).
Xylan is the most abundant hemicelluloses so that xylanases are one of the major
hemicellulases which hydrolyse the -1,4 bond in the xylan backbone yielding short
xylooligomers which are further hydrolysed into single xylose units by -xylosidase.
Hemicellulases, no doubt, are more complex and more numerous than the cellulose-degrading
enzymes. This reflects the complexity of hemicelluloses structure which comprises of several
sugars with several connections and osidic bonds, unlike cellulose which has a linear sequence of
glucose as its only monomeric unit. Hemicellulases and cellulose-degrading enzymes are similar
in the sense that they are majorly extracellular, requiring no cofactor. They are also characterized
by the presence of two acidic residues in their catalytic site. Their genes are controlled positively
by the substrate of degradation but negatively by the end-product of hydrolysis.
1.3.3 Cellulose-degrading enzymes
Thieman and Palladino (2009) reported that the breaking apart of sugar molecules that
make up cellulose is already being accomplished not only by the enzymes from fungi, the gut of
termites, but it can also come from the lab of genetic engineers. Similarly, cellulolytic property of
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bacterial species like Pseudomonas, Cellulomonas, Bacillus, Micrococcus, Cellovibrio and
Sporosphytophaga spp. were also reported (Nakamura and Kappamura, 1982; Immanuel et al.,
2006). Although a large number or array of microorganisms have the capacity of degrading
cellulose, only a few of them produce significant amounts of cell free or extracellular enzymes
that can completely hydrolyse crystalline cellulose in vitro.
Cellulases (1,4--D-glucanohydrolase, EC: 3.2.1.4) are multienzyme complexes,
comprising three main components; endo--glucanase (EC 3.2.1.4), exo--glucanase (EC
3.2.1.91) and -glucosidase (EC 3.2.1.21), which have been shown to act synergistically in the
hydrolysis of cellulose (Emert et al., 1974; Ryu and Mandels, 1980). The endoglucanase attacks
cellulose randomly along the cellulose chain yielding glucose and cello-oligosaccharides while the
exoglucanase cleaves from the non-reducing end of cellulose yielding cellobiose units. -1,4-
glucosidase (cellobiase) finally hydrolyses cellobiose to glucose (Okafoagu and Nzelibe, 2006).
Purified celluloses used as model substrates for studies of hydrolysis vary in structural
details. Bacterial microcrystalline cellulose, BMCC, synthesized by Gluconoacetobacter xylinus
(formerly Acetobacter xylinum), is the most highly crystalline substrate, but it contains mainly
allomorph I, which will have positive implications for the enzymatic utilization (Hayashi et al.,
1997). Avicel and Sigmacett are also highly crystalline, nearly pure celluloses from plant origin
prepared by removal of hemicelluloses and amorphous regions. The insoluble substrate is difficult
to attack by enzymes since the individual molecules are packed so tightly that not even water
molecules can enter the crystalline array. Phosphoric acid swollen cellulose, PASC, is an example
of amorphous substrate where the hydrogen bonds have been removed via acid treatment. The
difficulty encountered while working with insoluble substrates has given rise to the use of soluble,
artificial derivatives, carboxymethlcellulose (CMC) and hydroxyethylcellulose (HEC). Many
enzymes active on PASC, CMC or HEC cannot hydrolyze crystalline cellulose, but are still
referred to as “cellulases”. Filter paper is considered to be highly crystalline cellulose and IUPAC
has recommended the hydrolysis of filter paper as a standard measurement for total cellulolytic
activity (Ghose, 1984).
In a typical cellulose-degrading ecosystem, a variety of cellulolytic bacteria and fungi
work in concert with related microorganisms to convert insoluble cellulosic substrates to soluble
sugars-primarily cellobiose and glucose, which are then assimilated by the cell (Bayer et al.,
1998). These microbes are capable of carrying out this function by producing a variety of different
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enzymes which are called cellulases. This need for an enzyme with three catalytic entities is
traceable to the very complex structure of cellulose with extensive intermolecular bonding pattern,
even though all cellulases cleave a single type of bond. According to Holker et al. (2004),
Ahamed and Vermette (2008), the most promising technology for the conversion of the
lignocellulosic biomass to fuel ethanol is based on the enzymatic breakdown of cellulose using
cellulases. Cellulose-degrading enzymes, according to Ahmed et al. (2009), provide a key
opportunity for achieving tremendous benefits of biomass utilization.
1.3.3.1 Endoglucanases (Egl)
(1,4--D-glucan 4-glucanohydrolase, EC 3.2.1.4). Endoglucanases or CMCases catalyse
the hydrolysis of internal 1,4--D-glycosidic linkages in cellulose, lichenin, and cereal -D-
glucans. Endoglucanases, often called carboxymethylcellulose (CM)-cellulases, are proposed to
initiate attack randomly at multiple internal sites in the amorphous regions of the cellulose fibre
opening up sites for subsequent attack by the cellobiohydrolases (Wood, 1991). A number of
endoglucanases are found in Trichoderma reesei culture filtrates. The primary structures of Egl I
and Egl II (formerly known as Egl III) are deduced from their respective nucleotide sequences
(Saloheimo et al., 1988). Egl I consists of 459 amino acid residues) with a calculated MW =
48,212. Its cellulose-binding domain is at the C-terminal region. The amino acid sequence of the
Egl II protein, as deduced from the nucleotide sequence, consists of 418 amino acids among
which 21 residues make up the leader sequence. Its molecular weight is 42,200 and unlike the Egl
I, its cellulose-binding domain is at the N-termius (Stahlberg et al., 1988).
Baker et al., (1992) observed that Egl II exhibits greater thermal stability than Egl I. Its
melting temperature Tm is 750C while that of Egl I is 64
0C. Both Egl I and Egl II hydrolyze cello-
oligosaccharides with the same efficiency, except that the latter does not hydrolyze cellobioside
(Saloheimo et al., 1988).
Two endoglucanases (Cen A and Cen B) have been isolated from the bacterium
Celulomonas fimi, and their primary structures deduced from the nucleotide sequences (Wong et
al., 1986); Meinke et al., 1991). The nucleotide sequence encodes 449 amino acids (31 being in
the leader peptide) with a calculated MW of 51,800. The CBD (~100 amino acids) at the N-
terminus links with the C-terminal core segment (~300 amino acids) via a short Pro/Thr-rich
linker (Gilkes et al., 1988). Cen B, according to Meinke et al. (1991), consists of 1,045 amino
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acids encoded by the gene sequence excluding 31 residues in the leader sequence. It has a
calculated MW of 106,905 and is the longest cellulase sequenced so far. Cen B has an optimum
pH of 8.5-9.0 for CMC hydrolysis while 7.0-7.5 is the optimum pH of Cen A.
Beguin et al. (1985) also isolated six endoglucanases from Clostridium thermocellum and
sequenced their genes designated as celA, celB, celC, celD, celE and celH. The gene product of
celA gene is celA and is a polypeptide of 477 amino acids having a MW of 52,503. Thirty-two of
the encoded amino acids are found in the signal sequence. CelB consists of 563 amino acid
residues with a MW of 63,857. The N-terminal sequence does not resemble other signal peptides,
suggesting that the protein is not secreted (Grepinet and Beguin, 1986). CelC is a protein of 343
amino acids (14 amino-acid signal peptide) with a calculated MW of 40,439 (Schwarz et al.,
1988). The nucleotide sequence of celD encodes 649 amino acids with 41 residues in the signal
sequence. It has a MW calculated to be 72,334. While celE contains 814 amino acids (34 in the
signal peptide), MW 90,211, the celH gene encodes 900 amino acids (44 amino acids forming the
signal peptide); the molecular weight calculated was 102,301 (Hall et al., 1988).
1.3.3.2 Exoglucanases
(1,4--D-glucan cellobiohydrolase, EC 3.2.1.91). Exoglucanases or cellebiohydrolases
catalyse the removal of cellobiose units from non-reducing ends of cellulose chains. Other names
of the enzyme include exocellulase, cellobiosidase and avicellase (Wong, 1995).
Two cellobiohydrolases (CbhI and CbhII) secreted by the fungus Trichoderma reesei have
been purified from culture filtrates. CbhI comprises about 60% of the total extra cellular proteins
in culture filtrate. It has a molecular weight of 65KD and 10% carbohydrate content with an
isoelectric point (pI) of 4.4. Nummi et al. (1983) observed also that the enzyme, CbhI attacks both
amorphous and crystalline celluloses but has no activity on substituted celluloses such as
carboxymethylcellulose and hydroxyethylcellulose, and even on other soluble substrates like
cellohexaose, p-nitrophenyl -glucoside. CbhII, a minor enzyme has MW of 53KD, 8%
carbohydrate content, and a pI of 5.0. The enzyme, like CbhI, is not active toward CMC (Wood et
al., 1989).
For bacterial cellulase systems, the sequence of Cex gene has been reported encoding an
exoglucanase, Cex, in Cellulomonas fimi (O’Neill, 1986). The gene encodes for 484 amino acids.
This includes 41 residues acting as the leader peptide. For the gram-positive, thermophilic,
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anaerobic Clostridium thermocellum, only one cellobiohydrolase has been isolated in a truncated
form from the cellulosome of this bacterium, in spite of the fact that six endoglucanase genes have
been isolated and sequenced (Morag et al., 1991). The active cellobiohydrolase fragment has a
MW of 68KD. The enzyme activity is unstable at temperatures above 500C, but can be enhanced
by the addition of calcium and thiol-reducing agents. It is interesting to note that the cellulose
activity of the cellulosome of Clostridium thermocellum is also activated by the presence of
calcium and dithiothreitol (Johnson et al., 1982; Johnson and Demain 1984).
1.3.3.3 -Glucosidase
(-D-glucoside glucohydrolase EC 3.2.1.21). -glucosidase or cellobiase catalyses the
hydrolysis of cellobiose and removal of glucose from non-reducing ends of cello-oligosaccharides
and glycosyl transfer to cellobiose (Wong, 1995).
Multiple forms of -glucosidase, both intracellular and extracellular, are reported
produced by Trichoderma reesei, and have pI values ranging from acidic to alkaline (pH 4.4-8.4),
and MW from 50 to 98kilo dalton (Inglin et al., 1980; Enari et al., 1981; Umile and Kubicek,
1986). Less than 50% of the -glucosidases is secreted into the culture medium; the rest remain
tightly associated with a cell wall polysaccharide consisting of mannose, galactose, glucose,
galacturonic and gluconic acids. This heteroglycan binds -glucosidase in vitro, and the binding
results in a 2-fold increase in the enzyme activity against p-nitrophenyl -glycoside (Messner et
al., 1990).
-Glucosidase represents about 1.0% of the total proteins secreted, which is insufficient
for the Trichoderma reesei cellulase system to achieve practical saccharification of cellulose. The
cellobiose, accumulated by the action of endoglucanases and cellobiohydrolases, exhibits end-
product inhibition. Trichoderma reesei mutants have been isolated with increased production (up
to 3 times more) of -glucosidase (Kawamori et al., 1986). Alternatively, complete conversion of
cellulose to glucose can often be achieved by supplementing the cellulase system with -
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glucosidase from a different source, for example, Aspergillus niger, which has a high production
of the enzyme (about 10 times that of Trichoderma reesei) (Sternberg et al., 1977).
Based on the characterization of two distinct genes bglA and bglB in Clostridium
thermocellum, there is definitely more than one -glucosidase produced by the bacterium
(Schwarz et al., 1985; Grabnitz and Staudenbauer, 1988). BglA, as deduced from the nucleotide
sequence of the recombinant gene in Escherichia coli, consists of 448 amino acids and has a MW
of 51,482. The enzyme has an optimum activity at pH 6.0-6.5 at 600C. The enzyme hydrolyzes
both cellobiose and p-nitrophenyl -glycoside. It also hydrolyzes cello-oligosaccharides but not
CMC (Wong, 1995).
The primary structure of BglB, as encoded by the nucleotide sequence, consists of 754
amino acids with a calculated MW = 84,100 (Grabnitz et al., 1989). The N-termini of BglA and
BglB contain no basic or hydrophobic regions characteristic of a leader peptide, suggesting that
these enzymes are localized in the cytoplasmic region (Grabnitz et al., 1991).
1.4 Production of cellulase
Fig. 4: Schematic representation of sequential stages in cellulolysis
Source: Deacon (1997)
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Successful utilization of cellulosic materials as renewable carbon sources is dependent on
the development of economically feasible process technologies for cellulase production. It is
already established that not only micro-organisms such as fungi, bacteria and actinomycetes
produce cellulase, but also plants. Even small animals such as ants and snails produce certain
amounts of cellulase to be able to metabolize their food.
In general, bacterial cellulases are constitutively produced whereas fungal cellulases are
produced in the presence of cellulose (Suto and Tomita, 2001). Many fungal strains secrete higher
amounts of cellulases than bacterial ones with Trichoderma as the leading one (Amouri and
Gargouri, 2006). Most commercial cellulases are mesophilic enzymes produced by the
filamentous fungus Trichoderma reesei and Aspergillus niger. Other cellulase-producing fungi
include Acremonium cellulolyticus, Aspergillus acculeatus, Aspergillus fumigatus, Fusarium
solani, Irpex lacteus, Penicillium funmiculosum, Phanerochaete chrysosporium, Sporotichum
cellulophilum, Talaromyces emersonii, Trichoderma koningii, T. viride, etc. Bacteria, because of
their high growth rate compared to fungi have good potential to be used in cellulase production.
However, the application of bacteria in producing cellulase is not widely used (Abou-Taleb et al.,
2009). This is probably because bacterial cellulases are usually cell-bound and not secreted
whereas fungal cellulases are extracellular enzymes which are secreted directly into the culture
medium. As a result, our discussion will focus majorly on fungi.
Cellulase yields appear to depend on a complex relationship involving a variety of factors
like inoculum size, pH, temperature, presence of inducers, medium additives, aeration, growth
time, etc (Immanuel et al., 2006). Carbon source and quality of cellulose used are also factors that
affect the yield.
1.4.1 Factors affecting cellulase production
1.4.1.1 pH
This is a measure of hydrogen ion activity of a solution and is defined as the negative
logarithm of the hydrogen ion concentration. It is not surprising that pH dramatically affects
microbial growth. Each species has a definite pH for optimum growth. Acidophiles have their
growth optimum between pH 0 and 5.5; neutrophiles, between pH 5.5 and 8.0; and alkalophiles
prefer the pH range of 8.0 to 11.5 (Willey et al. 2008). A knowledge of the pH at which each
strain of microorganism produces cellulases maximally is of great importance to an industrial
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enzymologist. Several studies have reported the optimum pH for cellulase production by some
micro-organisms and are shown in Table 2 below.
Table 2: Optimum pH for cellulase production by microorganisms.
Organism pH Source
T. harzianum
T. viride
Bacillus subtilis
Cellulomonas spp
A.flavus MAM-F35
Aspergillus fumigatus
5.5
5.0
7.0-7.5
7.0
4.0
6.5
Ahmed et al. (2009)
Abo-State et al. (2010)
Ray et al. (2007)
Immanuel et al. (2006)
Abo-State et al. (2010)
Gilna and Khaleel (2011)
From Table 2 above, it is seen that the optimal pH for growth and cellulase production
varies from organism to organism. Most fungi prefer more acidic surroundings, about pH 4 to 6.
1.4.1.2 Temperature
Environmental temperature profoundly affects microorganisms, like all other organisms.
Indeed, microorganisms are particularly susceptible because their temperature varies with that of
the external environment (Willey et al., 2008). Generally, microorganisms are classified into four
groups based on their temperature tolerance. Psychrophiles grow well at 00C. However, their
optimum growth may be up to 150C or lower. Many species can grow at 0 to 7
0C though
optimally between 20 and 300C. These are called facultative psychrophiles or psychrotrophs and
spoil refrigerated foods. Mesophiles are microorganisms with growth optima around 20 to 450C.
Some others can grow at temperatures of 550C and above. They are called thermophiles. Table 3
below shows the optimum temperatures for growth and cellulase production in a few species of
micro-organisms.
Table 3: Temperature optima for cellulase production in microorganisms
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Organism Temperature (0C) Source
P. fluorescence
Bacillus circulans
B. subtilis
T. harzianum
Aspergillus fumigates
Aspergillus fumigatus
35
40
40
28
40
32
Bakare et al. (2005)
Ray et al. (2007)
Ray et al. (2007)
Ahmed et al. (2009)
Sherief et al. (2010)
Gilna and Khaleel (2011)
The temperature of the natural habitat of a cellulolytic organism can give a clue about its
possible temperature optimum for cellulase production. For instance, an organism isolated from
hot spring will require high temperature environment for maximum cellulase production.
1.4.1.3 Duration of incubation
This is a crucial factor in microbial cellulase production and growth. For an organism
growing by binary fission in a batch culture, the logarithm of the number of viable cells can be
plotted against duration or time of incubation to get a growth curve with four distinct phases.
During the lag phase, no growth occurs. But in the exponential or log phase, the microorganisms
are growing at the maximal rate possible depending on their genetic potential, nature of media and
other conditions of growth. Stationary phase is characterized by cessation of growth and
dormancy so that the curve becomes horizontal at that point. This may be due to depletion of
essential nutrients including oxygen (for aerobic organisms). It can also be as a result of
accumulation of toxic metabolites.
Depletion of oxygen in the case of aerobic organisms, can be remedied to some extent by
shaking the flask containing the culture. However, once a critical population is attained, the
microorganisms stop growing and it is expected that at this time, the batch culture be harvested.
The last phase is the death phase which is marked by decline in the number of viable cells.
Duration of incubation is really a factor to be considered in cellulase production and varies
from organism to organism and even among strains of the same organism. This is why many
authors have reported different durations at which peak enzyme activity was observed during
microbial cellulase production as shown in Table 4 below.
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Table 4: Optimal duration of incubation for microbial cellulase production
Organism Duration (Days) Source
Aspergillus niger
Aspergillus fumigatus
Bacillus coagulans
A. terreus
A. niger
T. harzianum
7
3
4
4
4
5
Sadaf et al. (2005)
Odeniyi et al. (2009)
Sherief et al. (2010)
Sherief et al. (2010)
Kang et al. (2004)
Ahmed et al. (2009)
From Table 4, it is seen that every organism has specific number of days or hours it should
be incubated for it to maximally produce cellulase. It is therefore necessary that a researcher
working with an organism whose optimal incubation period has not been ascertained yet, to carry
out an experiment first in that line.
1.4.1.4 Effect of metal ions on cellulase production
For many years heavy metals such as mercury, silver, arsenic, zinc and copper were used
as germicides more especially as bacteriostatic agents. Copper has proven to be an effective
algicide in lakes and swimming pools (Willey et al., 2008). Most heavy metals inactivate proteins
by reacting with their sulfhydril groups. They can precipitate cell proteins and act as protoplasmic
poisons at high concentrations.
Several microbial cellulases appear to be either activated or stabilized by the presence of
metals such as iron. While some metals are inhibitory to microbial growth, others are stimulatory.
Relatively high concentrations of Ca stimulate induction of cellulase production in the fungus
Trichoderma viride (Mandels and Reese, 1957). In some cases, a metal may act as a micronutrient
(micro-element) in an organism, meaning that high concentrations of it would be inhibitory to the
growth of the same organism. On the other hand, the same element may serve as macro-element
in another organism in which case high concentrations of it would be stimulatory to the growth of
the organism. Such microorganisms that grow in high concentrations of heavy metals are used in
bioremediation processes.
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1.4.1.5 Carbon source
In most fungal populations, the production of the enzyme cellulase is induced by the
presence of cellulose in the media or production broth (Suto and Tomita, 2001). The quality and
quantity of cellulose in a carbon source are important factors to be determined in cellulase
production so as to be able to make appropriate choice of the carbon source to use. A carbon
source made of almost pure cellulose like cotton will induce greater cellulase production.
Pretreated lignocellulosic material from which lignin has been removed will prove to be a better
source of carbon compared to native, untreated ones.
In some cases, composite carbon sources might give better yield of the enzyme than in the
case of single carbon source. The ability to utilize a carbon source maximally still lies in the
nature of the organism used. One organism may prefer a particular carbon source whereas another
may not. Several carbon sources have been employed in attempts to produce microbial cellulase.
Some of them are synthetic while others are natural. Abou-Taleb et al. (2009) used
carboxymethylcellulose, filter paper and cellobiose, and reported that carboxymethylcellulose was
the most effective sole carbon source for cellulase production in Bacillus alcalophilus S39 and
Bacillus amyloliquefaciens C23. The CMCase activities in the two organisms were reported as
1.8U/ml and 1.88U/ml and 0.86U/ml. -glucosidase activity was reported as 1.31U/ml and
1.41U/ml respectively. Narashimha et al. (2006) and Niranjane et al. (2007) compared
carboxymethylcellulose, cellulose powder and native carbon sources and found out that
carboxymethylcellulose is the best carbon source for cellulase production followed by cellulose
powder. Abo-State et al. (2010) studied cellulase production by two species of Aspergillus
(Aspergillus terreus Mam-F23 and Aspergillus flavus Mam-F35) using wheat straw as carbon
source. They reported that the enzymes produced showed CMCase, FPase and avicelase activities.
The highest avicelase activity was observed after twelve hours of incubation but the maximum
CMCase activity was observed after six days of incubation. Comparing cellulase production in the
two organisms with that of Trichoderma viride, they observed that the trend of cellulase
production in the three organisms using the same carbon source was almost the same. Ahmed et
al. (2009) compared the production of cellulase in Trichoderma harzianum using different carbon
sources including glucose, CMC, corn cobs, birchwood xylan and wheat bran. Although
researchers have attempted the use of corn cob as a substrate for cellulase production, none of
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them has investigated the possibility of cellulase production using the pulp of corn cob separated
from the hard outer part. Even though several carbon sources can be used in cellulase production,
it is still important that the carbon source to be used for industrial production of the enzyme be
sourced cheaply. This will help the economics, viability and sustainability of the industrial
process.
1.5 Structure of cellulase
Cellulases are modular enzymes that are composed of independently folding, structurally
and functionally discrete units referred to as either domains or modules (Henrissat et al., 1998).
Fungal EGs and CBHs have a two-domain structure. Specifically, a large catalytic core domain is
connected to a cellulose binding domain (CBD) with a flexible linker (Schulein, 1997). Lynd et
al. (2002) observed that one or two catalytic modules linked to one or several carbohydrate-
binding module is a frequent arrangement in aerobic microbes.
1.5.1 Catalytic domain (CD)
The active site in the catalytic core domain is divided into three distinct structures: a tunnel,
a cleft and a pocket. The substrate specificity for a given cellulolytic enzyme is often predicted
from the active site structure. The pocket and the tunnel shape characterize exo-acting enzymes,
because a chain needs to enter with one end first.
In the crystal structure of the predominant cellulolytic enzyme produced by T. reesei
CBHI, the active site is positioned in a long tunnel (Divne et al., 1994) as shown in Fig. 5 The
detailed structure of this long tunnel has been identified in cellooligomers and catalytic deficient
mutants of CBHI. Ten well-defined subsites for the cellulose chain have been found from
positions -7 to +3 with the active site being in positions -1 and +1 (Divne et al., 1998). The long
tunnel makes CBHI a very specific exo-acting enzyme that processes cellulose from one end of
the chain.
1.5.2 Linker
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The catalytic core and the cellulose-binding domain are connected via a flexible linker of
22-44 kDa long. Longer linkers have been reported by (Saloheimo et al. (1997). Because of the
flexible nature, it is difficult to obtain structural data on the linker. Therefore, the linker and CBD
are seldom depicted on 3D-models. The linker is rich in serine and threonine residues. In addition,
the linker is typically glycosylated (Harrison et al., 1998; Maras et al., 1999).
Fig. 5: Schematic representation of TrCBHI catalytic domain with a bound cellooligomer.
Secondary structure elements are coloured as follows: -strands, blue arrows; helices, red
spirals; loop regions, yellow coils. The cellooligomer is shown in pink as a ball-and-stick
object (Divne et al., 1998).
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The glycosylation is believed to provide rigidity and to ensure that the catalytic domain
and the CBD are correctly positioned next to each other (Srisodsuk et al., 1993).
1.5.3 The Cellulose binding domain (CBD)
The CBD binds to cellulose to bring the catalytic core in close contact with the cellulose
chain. This enhances the rate of hydrolysis. Removal of the CBD reduces hydrolytic activity by
reducing the number of interactions between cellulose and the enzyme (Tomme et al., 1988).
Removal of the CBD from CBHI (Cel 7A) and CBHII (Cel 6A) from T. reesei, for example,
reduced the adsorption of the catalytic core to bacterial microcrystalline cellulose by an order of
magnitude (Palonen et al., 1999). The CBHI CBD from Penicillium janthinellum bound to and
disrupted cotton fibres, and during hydrolysis the CBD showed synergy with an EG (Gao et al.,
2001). It is quite a surprise that although CBD has no active site, it has been shown to play yet
another role in the hydrolysis of cellulose, that is, non hydrolytic disruption of the structure of
cellulose.
CBDs are also called CBM (carbohydrate-binding module), a classification system used in
Carbohydrate Active enZymes (CAZy) database. CBM is defined as “a contiguous amino acid
sequence within a carbohydrate-active enzyme with a discrete fold having carbohydrate-binding
activity”. As with glycoside hydrolase (GH) classification scheme, CBMs are classified into
families based on amino acid sequence similarities and hydrophobic cluster analysis. Fungal
CBDs are all classified into the CBM I family, which contains proteins comprising 36 to 38 amino
acids in a wedge-like structure (Kraulis et al., 1989). The cellulose binding function is mediated
by aromatic residues on a flat surface in the CBD (Mattinen et al., 1998). Strikingly, the distance
between these aromatic residues matches the distance between repeating units in cellobiose
(Tomme et al., 1995). It has recently been suggested by (Mulakala and Reilly (2005) that the
CBM wedges itself under a free cellulose chain on the crystalline cellulose surface which then
“feed” into the catalytic domain of the cellulolytic enzyme.
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1.6 Mechanism of action of cellulase
All cellulases catalyze the cleavage of -1,4-glycosidic bonds. The reaction takes place via
general acid catalysis that requires two essential amino acid residues on the protein (Sinnott,
1990). In nearly all cases so far described these two residues are aspartates or glutamates (Davies
and Henrissat, 1995). The hydrolysis occurs via two major mechanisms giving rise to either
overall retention, or inversion of the anomeric configuration of the substrate. For enzymes that
perform catalysis with inversion, one catalytic residue is a proton donor which protonates the
glycosidic oxygen and thus promotes leaving group departure. The other catalytic residue acts as a
general base to activate the nucleophilic water by deprotonation. An oxocarbenium-ion-like
transition state is formed where electron density is distributed over no less than 8 atoms, and three
bonds are broken and formed almost simultaneously (Withers, 2001). For enzymes that catalyse
glycosidic bond cleavage with retention, these two residues consist of a catalytic nucleophile,
which forms a covalent glycosyl-enzyme intermediate, and a general acid/base residue. The
general acid/base residue first protonates the leaving group then, following the formation of the
covalent intermediate, activates the incoming nucleophile, often a water molecule.
The glycosyl-enzyme intermediate is formed and hydrolysed via an oxocarbenium ion-like
transition state. In addition to hydrolytic reactions, retaining enzymes can also perform
transglycosylation reactions under low water content or high substrate concentrations. In a
transglycosylation reaction, a glycosidic bond is usually broken and the glycosyl-enzyme
intermediate is formed. However, in contrast to hydrolytic reactions, an acceptor molecule other
than water is protonated to attack the glycosyl-enzyme intermediate to form a glycosidic bond.
Inverting enzymes lack the capability of performing transglycosylation reactions because it
requires the enzyme to be anomerically indiscriminate. For instance, if an acceptor such as a sugar
attacked the glycosidic carbon in a reaction which is catalyzed by an inverting enzyme, the
product would have a glycosidic bond with an inverted configuration. This is so because under
thermodynamically favoured conditions, any given reaction can proceed in the reverse direction
such that inverting enzyme would have to accept as substrate both and configurations.
1.6.1 Synergism in cellulose hydrolysis
In enzymatic reactions, synergism occurs when the combined activity of two enzymes
exceeds the sum of the individual enzyme activities. In the hydrolysis of filter paper, a one to one
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mixture of CBHI and EGI from T. reesei resulted in 23% (w/w) conversion whereas the individual
conversion was 6% and 8% using CBHI and EGI, respectively (Gama et al., 1998). This
synergism is explained by the well described endo-exo model (Stahlberg et al., 1993; Teeri,
1997). Exoglucanases make random cuts in the cellulose chain creating new starting points for the
progressive action of the CBHs. Figure 6.0 (I). The endo-exo model has been extended to the
obstacle model to account for synergism in the early phase of cellulose hydrolysis when there
should be sufficient free cellulose end for full CBH activity and therefore no synergism (Eriksson
et al., 2002b, Karlsson et al., 1999). In the obstacle model, CBHs remain adsorbed to the cellulose
chain due to tight binding between cellulose and the substrate binding site in tunnel of the CBH,
even though the CBH encounters an obstacle. Figure 6 (2A). Here, the CBH is not active before
the obstacle is removed. The obstacle can be an overlying cellulose chain (Valjamae et al., 1998).
Another model for synergism is the exo-exo model, in which synergism is found between
a CBH acting from the reducing end of cellulose chain and CBH acting from the non-reducing
end (Wood, 1985). The exo-exo synergism is less understood than the endo-exo model. However,
one explanation can be found in the obstacle model in which an overlying cellulose chain can be
removed by a CBH acting from the other end of the cellulose chain than the CBH being stuck on
the underlying chain. Figure 7(2A) (Mansfield et al., 1999).
Fig. 6: Model describing the degradation of cellulose by cellulases I. The traditional endo-exo
model. Endoglucanases make random cuts in the cellulose chain making new ends available for
CBHI
1
2A
2B
CBHII
-glucosidase
Cellobiose
Reducing end
Endoglucanase
Non-reducing
end
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the cellobiohydrolases (CBHI and CBHII) which cleave off cellobiose units from the ends of the
cellulose chains. -glucosidases act on cellobiose giving glucose (2A and 2B). The figure shown
above indicates an extension of the endo-exo model, namely, the obstacle model. In 2A, a CBHI
has been blocked by an overlapping cellulose chain (an obstacle). Due to the strong binding of
CBHI to the cellulose chain by the cellulose binding domain and the glycosyl binding sites in the
tunnel, the enzyme cannot be released from the cellulose chain and becomes unreproductively
bound. In 2B, an endoglucanase cuts the cellulose chain before CBHI reaches an obstacle, and
thereby preventing the unproductive binding of CBHI (Olsson et al., 2005).
1.7 General features of Aspergillus fumigatus
Aspergillus is a hyphomycetous genus with approximately 150 recognized species.
Members of the genus occur in a wide variety of habitats, but are especially common as
saprophytes in soils, stored food and feed products, and decaying vegetation in tropical and
subtropical regions (Domsch et al., 1980; Christensen and Tuthill, 1985). Some species are
economically important in the manufacture of fermented food, as sources of enzymes and in
chemical production (Klich and Pitt, 1992). There are also some species that produce potent
toxins that cause serious health effects.
Aspergillus fumigatus is a well – known member of the Aspergillus genus. Its life cycle
consists of two phases: a hyphal growth phase and a reproductive sporulation phase during which
secondary metabolites are produced depending on the material on which it grows. The organism
can survive on a variety of nitrogen sources and plays an essential role in carbon and nitrogen
recycling (Klich and Pitt, 1992).
For so many years, A. fumigatus was thought to only reproduce asexually, as neither
mating nor meiosis had ever been observed. In 2008, however, A. fumigatus was shown to possess
fully functional sexual reproductive cycle, 145 years after its original description by Fresenius
(O’Gorman et al., 2008). Its conidia is greyish turquoise or dark turquoise to dark green or dull
green with white mycelium; exudates when present, are uncoloured. Soluble pigment is usually
absent. According to Klich and Pitt (1992), its texture is velutinous to floccose, plane or radially
furrowed. Colony diameter varies depending on the media.
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The fungus is distinguished from other species by its rapidly growing colonies in turquoise
to dark green shades. It has phialides curving to be roughly parallel to each other and the axis of
the stipe. Its conidia is small (2-3µm) and borne in columns. Aspergillus fumigatus grows well
over a relatively wide temperature range and is ubiquitous (Klich and Pitt, 1992). It is a human
and animal pathogen, responsible for systemic mycoses usually resulting from invasion of the
lungs or respiratory tract. However, it is only immune-suppressed or immune-deficient individuals
that are susceptible to the fungal invasive infection which most commonly manifests as
aspergillosis. The fungus is capable of growth at 370C and up to temperatures of 50
0C with
conidia surviving at 700C. Its spores are ubiquitous in the atmosphere, and it is estimated that
everybody inhales several hundred spores each day but they are quickly eliminated by the immune
system in healthy individuals.
Aspergillus fumigatus is found in a wide range of substrates such as soils, plants, seeds,
sludge, wood chips, compost, cotton and even in penguin excreta (Klich and Pitt, 1992). It has a
stable haploid genome of 29.4 million base pairs. When fermentation broth of A. fumigatus was
screened, a number of indolic alkaloids with antimitotic properties were discovered. This points to
its potential in secreting anti-cancer substances.
1.7.1 Scientific classification
Kingdom: Fungi
Phylum: Ascomycota
Class: Eurotiomycetes
Order: Eurotiales
Family: Trichocomaceae
Genus: Aspergillus
Species Fumigatus
Source: Klich and Pitt (1992).
1.8 Rationale of the research
Production of ethanol, a multi-purpose chemical has long been a conventional process
involving the fermentation of sugar embedded in carbohydrate-rich human and animal food.
Sometimes, these essential foodstuffs are exported from developing countries for use in biofuel
industries in developed economies. The helpless masses are left without adequate food for
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survival. Conventional ethanol production from foodstuff is never a cheap process in a hunger-
stricken, populous world such as ours. The paradox in conventional ethanol is that it solves our
secondary problems but aggravates our primary problem of food inadequacy. We can generate
cellulosic ethanol from abundant lignocellulosic wastes by simple technologies. These
technologies involve the use of microorganisms to produce cheap cellulases employable in
hydrolyzing the sugars in cellulosic biomass. Corn cob is one of such cellulosic biomass which on
accumulation causes environmental pollution and even when burnt, generates CO2 implicated in
global warming. This research is therefore an attempt to use corn cob as a carbon source to
produce cheap cellulases which are starting material in cellulosic ethanol production. Apart from
advancing biofuel industries, cellulases are useful in textile, animal feed, detergent, food, pulp and
paper industries. Cellulase is used extensively in plant protoplast isolation (Ray et al., 2007).
1.9 Objectives of the research
i. To screen for a competent cellulose-degrading fungus from the local environment.
ii. To test its ability to adapt on media composed of corn cob pulp and the hard outer part
used as distinct carbon sources.
iii. Test the production of cellulase using the isolate on each of the two carbon sources.
iv. Characterize partially, the cellulases produced with respect to pH, temperature and heat
stability.
v. Assess which of the two carbon sources has a higher industrial potential.
CHAPTER TWO
MATERIALS AND METHODS
2.1 MATERIALS
2.1.1 Plant material
Mixed varieties of corn cobs were collected from the National Cereals Research Institute,
Yandev, Benue State. To complement this first batch of corn cobs, local varieties were gotten
from Agric Farms, University of Nigeria, Nsukka and Ogige Market Nsukka, Enugu State.
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2.1.2 Cellulolytic microorganism
A total of four microorganisms were isolated and screened for cellulase production but the
one used was the fungus, Aspergillus fumigatus, isolated from sewage sample obtained from the
University of Nigeria Central Sewage Treatment Plant.
2.1.3 Chemicals/Reagents
Ammonium chloride.(Fischer Scientific Company, USA).
Agar – Agar. (Hopkins and Wichans England).
Ammonium sulphate. (British Drug House (BDH) Chemicals Ltd. Poole, England).
Dipotassium hydrogen orthophosphate trihydrate. (Kermel,India).
Magnesium tetraoxosulphate(vi) heptahydrate. (Merck, Germany).
Sodium potassium tartarate (Rochelle salt) (Merck, Germany).
Sodium acetate. (Vickers laboratories Ltd, West Yorkshire, London).
Acetic acid. (May and Baker Ltd. England).
Sodium hydroxide. (Merck, Germany).
Folin-Ciocalteau Phenol Reagent. (Sigma-Aldrich, Germany).
Standard buffers pH 4.01, 7.01 and 9.0. (Fluka chemical company).
3,5-dinitrosalicylic acid (DNS). (Lab. Tech Chemicals, Avighkar India).
Hydrochloric acid concentrated.
Sabouraud dextrose agar (SDA)(Fluka chemical company,Germany).
Calcium carbonate. (M&B Laboratory Chemicals Dagenham England).
Sodium carboxymethyl cellulose. (British Drug House Chemicals Ltd Poole England).
Copper tetraoxosulphate(vi) pentahydrate. (Merck, Germany).
Bovine Serum Albumin (BSA)(Merck, Germany).
D (+) Glucose (Analar R) ( British Drug House Chemicals Ltd Poole England).
Lactophenol cotton staining blue. (Kermel chemicals, China).
Whatman No.1 Filter paper. (Whatman International Ltd, Maldstone England).
2.1.4 Equipment/Apparatus
Autoclave UDAY BURDON’S Patent Autoclave, Made in India.
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Centrifuge: Finlab Nigeria 80 – 2B.
Incubator: B & T Trimline Incubator.
Magnetic Stirrer: AM – 3250B Surgifriend Medicals, England.
pH Meter: Model PHS – 3C, Search Tech. Instruments.
Sensitive Weighing Balance: B2404 – 5 Mettler Toledo Made in Switzerland.
UV/Visible Spectrophotometer: Jenway 6405.
Water bath: Model DK .
Pressure Pot: MC-PC12000D Master Chef.
Weighing Balance: Ohaus Dial – O – Gram. Ohaus Corporation, N.J. USA.
Light Microscope: Union Tokyo. No. 35122
2.2 METHODS
2.2.1 Processing of corn cobs
All the corn cobs were sun-dried for three days and then broken to separate the hard outer
part from the soft inner part (the pulp). Each was milled separately and the hard outer part was
labeled CC-Outer while the pulp was labeled CC-Inner.
2.2.2 Isolation of cellulolytic microorganisms
2.2.2.1 Collection of soil samples
Five samples were collected from different sites. The first three were soil samples from
decomposing saw dust (DSD) at Nsukka timber shade, decomposing litter behind Mbanefo
Students’ Hostel (DL – MSH) and Garden Soil behind Mbanefo Students’ Hostel, University of
Nigeria, Nsukka (GS – MSH). The last two were samples from the University of Nigeria Central
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Sewage Treatment Plant. They were sewage water and sewage sediment labelled as SW –UNN
and SS-UNN respectively. All the samples collected were transferred to the laboratory.
2.2.2.2 Determination of soil pH
The pH was standardized using standard buffers of pH 4.01, 7.01 and 9.02 as specified by
the manufacturer according to the method of Forster (1995) and is based on the potentiometric
determination of hydrogen ion activity in soil suspended in water.
A known amount (10g) of each of the soil samples was weighed into a clean dry beaker
and 20ml of distilled water was added to each, stirred vigorously to get a homogenous suspension
which was allowed to equilibrate properly in water. The suspension was again stirred and the
supernatant was obtained by means of decantation for the pH measurement.
2.2.2.3 Preparation of soil extract enrichment
Ten grams of the garden soil was weighed into a clean dry beaker and 20 ml of distilled
water was added. This was followed by the stirring and filtration through Whatman No.1 filter
paper. The filtrate was used to enrich the broth by supplying the minerals necessary for the growth
of microorganisms in their natural environment.
2.2.2.4 Preparation of mineral salt medium for isolation
A weight of 0.4g each of NH4Cl, K2HPO4.3H2O and MgSO4.7H2O was weighed into a
beaker containing 400ml of distilled water. A known volume (100 ml) of it was measured into
each of four labelled 250ml Erlynmeyer flasks. A measured volume (2.5 ml) of soil extract
enrichment (see section 2.2.2.3) was added to each of the labeled flasks containing the minerals.
After thorough mixing, their pH was adjusted to 7.0 by drop-wise addition of 0.1M sodium
hydroxide solution. Equal number of filter paper strips were shredded and introduced into each
flask and plugged with aluminium foil-wrapped cotton wool. The four flasks were autoclaved at
121oC at 15 pounds per square inch (psi) for 15 min and allowed to cool.
2.2.2.5 Inoculation of the first isolation broth
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Five milliliters of each of the supernatants from the soil samples from DSD and DL-MSH
prepared in section 2.2.2.2, was transferred to the appropriate autoclaved flasks following the
labels on them. Five milliliters each of the sewage water and sewage sediment was also used to
inoculate appropriate flasks. All the inoculated flasks were wrapped with carbon paper and kept in
a dark cupboard at room temperature for fourteen days. Penetration of light was avoided by
keeping them in the dark so as to prevent algal growth.
2.2.2.6 Sub-culturing into another isolation broth
Another mineral salt broth was prepared exactly as before and 2.5 ml of soil extract
enrichment was obtained from the remaining of the earlier garden soil air-dried. Mineral salts
medium was prepared as in section 2.2.2.4 but this time 95 ml of the mineral broth was measured
into each of four labelled Erlynmeyer flasks and sterilized. After autoclaving and cooling, 5 ml
from the fourteen-day cultures were used to inoculate the freshly prepared broth and incubated for
14 days just as in section 2.2.2.5.
2.2.2.7 Sub-culturing on solid media (SDA)
The solid medium was Sabouraud Dextrose Agar medium which is a selective medium for
isolating fungi. A quantity (6.2g) of SDA was dissolved in 100ml of distilled water as specified by
the Manufacturer. The 250 ml Erlynmeyer flask containing the mixture was plugged with cotton
wool covered with aluminium foil and sterilized for 20 min in a pressure pot. It was allowed to
cool to 50oC before the content was poured into four labelled sterile petri dishes and allowed to
cool completely, ready for inoculation.
2.2.2.8 Inoculation of plates
A sterile wire loop was dipped into the Erlynmeyer flask, then removed and used to
inoculate the media by streaking method. The plates were incubated at 37oC until visible colonies
were observed, thereafter successive sub-culturing was done by identifying fungal colonies with
different morphological features and plating them on separate plates. This process was continued
until pure fungal cultures were obtained.
2.2.2.9 Storage on SDA slants
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Sabouraud Dextrose Agar medium was prepared as in section 2.2.2.7, aseptically poured
into sterile test tubes, capped and slanted to cool at room temperature. A sterile wire loop was
used to inoculate the slants with the pure fungal isolates. The inoculated slants were incubated at
37oC. All the fungi grew well on SDA.
2.2.2.10 Macroscopic photographs of the fungal isolates
Three-day old cultures of the four fungi isolated but now grown on SDA, were observed.
Colour, nature of spores and growth patterns were noted. Snapshots of the cultures were equally
taken.
2.2.2.11 Culturing of the isolates on corn cob media
The media were composed of 0.1% w/v of each of the following salts: MgSO4.7H2O,
K2PO4.3H2O and NH4Cl. A quantity (1.0% w/v) of CC – Inner was added to obtain CC – Inner
Media, and 1.0% w/v of CC – Outer was added to obtain CC – Outer media. To each of the
media, 1.5g/100 ml (i.e 1.5% w/v) agar – agar (the gelling agent) was added. The two separate
media were sterilized for 20 min using pressure pot. They were allowed to cool to 50oC and
poured into plates which later were inoculated with the pure fungal isolates using sterile wire
loop. The plates were incubated at 37oC. Since only one of the four fungal isolates grew very well
on both CC – Inner and CC – Outer media, it was preserved on their respective slants.
2.2.3 Method of protein determination
The method of Lowry et al. (1951) was adopted for protein determination in both crude
and partially pure cellulases. A volume (0.1ml) of each enzyme sample was made up to 1.0 ml
using 0.2M sodium acetate buffer pH 5.5. Five milliliters of alkaline solution (Solution D
prepared as in appendix iv) containing sodium carbonate, sodium hydroxide and sodium tartarate,
was added and left to stand for 10 min at room temperature. A volume (0.5 ml) of diluted Folin
Ciocalteau Phenol reagent (Solution C ) was added with immediate mixing. Absorbance was
taken at the wavelength of 750nm after allowing a time course of 30 min.
2.2.3.1 Protein calibration curve
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Ten test tubes were arranged in duplicate containing 0.0 to 1.0 ml of 2 mg/ml protein stock
solution. The volumes were made up to 1 ml using 0.2M sodium acetate buffer pH 5.5. Five
milliliters of the alkaline solution D was added and left to stand for 10 min at room temperature. A
volume (0.5 ml) of solution C (diluted Folin Ciocalteau Phenol Reagent) was added and mixed
immediately. A time course of 30 min was allowed before the absorbance was taken at the
wavelength of 750 nm. A standard curve of absorbance against standard BSA concentration was
generated and calculation of protein concentration per ml in each of the test tubes was done using
the equation of straight line: Y = 0.8038x. Where Y is the absorbance and x is the protein
concentration in mg/ml
2.2.4 Method of glucose determination
The method of Miller (1959) was adopted for glucose determination. This involved
incubating a volume (0.1 ml) of each enzyme sample with one filter paper strip (1cm x
3cm) for 6hr at 500C in the presence 0.2M sodium acetate buffer pH 5.5. A volume (2.4 ml) of
the buffer was used. DNS reagent (0.5 ml) was added to the reaction mixture after the 6hr
incubation and mixture boiled for 10 min. Absorbance was then taken at 540nm when the
mixture had cooled to room temperature.
2.2.4.1 Glucose calibration curve
Nine test tubes were arranged in duplicates containing 0.0 to 2.4 ml of 5mM glucose stock
solution. The volumes were made up to 3 ml with 0.2M sodium acetate buffer pH 5.5. Then, 0.5
ml of DNS (3,5 – dinitrosalicylic acid) reagent was added with shaking and the reaction mixture
boiled for 10 min and allowed to cool so that absorbance was taken at 540 nm. A standard curve
of absorbance against standard glucose concentration was generated. Glucose concentration was
calculated per ml in each test tube using the formular: Y = 0.0004x2. Where Y is the absorbance
and x2 is glucose concentration in μmoles/ml.
2.2.5 Sub-merged fermentation experiments
2.2.5.1 The fermentation broth
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The broth used for fermentation in the case of CC-Inner contained 1% w/v CC – Inner,
0.1 % w/v MgSO4. 7H2O, NH4Cl, and K2HPO4 in distilled water. The broth (50 ml) was measured
out into clean 250 ml Erlynmeyer flasks before the carbon source was added. Exactly the same
was also done in the case of CC – Outer except that the CC – Inner was replaced with CC – Outer.
All the flasks were then stoppered and sterilized for 20 min in a pressure pot.
2.2.5.2 Inoculation of the broth
Fresh cultures were prepared by preparing fresh plates as described in section 2.2.2.4, and
inoculating them. When the organism had covered the plates evenly, it was used to inoculate the
flasks. A sterile 6.0 mm2 cork borer was used to inoculate the sterile broth. All inoculated flasks
were left to incubate at room temperature.
2.2.5.3 Harvesting the fermented broth
Each day of harvest, appropriate flasks were selected and the contents were filtered
through Whatman No. 1 filter paper. The filtrate collected was used for cellulase activity assay.
Protein determination was also carried out on the filtrate. This was a pilot study to determine the
actual day of peak cellulase production.
2.2.6 Cellulase assay
Cellulase activity was assayed by the determination of reducing sugar released from filter
paper strips. A volume of 0.1 ml enzyme was incubated with one strip of filter paper in 0.2M
sodium acetate buffer at 500C for 6 hr. The reducing sugar produced was assayed by the
dinitrosalicylic acid (DNSA) method (Miller, 1959), using glucose as the sugar standard. One unit
of cellulase was defined as the amount of enzyme which produced one micromole glucose per
minute under the assay conditions.
2.2.6.1 Preparation of filter paper strips
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Filter paper strips of dimensions 1cm x 3cm were cut out and used as substrate for the
cellulase assay. By convention, each 1cm x 3cm filter paper strip contains an equivalent of 25mg
of glucose.
2.2.6.2 Cellulase assay mixture
Two milliliters of 0.2M sodium acetate buffer pH 5.5, was dispensed into the test tube and
0.5 ml of the filtrate was added. A strip of filter paper was submerged in the mixture and
incubated at 500C for six hr. An enzyme blank was prepared in the like manner but was devoid of
the filtrate (enzyme). Also, substrate blank was prepared and contained 0.5 ml of the filtrate
(enzyme), 2.0 ml of buffer but no substrate was added to it. At the completion of the incubation
time, the test tubes were removed and 0.5 ml of DNS reagent was added. After boiling for 10 min
and cooling, absorbance was read at the wavelength of 540 nm. The blank was used to zero the
spectrophotometer. The reducing sugar formed per ml was calculated from the glucose calibration
curve.
2.2.7 Protein assay mixture
Sodium acetate buffer pH 5.5 (0.9 ml, 0.5 ml) was pipetted into separate test tubes and the
volumes made up to 1ml with the filtrate. Alkaline solution D (5 ml) was added and the reaction
allowed to stand for ten minutes at room temperature. A volume of 0.5 ml solution C (diluted
Folin Ciocalteau Phenol Reagent) was added with rapid mixing and absorbance was taken at 750
nm after 30 min. The blank used to zero the instrument contained O ml of the filtrate. All these
(cellulase activity and protein determination) were carried out on crude enzymes from pilot study
as well as those from mass production before partial purification was done.
2.2.8 Partial purification of the crude enzyme filtrate
2.2.8.1 Ammonium sulphate saturation and centrifugation
A total of 1 litre of each of the crude enzyme filtrate was subjected to 50% ammonium
sulphate saturation by dissolving 291g of the salt in the cold filtrate until the salt became
completely dissolved. The solution was then stored at 40C. Centrifugation was carried out at 3,000
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rpm for 30 min and the precipitate was re-dissolved in the sodium acetate buffer pH 5.5 and stored
at 40C.
2.2.8.2 Dialysis
Dialysis bag / tube preserved in 90% ethanol was used. It was first rinsed severally with
distilled water till it was free of ethanol. A volume (15 ml) of the cellulases was each dialysed
against 300 ml of the sodium acetate buffer for 12 hr with constant stirring via magnetic stirrer at
40C. The buffer was changed after seven hr. The partially purified enzyme was stored at 4
0C.
2.2.8.3 Studies on partially purified cellulase samples
Two cellulase samples obtained from the partial purification were used for further studies
which included the effect of pH and temperature on cellulase activity. Then the heat-stability of
the enzymes was also investigated. These two enzymes secreted by the organism using CC-Outer
and CC-Inner as the carbon sources had earlier been labelled CC-OUTER and CC-INNER
respectively. Protein determination and cellulase activity were also carried out on the partially
pure cellulases as in sections 2.7 and 2.6 respectively
2.2.8.3.1 Effect of pH on enzyme activity
A volume (0.1 ml) each of the partially pure cellulases (CC-OUTER and CC-
INNER) was incubated with a strip (1cm×3cm) of filter paper in the presence of 0.2M sodium
acetate buffer of varying pH for 6 hr at 500C and absorbance was read at 540nm. The pH range
covered was 3.5-9.0. The concentration of glucose released from the assay showed the optimum
pH of the enzymes.
2.2.8.3.2 Effect of temperature on enzyme activity
A volume (0.1 ml) of CC-OUTER, one strip of the same dimension of filter paper and 2.4
ml of acetate buffer (pH 5.5) were incubated for 6 hr at various temperatures: 25, 30, 35, 40, 45,
50, 55, 60, 65 and 700C. The concentration of glucose released in each case was determined by
DNSA method and the optimum temperature of the cellulase was deduced. The second enzyme
sample (CC-INNER) was subjected to the same treatment and its optimum temperature was
equally determined.
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2.2.8.3.3 Heat-stability study
A volume (0.1 ml) of CC-OUTER was incubated alone for one hour at each of the
following temperatures: 35, 40, 45, 50, 55, 60, 65 and 700C and cooled for 30 mins before
incubated with the substrate (filter paper) and appropriate buffer for 6 hrs at 500C. The
concentration of glucose released in each case was then determined by DNSA method. The same
treatment was also carried out on the second cellulase sample.
2.2.9 Fungal identification
Partial microscopic identification was done by staining a tuft of the mycelia of the
organism on microscopic slide using lactophenol staining blue. The identification was done by
placing the slide on the microscope matching the image on it with the one on Colour Atlas of
Diagnostic Microbiology.
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CHAPTER THREE
RESULTS
3.1: Pilot study for determination of day of peak enzyme production
Cellulase production by Aspergillus fumigatus grown on CC-Inner, increased steadily from
day 1 until a peak was reached on the 3rd
day (4.083Units). Thereafter, the enzyme production
became static till day 6 before declining as shown in Fig. 7. Cellulase production by the same
organism grown on CC-Outer, showed a more gradual increase from day 1 and reached a peak
(3.819Units) on the 4th
day. Beyond the 4th
day, a steady decrease in the enzyme production was
observed as shown in Fig. 8. Similar trends were also observed when the specific activities of the
two cellulases were plotted against duration of incubation as shown in Figs. 9 and 10.
Fig. 7: Pilot study for determination of day of maximum enzyme production using CC-Inner
(Total activity in micromoles/min).
0 1 2 3 4 5 6 7 8
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Fig. 8: Pilot study for determination of day of maximum enzyme production using CC-outer as
carbon source. (Total activity in micromoles/min).
0 1 2 3 4 5 6 7 8 9
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Fig. 9: Pilot study for determining the day of maximum enzyme production using CC-Inner as
carbon source. (Specific activity in micromoles/min/mg protein).
0 1 2 3 4 5 6 7 8 9
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3.2 Studies on the crude enzymes.
3.2 Studies on mass-produced crude cellulases
3.2.1 Extracellular protein secretion
Fig. 10: Pilot study for determining the day of maximum enzyme production using CC-
outer as carbon source. (Specific activity in micromoles/min/mg protein).
0 1 2 3 4 5 6 7 8
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Aspergillus fumigatus also secreted higher amount of extracellular protein during larger
scale production of cellulase when grown on CC-INNER (0.164 mg/ml) than on CC-OUTER
(0.112 mg/ml) as shown in Fig. 11. The assay was carried out on the crude cellulase samples just
before ammonium sulphate saturation.
Fig. 11: Comparison of extracellular protein secreted by the organism using CC-Outer and CC-
Inner as substrates. The protein assay was done in duplicates.
3.2.2 Cellulase activities of crude CC-OUTER and CC-INNER
Carbon source
used
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Total cellulase assay carried out on crude sample of the mass-produced CC-INNER showed a
higher activity (26.111Units) than that obtained from crude sample of the mass-produced CC-
OUTER (14.861Units). This is shown in Fig. 12.
Fig. 12: Comparison of total cellulase activities of crude CC-OUTER and CC-INNER. Total
cellulase activity was given in Units. (Units = micromoles/min).
3.3 Studies on partially purified cellulases
3.3.1a Purification for CC-OUTER
Carbon source used
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Purification for CC-OUTER, showing the total proteins, total activities, specific activities
and purification factors of the crude and partially purified enzyme obtained after 50% ammonium
sulphate saturation and dialysis is shown in Table 5. The specific and total activities of the
dialyzed enzyme were higher than those of the crude and un-dialyzed enzymes. A twenty-nine-
fold purification was achieved in the dialyzed enzyme.
Table 5: Purification for CC-OUTER
procedure Total
protein
(mg)
Total
activity
(units)
Specific
Activity
(units/mg)
Purification
factor
Crude cellulase filtrate
Undialyzed ppt at 50% salt
Dialyzed ppt at 50% salt
56
4.14
5.964
14.861
13.056
45.833
0.265
3.154
7.685
1
11.190
29
Ppt. : precipitate
3.3.1b Purification for CC-INNER
Purification for CC-INNER, showing the total proteins, total activities, specific activities
and purification factors of the crude and partially purified enzyme obtained after 50% ammonium
sulphate saturation and dialysis is shown in Table 6. The specific and total activities of the
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dialyzed enzyme were higher than those of the crude and un-dialyzed enzymes. Approximately
two -fold purification was achieved in the dialyzed enzyme.
Table 6: Purification for CC-INNER
procedure Total
protein (mg)
Total
activity
(units)
Specific
Activity
(units/mg)
Purification
factor
Crude cellulase filtrate
Undialyzed ppt at 50% salt
Dialyzed ppt at 50% salt
82
11.24
14.882
26.111
13.472
92.917
3.18
1.20
6.24
1
0.377
1.960
Ppt. : precipitate
3.3.2 pH profile of the partially purified cellulases
3.3.2a The pH profile of partially purified CC-INNER
The pH profile of the dialyzed CC-INNER showed a sudden rise in cellulase activity
between the pH 4.0 (17.778 Units) and pH 5.0 (79.583 Units). The optimum activity of the
enzyme was obtained at pH 6.0 (88.819 Units) as shown in Fig. 13.
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Fig. 13: Optimum pH of the partially purified CC-INNER. The assays were carried out in
duplicates. (Units = micromoles/min).
3.3.2b The pH profile of partially purified CC-OUTER
The pH profile of the dialyzed CC-OUTER also showed a sudden rise in cellulase activity
that was between pH 5.0 (9.306 Units) and pH 6.0 (46.181 Units). The enzyme is a neutral
cellulase and gave optimum activity (57.153 Units) at pH 7.0 as shown in Fig. 14.
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Fig. 14: Optimum pH of the partially purified CC-OUTER. The assays were carried out in
duplicates.
3.3.3 Optimum temperature
3.3.3a Optimum temperature for CC-INNER
The temperature profile of dialyzed CC-INNER showed an increase in cellulase activity as
temperature increased from 25 to 50˚C Optimum activity (278.667 Units) was obtained at 55
0C as
shown in Fig. 15. Above the optimum temperature, activity declined sharply.
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Fig. 15: Optimum temperature of the partially purified CC-INNER.
3.3.3b Optimum temperature for CC-OUTER
The temperature profile of the dialyzed CC-OUTER also showed an increase in activity
with increasing temperature. Fig. 16 shows that the enzyme has optimum temperature of 50oC at
which a peak activity of 43.333 Units was observed. Thereafter, there was a steady decrease in
activity.
.
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Fig. 16: Optimum temperature of the partially purified CC-OUTER.
3.3.4 Heat stability studies
3.3.4a Heat stability study on CC-INNER
The heat stability study on the enzyme as depicted in Fig. 17, revealed a triphasic pattern.
In the first phase (35-50oC), it was relatively stable whereas the second phase (50-60
oC) showed a
pronounced decrease in stability. The last phase (60-70oC) also showed a decrease in stability of
the enzyme to heat but the decrease was not as sharp as in the second phase.
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Fig. 17: Heat stability of the partially purified CC-INNER
3.3.4b Heat stability on CC-OUTER
study conducted on CC-OUTER showed an enzyme that was rather stable when stored
between the 35 and 45oC for one hour. Its stability however, dropped between 45 and 50
oC but
was not lowered between 50 and 55 o
C. Beyond 55 o
C, the stability of the enzyme continued to
decline as shown in Fig. 18.
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Fig. 18: Heat stability of the partially purified CC-OUTER. The assay was done in duplicates.
3.3.5 Percentage loss in the activities of CC-INNER and CC-OUTER from 35-
70˚C
Figs. 19 and 20 show the percentage losses in the activities of CC-INNER and CC-
OUTER when stored between 35-70 ˚C for 1 hr. In the case of CC-INNER, activity losses of
9.975% and 26.378% were observed at 50 and 55˚C respectively. But at 70 ˚C, as much as
58.137% loss in activity was observed. The loss in activity of CC-OUTER was not as consistent
as that of CC-INNER. However, loss in its activity became significant from 60˚C, and at 70˚C as
much as 45.854% loss in activity was observed.
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Fig. 19: Percentage loss in CC-INNER activity under varying temperatures (35 - 70oC). The assays
were carried out in duplicates and mean values used.
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Fig. 20: Percentage loss in CC-OUTER activity under varying temperatures (35 - 70oC). Assays were
carried out in duplicates and mean values used.
3.4: Macroscopic photographs of isolated fungi
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Plates 1-4 are the macroscopic photographs four fungi isolated from sewage water. Each
plate shows a three-day old culture of the fungus specified as grown on SDA (top view). The
fungus on Plate 3 is Aspergillus fumigatus and was the organism used in cellulase production.
Plate 1: Aspergillus niger grown on Sabouraud Dextrose Agar.
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Plate 2: Aspergillus spp grown on Sabouraud Dextrose Agar.
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Plate 3: Aspergillus fumigatus grown on Sabouraud Dextrose Agar.
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Plate 4: Rhizopus spp grown on Sabouraud Dextrose Agar.
CHAPTER FOUR
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DISCUSSION
The pH of most of the samples from which the organisms were isolated were slightly
alkaline. The two sewage samples SS-UNN and SW-UNN had pH values of 7.33 and 7.37
respectively while DSD gave a slightly acidic pH of 6.05. The garden soil (GS-UNN) which was
used for enrichment was neutral (7.02) while DL-MSH gave a pH value of 7.61.
The alkalinity observed in the two sewage sites could be the reason sewage is used as
organic manure especially in growing vegetables though it has been recently found to contain high
amounts of heavy metals. The neutrality observed in the garden soil could be as a result of balance
in nutrients. In soil ecology, sets of microorganisms carry out the degradation of materials and the
pH of the soil or site depends on the population of organisms at work at the prevailing moment.
One set initiates degradation, another continues while the last completes the natural process. Some
of these organisms release organic acids as end products which affect the soil pH.
The results of the pilot study on cellulase production using the two distinct parts of corn
cob, showed varied durations of time for enzyme production even though the same organism was
used. The crude CC-OUTER showed peak cellulase activity on the 4th
day (3.82 µmole/min)
while the crude CC-INNER had its peak activity on the 3rd
day (4.083µmole/min). Cellulase
activity of CC-INNER is slightly higher than that observed in CC-OUTER as shown in Figs.7
and 8. Sherief et al. (2010) investigated cellulase production from Aspergillus fumigatus using
both single and mixed substrates. The result of their study on mixed substrate of rice straw and
wheat bran, showed peak CMCase, endoglucanase and β-glucosidase activities on the 4th
day.
This finding is in agreement with the work of Sherief et al. (2010). Variation in the case of CC-
INNER may be attributed to the nature of the substrate used. The fact that the organism secreted
higher amount of cellulase when grown on CC-Inner than when grown on CC-Outer suggests that
the corn cob pulp is a better substrate for cellulase production than the outer part.
The same study by Sherief et al (2010) reported the activities of cellulase from A.
fumigatus grown on different carbon sources including alfa alfa, corn cob, saw dust, wheat straw,
rice straw and wheat bran. Corn cob and sawdust proved to be the poorest among the substrates
investigated in supporting cellulase secretion. This poor cellulase secretion could be because the
corn cob was used whole. Most of the previous works on cellulase production from Aspergillus
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fumigatus using corn cob have been on corn cob as a whole, and not on the distinct parts of the
cob.
Several factors affect the extracellular secretion of cellulase during fermentation causing
either increase or decrease in extracellular enzyme activity. The depletion of micro- and macro-
nutrients in the medium as fermentation progresses could limit the potentiality of the organism.
Some toxic metabolites secreted into the medium by the same organism could also result in a
change in the pH of the medium. This in turn could have a critical effect on cellulase production.
Glucose, the end product of cellulase action, is a potent catabolite repressor of cellulase
biosynthesis (Ilmen et al., 1997; Zhang and Lynd, 2005). Cellobiose inhibits both endoglucanase
and β-glucosidase (Ojumu et al., 2003).
All through the work, the cellulase activity and total protein obtained for CC-OUTER
were always lower than those obtained with CC-INNER. This again suggests that CC-Outer did
not support cellulase production as much as CC-Inner. The outer part of the corn cob is very hard
compared to the inner part (the pulp) which is very soft and light. CC-Outer is probably rich in
lignin since hardness of plant materials is associated with lignifications which are also associated
with age of the plant material. Lignin shields the cellulose and makes it inaccessible to hydrolytic
action of enzymes.
Partially purified cellulases from CC-Outer (that is, CC-OUTER) maintained a fairly low
activity over pH range of 3.5 and 5.0 after which the activity increased sharply as shown in
Fig.14. This increase in activity was sustained over the pH range of 6.0 and 9.0 with an optimum
pH of 7.0, suggesting a neutral cellulase. This finding agrees with that of Immanuel et al (2006).
Ray et al (2007) also reported neutral cellulase. More so, Abdelnasser and El-diwany (2007) were
able to isolate from a thermophilic bacteria, cellulases with peak activity at pH 7.0. These reports,
however, disagree with that of Abo-State et al (2010). They carried out a work on cellulase
production using Aspergillus terreus Mam-F23 and Aspergillus flavus Mam-F35 and reported
optimum pH of 4.5 and 4.0 for CMCase and Fpase activities respectively. These were for
Aspergillus terreus. In the case of A. flavus, they reported pH optima of 4.0 for both CMCase and
Avicelase activities.
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The partially purified enzyme, CC-INNER maintained a fairly low activity over pH range
of 3.5 and 4.0. Thereafter, a sudden increase in activity was also observed. Between the pH range
of 5.0 and 9.0, it showed high activities with a peak at pH 6.0. This is seen in Fig.13. This finding
agrees with some previous works. Lowe et al (1987) reported optimal pH of 6.0 for both CMCase
and FPase. Ponpium et al (2000) also reported a cellulolytic multienzyme complex with pH
optima of 6.0. However, Gilna and Khaleel (2011) reported pH optima of 6.5 for a strain of A.
fumigatus. This disagrees with the current findings and those of Ahmed et al. (2009). They
produced cellulases from the fungus Trichoderma harzianum and reported optimum pH of 5.5 for
exoglucanase, endoglucanase and β-glucosidase. Coral et al. (2001) reported cellulase from
Aspergillus niger, with CMCase activity over a broad range of pH but having peak activities at pH
4.5 and 7.5. These multiple peaks and hence variation from our finding could be as a result of
isoforms of the same enzyme being present but acting maximally at different pH. It may also be
traced to difference in the species of organisms used. The fairly low activity observed in CC-
OUTER over the low pH range of 3.5 to 5.0, and in CC-INNER over the pH range of 3.5 to 4.0
could probably be due to deprotonation of the catalytic amino acid residues in the active and
binding sites of the enzyme protein leading to marked reduction in activity. This might explain
why there was a sudden rise in activities of the enzymes as pH increased.
One of the factors that have profound effect on the biological activity of proteins including
enzymes is pH. Its alteration could denature an enzyme changing its three-dimensional structure
as well as its active site configuration or ionic state which has great relevance to its activity. This
is because electrostatic bonds contribute to the maintenance of the three-dimensional structure of
native proteins so that a change in pH can lead to unfolding and dis-orientation of the polypeptide
chains.
The two enzyme samples showed increase in activity with increasing temperatures but
with temperature optima between 50oC and 55
oC. For CC-OUTER, the optimum activity was
observed between at 50oC while CC-INNER showed peak activity at 55
oC as shown in Figs. 16
and 15 respectively. At temperatures above 55oC activity decreased steadily up to 70
oC. Lowe et
al (1987) reported temperature optimum of 50oC for CMCase and β-glucosidase activities but
45oC for FPase activity. Several researchers have equally reported different temperature optima.
While Nipa et al (2006) reported optima of 40oC as well as Ray et al (2007), Haq et al (2005)
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reported 30oC. Immanuel et al (2007) reported 40
oC for cellulase from a strain of Aspergillus
fumigatus whereas Gilna and Khaleel (2011) reported 32oC as temperature optima for cellulase
produced by another strain of A. fumigatus. These differences could be attributed to differences in
the strain and species of organisms used in the enzyme production. Strains of the same organism
isolated from different sources synthesize proteins (enzymes) with varying characteristics.
Temperature like pH, is one of the critical factor affecting enzyme-catalysed reactions.
Like other chemical reactions, the rate of an enzyme-catalyzed reaction increases with modest
increase in temperature. This is true only over a strictly limited range of temperatures. When the
temperature of a reaction is raised, there is sufficient energy to overcome the energy barrier and so
cause an increase in the number of collision between the enzyme involved and its substrate. The
result is an increase in the rate of the reaction so as to reach its maximum. Beyond this optimum
temperature, every further increase in temperature introduces vibrational energy that weakens the
three-dimensional structure of the enzyme. Once the hydrogen bonds and hydrophobic bonds
holding the native structure together are broken or disrupted, the enzyme is denatured and the
reaction stops. The temperature range over which any enzyme is stable and catalytically active
depends on the temperature of the cell in which the enzyme is found.
Temperature does not affect enzyme activity only during temperature-activity assay but
also affects the enzyme protein on storage. This was corroborated by assays carried out to
determine the heat or thermal stability of the two enzyme samples. Processes that disrupt or
unfold protein structures make them lose their activity through denaturation. Extremes of
temperature apart from causing a loss of activity of water-soluble proteins might also precipitate
them out of solution. Therefore, the thermal stability of an enzyme refers to the degree to which
the secondary, tertiary and quarternary structures are affected by temperature changes. This
reflects in an enzyme’s activity after its exposure to heat especially in absence of its substrtate.
The heat stability study on CC-INNER as shown in Fig. 17, indicated a triphasic pattern.
In the first phase (35-50oC), it was relatively stable whereas the second phase (50-60
oC) showed a
pronounced decrease in stability. The last phase (60-70oC) also showed a decrease in stability of
the enzyme to heat but the decrease was not as sharp as in the second phase. On the other hand,the
same study conducted on CC-OUTER showed an enzyme that was rather stable when stored
between the 35 and 45oC for one hour. Its stability however, dropped between 45 and 50
oC but
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was not lowered between 50 and 55 o
C. Beyond 55 o
C, the stability of the enzyme continued to
decline as shown in Fig. 18.
Though the two enzymes were observed to be stable between 35oC and 40
oC within the
duration allowed by the study, but as the temperature increased to 60 o
C, a steady decline in
activity was the result.
Figs. 19 and 20 show the percentage losses in the activities of CC-INNER and CC-
OUTER when stored between 35 and 70 ˚C for 1 hr. In the case of CC-INNER, activity losses of
9.975% and 26.378% were observed at 50 and 55˚C respectively. But at 70 ˚C, as much as
58.137% loss in activity was observed. The loss in activity of CC-OUTER was not as consistent
as that of CC-INNER. However, loss in its activity became significant from 60˚C, and at 70˚C as
much as 45.854% loss in activity was observed. These losses in activities of the two enzymes
substantiates the fact that storage temperature affects the stability and hence, the activities of
enzymes on storage. However, the temperature of an organism’s natural habitat determines the
nature of the enzymes or proteins it produces. A thermophile will usually secrete a thermo stable
enzyme. Ponpium et al (2000) reported a cellulosome-type multienzyme complex from a strain of
thermophilic bacteriod species with optimum temperature of 60oC. Abdelnasser and El-diwany
(2007) also reported cellulases with optimum temperature of 75oC, produced by thermophilic
bacteria from Egyptian hot spring.
The temperature of the natural habitat of the organism used in cellulase production affects
the thermal stability of the cellulase produced. The thermostability studies have shown that
enzyme preparations are best stored at very low temperatures for them to still retain their
activities. Even some soluble enzyme preparations stored at temperatures as low as +4oC lose
some of their activities on long storage. This leaves enzyme immobilization and lyophilisation as
better choices for costly enzymes that lose appreciable part of their activity even when stored at
low temperatures.
4.2 Conclusion
This study has shown that the fungus ( Aspergillus fumigatus), isolated from the native
environment has potential of being a source of cellulases using native carbon sources. Two
distinct parts of corn cob (CC-Outer and CC-Inner) were successfully used in the production of
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the enzyme. Corn cob, being an agricultural waste littered in the environment can be utilized to
produce cheap but viable cellulases.
Among the two distinct parts of corn cob used, the pulp (CC-Inner), though very light,
small in quantity and difficult to separate from the other part, has proven to be a better substrate
for fungal cellulase production.
4.3 Recommendations
Based on the findings from this work, the following recommendations are made:
i. Aspergillus fumigatus is a good source of cellulases using corn cob pulp as carbon source,
but the hard outer part may require some form of delignification as pre-treatment. The
organism may however do better on it perhaps by being genetically improved.
ii. An investigation into the use of corn cob pulp as a component of commercial media
formular for cellulotytic fungal isolation will be both revealing and rewarding.
However, a cheap technology will be necessary for efficient separation of the pulp
from the outer part of corn cob even with ease.
iii. Corn cob could be used by chemical industries for both sugar production and cellulosic
ethanol production instead of foodstuff.
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APPENDICES
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Appendix I: Glucose standard curve
Appendix II: Protein standard curve
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Appendix III The pH of soil samples
Soil Samples pH Temperature
DSD 6.05 30.5
DL-MSH 7.61 29.3
GS 7.02 29.5
SS-UNN 7.33 28.2
SW-UNN 7.37 26.8
Appendix iv: Preparation of reagents
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Solution A: Alkaline sodium carbonate (Na2CO3) solution prepared by dissolving 2g of
Na2CO3 in 100ml of 0.1M NaOH (0.4g of sodium hydroxide pellets were dissolved in 100ml of
distilled water).
Solution B: Prepared by dissolving 0.5g of CuSO4 and 1g of sodium potassium tartarate in 100ml
of distilled water.
Solution C: Dilute Folin-Ciocalteau Reagent prepared by diluting the commercial stock reagent
with equal volume of water, ( i.e, in the ratio of 1:1).
Solution D: Prepared by mixing 50ml of freshly prepared solution A with 1ml of solution B.
Solution E: Standard Solution of pure protein (Bovine Serum Albumin, BSA) 2mg/ml was
prepared by dissolving 0.1g of BSA in 50ml of distilled water. All these solutions were
prepared fresh at each time of use.
Preparation of acetate buffer
Sodium acetate buffer (0.2M pH 5.5) was prepared by dissolving 16.40g sodium acetate
(solid) in 0.5 litre of distilled water and stirred with magnetic stirrer till a homogenous solution
was obtained. The solution was titrated against acetic acid till a pH of 5.5 was obtained.
Preparation of DNS reagent
DNS (2.5g) was dissolved in 50 ml of 2M NaOH solution. 75g of sodium potassium
tartarate was dissolved in 125 ml of distilled water and mixed with the initial 50 ml DNS – NaOH
solution and then made up to 250 ml with distilled water. The reagent was filtered into an amber
bottle , capped and kept inside a dark cupboard.
Preparation of 5mM glucose
Analar grade glucose (0.09g) was dissolved in 100 ml of distilled water and then shaken
thoroughly to get a homogenous solution.
