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TITLE PAGE
EVALUATING THE POTENTIAL OF WILD COCOYAM (Caladium bicolor) FOR BIO-ETHANOL PRODUCTION USING INDIGENOUS FUNGAL ISOLATES
BY
ONYENMA, NGOZI CHIZURUM
PG/M.Sc./12/61544
A DISSERTATION SUBMITTED TO THE SCHOOL OF POST GRADUATE STUDIES, UNIVERSITY OF NIGERIA, NSUKKA, IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE AWARD OF MASTER OF SCIENCE (M.Sc.) DEGREE IN INDUSTRIAL MICROBIOLOGY
SUPERVISOR: PROF. A. N. MONEKE
NOVEMBER, 2016 ii
CERTIFICATION
Onyenma, Ngozi Chizurum (Reg. No. PG/M.Sc./12/61544), a postgraduate student in the Department of Microbiology, majoring in Industrial Microbiology, has satisfactorily completed the requirements for course work and research for the degree of Masters in Science (M.Sc.) in Microbiology. This work embodied in her project is original and has not been submitted in part or full for either diploma or degree of this university or any other university.
………………………………… …………………………………… Prof. L.I. Ezeogu Prof. A.N. Moneke Head, Supervisor, Department of Microbiology, Department of Microbiology, University of Nigeria, Nsukka. University of Nigeria, Nsukka.
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DEDICATION
This work is dedicated to the memory of my father Elder Johnson Onyenma, who passed on to me a love of reading and respect for education, and without whose caring support it would not have been possible.
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ACKNOWLEDGEMENTS
I have made efforts in this project work. However, it would not have been possible without the kind support and help of many whom I would like to extend my sincere thanks to.
I wish to thank most the Almighty God for giving me the determination to complete this project and to improve myself in a situation that I never imagined that I can surpass.
My profound gratitude is extended to my HOD and my research supervisor, Prof. A. N. Moneke, who gave me a golden opportunity to do this wonderful project work, which helped me to learn about so many new things. I am also particularly grateful for the assistance given by Dr. Mrs. O. C. Amadi and Dr. Mrs. T. N. Nwagu. Their advice, assistance in keeping my progress on schedule and their willingness to give their time so generously has been very much appreciated.
To my super mum and siblings, your prayers, unconditional support (both financially and emotionally) and your encouragement throughout my study saw me through.
Finally, my thanks and appreciations go to my colleagues and friends who helped me a lot in developing and finalizing this project work with their abilities.
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ABSTRACT
Diminishing fossil fuel reserves and environmental pollutions associated with the use of fossil fuels prompts the search for alternative energy sources that are renewable, sustainable, cost effective and safe. This study evaluates the potential use of Caladium bicolor (fleshy part of corm, peel of corm, leaf and stalk) for ethanol production. Eighteen fungal strains were isolated from healthy and rotten corms of C. bicolor and also from fresh palm wine. They were screened for amylase activity, cellulase activity and only the yeast strains screened for ethanol fermentation ability. Hydrolysis was carried out in flasks containing mineral salts medium (MSM), at pH 5.0, 4% substrate concentration and 1.0 x 108 spores/ml inoculum size at 28 ± 2°C for 7 days and amount of reducing sugar produced determined. Effects of time, pH, substrate concentration, nitrogen source, and inoculum size on hydrolysis of the various substrates were studied. Optimal conditions were combined and the time course of the enzymatic hydrolysis obtained. Following hydrolysis, Saccharomyces sp. (4.0 x 107 cells/ml) isolated from fresh palm wine was used to ferment the sugar for 7 days. The fermented liquid was distilled and maximum ethanol yield determined. The results obtained from the screening showed that nine isolates (three organisms: Aspergillus spp., Penicillium spp. and Rhizopus sp.) showed enzymatic activity. Six showed multi-enzyme activity while three isolates showed just amylolytic activity. Aspergillus sp. (Org 2) showed the highest amylolytic and cellulolytic ability and thus used for further study. Maximum reducing sugar yield was achieved on day 5 when the Aspergillus sp. was cultured on the media containing fleshy part of corm (23.297g/L) and stalk (15.320g/L), while the maximum reducing sugar yield for peel of corm and leaf were 18.013g/L and 6.667g/L, respectively on day 4. Optimal reducing sugar yield was achieved with pH 5 for all the substrates. Optimal ethanol yield from fleshy part of corm, peel of corm and leaf were 0.485%, 0.210%, and 0.380%, respectively on day 5 while that of stalk and mixed substrate were 0.280% and 0.280% on day 3 and 6, respectively. This study therefore demonstrates the potential of utilizing fleshy part of corm, peel of corm, leaf and stalk of C. bicolor for glucose production which can be fermented for bio-ethanol production especially in areas where they are in abundance.
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TABLE OF CONTENTS
Title page i
Certification ii
Dedication iii
Acknowledgements iv
Abstract v
Table of Contents vi
List of Tables xii
List of Figures xiii
List of Appendices xvi
CHAPTER ONE: INTRODUCTION AND LITERATURE REVIEW
1.1 Introduction 1
1.1.1 Statement of Problem 2
1.1.2 Aim of Study 2
1.1.3 Objectives of the Study 3
1.2 Literature Review 3
1.2.1 Overview of Ethanol 3
1.2.2 Caladium bicolor (Wild Cocoyam) 5
1.2.3 Raw Materials for Ethanol Production 7 vii
1.2.3.1 Ethanol from Sugar 7
1.2.3.2 Ethanol from Starches 7
1.2.3.3 Ethanol from Lignocelluloses 8
1.2.4 Pretreatment of Lignocellulose Biomass 12
1.2.4.1 Physical Pretreatments 14
1.2.4.2 Chemical Pretreatments 17
1.2.4.3 Biological Pretreatments 18
1.2.4.4 Combined Pretreatments 19
1.2.5 Enzymatic Hydrolysis 19
1.2.5.1 Amylolytic Enzymes and their Producer Organisms 20
1.2.5.2 Cellulolytic Enzymes and their Producer Organisms 23
1.2.6 Yeast and Its Fermentative Ability 26
1.2.7 Hydrolysis and Fermentation Strategies 28
1.2.7.1 Separate Hydrolysis and Fermentation (SHF) 28
1.2.7.2 Simultaneous Saccharification and Fermentation (SSF) 28
CHAPTER TWO: MATERIALS AND METHODS
2.1.1 Collection of Sample (Substrate) 30
2.2 Identification of Sample 30
2.3 Proximate Analysis of Sample 30
2.4 Processing of Sample 30 viii
2.5 Isolation of Microorganisms 31
2.5.1 Samples 31
2.5.2 Media for Isolation 31
2.5.3 Isolation Procedure 31
2.6 Purification / Preservation of Isolates 32
2.7 Screening of the Isolated Microorganisms 32
2.7.1 Qualitative Screening of Isolates for Amylase Production 32
2.7.2 Qualitative Screening of Isolates for Cellulase Production 32
2.7.3 Quantitative Screening of Isolates for Ability to Generate Reducing Sugars from
C. bicolor 33
2.7.4 Screening Yeast Isolates for Alcohol Production 33
2.8 Characterization/Identification of Selected Isolates 34
2.8.1 Characterization/Identification of Mould 34
2.8.1.1 Macroscopic Identification of Mould 34
2.8.1.2 Microscopic Identification of Mould 34
2.8.2 Characterization/Identification of Yeast Strains 35
2.8.2.1 Colonial Morphology of Yeast Isolates 35
2.8.2.2 Microscopy 35
2.8.2.3 Assimilation of Nitrogen Compound 36
2.8.2.4 Sugar Fermentation Test 36 ix
2.9 Preparation/Standardization of Inoculum 37
2.9.1 Preparation of Mould Inoculum 37
2.9.2 Preparation of Yeast Inoculum 37
2.9.3 Inoculum Standardization 37
2.10 Preparation of Standard Graphs 38
2.10.1 Preparation/Standardization of Glucose Concentration 38
2.10.2 Preparation/Standardization of Starch Concentration 39
2.10.3 Preparation/Standardization of Cellulose Concentration 39
2.11 Enzymatic Hydrolysis of Caladium bicolor 40
2.12 Analytical Methods in Enzymatic Hydrolysis of Caladium bicolor 40
2.12.1 Determination of Reducing Sugar Concentration 40
2.12.2 Determination of the Starch Content of Caladium bicolor (the fleshy part of corm) 41
2.12.3 Determination of the Cellulose Content of Caladium bicolor 41
2.13 Determination of Optimal Conditions on Hydrolysis of Caladium bicolor using the
Selected Test Organism Aspergillus sp. (Org 2) 42
2.13.1 Effect of Incubation Time 42
2.13.2 Effect of Varying Initial pH of the Medium 42
2.13.3 Effect of Varying Substrate Concentration 42
2.13.4 Effect of Different Nitrogen Sources 43 x
2.13.5 Effect of Varying Inoculum Size 43
2.13.6 Time Course of Enzymatic Hydrolysis of Caladium bicolor 43
2.14 Fermentation 43
2.15 Measurement of Ethanol Concentration 44
2.16 Statistical Analysis 45
CHAPTER THREE: RESULTS
3.1 Proximate Analysis of Caladium bicolor (Wild Cocoyam) 46
3.2 Screening of the Isolated Microorganisms 48
3.2.1 Qualitative Screening of Isolates for Enzyme Production 48
3.2.2 Quantitative Screening of Isolates for Ability to Generate Reducing Sugar from
C. bicolor 50
3.2.3 Screening Yeast Isolates for Alcohol Production 56
3.3 Characterization/Identification of Selected Isolates 58
3.3.1 Characterization/Identification of Mould Isolates 58
3.3.2 Characterization/Identification of Yeast Strains 58
3.4 Determination of Optimal Conditions on Hydrolysis of Various Parts of Caladium
bicolor using the Selected Test Fungi Aspergillus (Org 2) 62
3.4.1 Effect of Incubation Time 62
3.4.2 Effect of Varying Initial pH of the Medium 68
3.4.3 Effect of Varying Substrate/Carbon Concentration 74 xi
3.4.4 Effect of Different Nitrogen Sources 80
3.4.5 Effect of Varying Inoculum Size (1.0 x 108 spores/ml) 86
3.4.6 Time Course of Enzymatic Hydrolysis of Various Parts of Caladium bicolor 92
3.5 Fermentation 98
CHAPTER FOUR: DISCUSSION AND CONCLUSION
4.1 Screening of Isolates for Enzyme Production 105
4.2 Optimal Production Studies (Hydrolysis of Starch and Cellulose) 106
4.3 Fermentation 110
4.4 Conclusion 110
REFERENCES 112
APPENDICES 136
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LIST OF TABLES
Table Title Page
1 Pretreatment Methods of Lignocellulosic for Enzymatic Hydrolysis 13
2 Major Microorganisms Employed in Cellulase Production 25
3 Proximate Analysis of Caladium bicolor (Wild Cocoyam) 47
4 Qualitative Screening of Isolates for Enzyme Production 49
5 Screening Yeast Strains for Ethanol Production 57
6 Characterization/Identification of Mould Isolates 59
7 Cell Morphology and Physiological Characterizations of Yeast Strains 61
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LIST OF FIGURES
Figure Title Page
1 Photographs Showing the Image of the Different Parts of “Caladium bicolor”
Plant Studied 6
2 Chemical Structure of Amylose 8
3 Chemical Structure of Amylopectin 8
4 Chemical structure of cellulose 10
5 Screening of the Fungal Isolates for Hydrolytic Ability using Fleshy Part of Corm
as Substrate 51
6 Screening of the Fungal Isolates for Hydrolytic Ability using Peel of Corm as Substrate 52
7 Screening of the Fungal Isolates for Hydrolytic Ability using Leaf as Substrate 53
8 Screening of the Fungal Isolates for Hydrolytic Ability using Stalk as Substrate 54
9 Comparison of the Hydrolytic Ability of the Various Organisms 55
10 Effect of Incubation Time on Hydrolysis of the Fleshy Part of Corm 63
11 Effect of Incubation Time on Hydrolysis of the Peel of Corm 64
12 Effect of Incubation Time on Hydrolysis of the Leaf 65
13 Effect of Incubation Time on Hydrolysis of the Stalk 66
14 Comparison of the Reducing Sugar Produced from the Various Substrates with
Effect to Incubation Time 67
15 Effect of Varying Initial pH on Hydrolysis of the Fleshy Part of Corm 69
16 Effect of Varying Initial pH on Hydrolysis of the Peel of Corm 70 xiv
LIST OF FIGURES CONTINUED
Figure Title Page
17 Effect of Varying Initial pH on Hydrolysis of the Leaf 71
18 Effect of Varying Initial pH on Hydrolysis of the Stalk 72
19 Comparison of the Effect of Varying Initial pH on Hydrolysis of the Different
Plant Parts 73
20 Effect of Varying Substrate/Carbon Concentration on Hydrolysis of the Fleshy
Part of Corm 75
21 Effect of Varying Substrate/Carbon Concentration on Hydrolysis of the Peel of Corm 76
22 Effect of Varying Substrate/Carbon Concentration on Hydrolysis of the Leaf 77
23 Effect of Varying Substrate/Carbon Concentration on Hydrolysis of the Stalk 78
24 Comparison of the Effect of Varying Substrate/Carbon Concentration on
Hydrolysis of the Different Plant Parts 79
25 Effect of Different Nitrogen Sources on Hydrolysis of the Fleshy Part of Corm 81
26 Effect of Different Nitrogen Sources on Hydrolysis of the Peel of Corm 82
27 Effect of Different Nitrogen Sources on Hydrolysis of the Leaf 83
28 Effect of Different Nitrogen Sources on Hydrolysis of the Stalk 84
29 Comparison of the Effect of Different Nitrogen Sources on Hydrolysis of the
Different Plant Parts 85
30 Effect of Varying Inoculum Size on Hydrolysis of the Fleshy Part of Corm 87
31 Effect of Varying Inoculum Size on Hydrolysis of the Peel of Corm 88 xv
LIST OF FIGURES CONTINUED
Figure Title Page
32 Effect of Varying Inoculum Size on Hydrolysis of the Leaf 89
33 Effect of Varying Inoculum Size on Hydrolysis of the Stalk 90
34 Comparison of the Effect of Varying Inoculum Size on Hydrolysis of the Different
Plant Parts 91
35 Time Course of Hydrolysis of the Fleshy Part of Corm 93
36 Time Course of Hydrolysis of the Peel of Corm 94
37 Time Course of Hydrolysis of the Leaf 95
38 Time Course of Hydrolysis of the Stalk 96
39 Comparison of the Sugar Yield from Different Plant Parts during Time Course
of Hydrolysis 97
40 Ethanol Production from Hydrolysate of Fleshy Part of Corm and Reducing Sugar Utilization 99
41 Ethanol Production from Hydrolysate of Peel of Corm and Reducing Sugar Utilization 100
42 Ethanol Production from Hydrolysate of Leaf and Reducing Sugar Utilization 101
43 Ethanol Production from Hydrolysate of Stalk and Reducing Sugar Utilization 102
44 Ethanol Production from Hydrolysate of Mixed Substrate (Various Plant Parts)
and Reducing Sugar Utilization 103
45 Percentage Ethanol Yield of the Different Substrates 104 xvi
APPENDICES
Appendix Title Page
1 Glucose Standard curve 136
2 Starch Standard Curve 137
3 Cellulose Standard curve 138
4 Reference Standard Table for Ethanol Determination 139
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CHAPTER ONE
INTRODUCTION AND LITERATURE REVIEW
1.1 Introduction
Energy in all its forms is essential to humanity and is central to the improvement in people’s quality of life. The continuous increase in energy demand, the inevitable decline in the availability of fossil fuels, and the growing concerns about climate change have sparked a number of initiatives from governments around the world to increase production of energy from renewable sources (Quintero et al., 2008). Bio-fuels, and in particular bio-ethanol, i.e. ethanol obtained from crops or lignocellulosic biomass, are getting a lot of attention as possible option for renewable transportation fuel.
Liquid fuels are used as energy sources throughout the world and there have been progressive increases in their utilization. In the year 2004, average liquid fuel consumption was 83 million barrels per day which is projected to be 97 million barrels per day in the year 2015 and 118 million barrels per day in 2030 (EIA, 2007). Brazil and the United States lead the industrial world in global ethanol production, accounting together for 70% of the world's production and nearly 90% of ethanol is used for fuel. In 2006, Brazil produced 16.3 billion liters of ethanol representing 33.3% of the world's total ethanol production. Sugar cane plantations cover 3.6 million hectares of land for ethanol production with a productivity of 7,500 liters of ethanol per hectare. The U.S. on the other hand produced 3,000 liters per hectare of maize ethanol (Cardona and Sanchez, 2007).
Ever increasing demand for liquid fuel will deplete currently available fuel resources and there is increased interest in exploring alternative sources for the production of this energy resource. The high prices of fossil fuel have led to energy crises in both developing and developed countries that are oil dependent. According to Naylor et al. (2007) “bio-fuels will remain a critical energy development target in many parts of the world if petroleum prices exceed US $ 55 – 60 per barrel.” At the global level, Brazil has been the world leading producer of bio-fuel and recently, Nigeria joined the league of bio-fuel users with the aim of generating wealth (Aisien et al., 2010). 2
Ethanol is one of the preferable liquid fuels due to its combustion properties and its use as an additive with gasoline (Galbe and Zacchi, 2002). Of importance is that a mix of ethanol and gasoline reduces green house gas emissions at certain levels, minimizes dependence on fossil fuel and reduces waste management problems. The design had generally been production from sugarcane and cassava. Klass (1998) categorized cassava alongside with sweet potato and yam as main starches that serve as staple foods for people through the world’s hot and humid regions. These plants are so proficient at supplying essential calories that they are considered the quintessential subsistent crops. However, the success of these starch crops as staple foods limits their potential development and general economic growth, for instance, cassava which has become an important bio-fuel crop is a crop crucial for food security especially in Nigeria. The implication of this is that threats to food security exist in the face of growing fuel ethanol demand. Perhaps, the diversion of food resource to bio-fuel production may to a large extent have fuelled the current food crises worldwide (Srinorakutara et al., 2008). It therefore becomes imperative that the searchlight be turned at present to the use of non-food starchy items for the production of fuel ethanol.
1.1.1 Statement of Problem
Gasoline usage has a higher demand every year and the world faces a crisis of diminishing fossil fuel reserves, thus an alternative energy source that is renewable, sustainable, efficient, cost effective, convenient and safe is sought after (Chum and Overend, 2001).
Most of the raw materials utilized for bio-ethanol production are corn grain, sugar cane (Mojovic et al., 2006), and cassava (Klass, 1998). These plants are so proficient at supplying essential calories and the implication of this is that threats to food security exist in the face of growing fuel ethanol demand. However, ethanol production from non-edible crops or lignocelluloses is considered more attractive as they will not compete with food sources that are proposed for ethanol production.
1.1.2 Aim of Study
To evaluate the potential of Wild Cocoyam “Caladium bicolor” for bio-ethanol production.
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1.1.3 Objectives of the Study
The following are the specific objectives of the study:
To isolate moulds and yeasts from rotten corms of C. bicolor, healthy corms of C. bicolor, and Palm wine; To screen isolates for both amylase and cellulase production; To screen the yeast isolates for ethanol production; To determine the optimum culture condition for the hydrolysis of the starch and cellulose present in C. bicolor; To determine the suitability of C. bicolor as source of sugar for bio-ethanol production.
1.2 Literature Review
1.2.1 Overview of Ethanol
Ethanol is known as ethyl alcohol or fermentation alcohol, and is often referred to as just
‘‘alcohol’’, and has the chemical formula of C2H5OH. It is a colorless, clear liquid that looks like water and is completely miscible with water. Ethanol has a somewhat sweet flavor when diluted with water; a more pungent, burning taste when concentrated; and an agreeable ether-like odor. However, it is more volatile than water, flammable, burns with a light blue flame, and has excellent fuel properties for spark ignition internal combustion engines (Wyman, 2004). In daily application, ethanol is mostly used as fuels (92%), industrial solvents and chemicals (4%) and beverages (4%) (Logsdon, 2006).
The idea to use ethanol as a source of energy is not new. The oldest evidence about alcohol used as an engine fuel comes from 1899. Between the world wars about 4 million cars used gasoline blended with 25% volume of ethanol (Chereminisoff, 1979). Bio-ethanol is a renewable energy source produced mainly by the sugar fermentation process; although it can also be synthesized by chemical processes such as reacting ethylene with steam (Anuj et al., 2007). The alcohol fermentation by biochemical process is brought about by the action of yeast through a process that transforms the natural sugar present in any starchy material into alcohol with the evolution 4
of carbon dioxide (CO2) under controlled environmental conditions (Dubey, 2005; Okorondu et al., 2009).
Sugar substances such as sugarcane juice, sugar beets and molasses can be fermented directly to ethanol while starchy and cellulosic materials must be hydrolyzed to monomer sugar solutions; before it can be fermented to ethanol by microorganisms (yeasts or bacteria); and purified by e.g distillation and dehydration (Palmqvist and Hahn-Hägerdal, 2000; Millati et al., 2002; Taherzadeh and Karimi, 2007; Sameera et al., 2011). During the fermentation process, part of the sugar is assimilated by the yeast cells and part is transformed into glycerol, acetaldehydes and lactic acid.
An important issue regarding the ethanol production is weather the process is economical. Research efforts are focused to design and improve a process, which would produce a sustainable transportation fuel. A low cost feedstock is a very important factor in establishing a cost effective technology (Mojovic et al., 2006). Therefore, a strong need exists for efficient ethanol production with low cost raw material and production process. Using biomass to produce energy can reduce the use of fossil fuels, reduce pollution and waste management problems and show environmental advantages in terms of life-cycle energy use and greenhouse gas (GHG) emissions (Marshall, 2007; Inderlwildi and King, 2009; Rettenmaeir et al., 2010; Fernando et al., 2010). Also, it had been pointed out that the use of firewood, kerosene and charcoal in households had adverse effects on human health (Adelekan and Adelekan, 2004). Ethanol represents closed carbon dioxide cycle because after burning of ethanol, the released carbon dioxide is recycled back into plant material because plants use CO2 to synthesize cellulose during photosynthesis cycle (Wyman, 2004). Ethanol production process only uses energy from renewable energy sources. Hence, no net carbon dioxide is added to the atmosphere, making ethanol an environmentally beneficial energy source.
Bio-ethanol production involves several steps starting from selection of proper feedstock, its pre- treatment, amylase and cellulase production, hydrolysis of feedstock using amylases and cellulases, fermentation of hydrolysate, and finally distillation of the fermented hydrolysate to obtain pure ethanol.
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1.2.2 Caladium bicolor (Wild Cocoyam)
The Araceae comprises a large family of herbaceous monocots that occurs in all continents, except the Antarctic, and has two main centers of distribution – tropical America and tropical Asia (Croat, 2000). Within the family Araceae, Caladium belongs to the tribe Caladieae, a neotropical group in the subfamily Aroideae. Caladiums are ornamental aroids grown and bred for their colourful and various-shaped foliage. It is used as a foliage plant, indoors and out, and as cut leaves due to its longevity in arrangements (Miller, 1997). They can be propagated from seed but are highly heterozygous and show great variability in the seedling population (Hartman et al., 1972). The majority of Caladium cultivars are therefore propagated asexually from tuber pieces. New cultivars are developed by hybridization, primarily based on foliage coloration but other important characteristics include plant growth habit, petiole strength, tuber production, and disease and pest resistance (Wilfret, 1993). The genus Caladium comprises 12 species (Mayo et al., 1997), of which C. bicolor is the major source of cultivars. C. picturatum and C. marmoratum are now considered synonyms of C. bicolor (Madison, 1981). These cultivars fall into two general categories: fancy leaf and strap leaf caladiums. Fancy leaf caladiums have broad heart-shaped leaves borne on erect petioles while strap leaf caladiums have narrow, lanceolate leaves on short petioles and produce a more compact plant than the fancy-leaved type. Caladium plants have not been well studied at the molecular level and so the naming and identification of species within the genus Caladium (Araceae) has been difficult and based primarily on morphology (Loh et al., 1999). However, the lack of up-to-date comprehensive reference material which illustrates all cultivars in colour makes identification of Caladium cultivars extremely difficult (Loh et al., 1999). Wilfret (1993) reported that there are over 2000 cultivar names for Caladium and over a hundred cultivars are grown today.
Caladium bicolor ‘Florida Clown’ is a wild cocoyam commonly known as “Ede Umuagbara”, “Ede Umunmo” or “Ede Obara Jesus” in the eastern part of Nigeria. They look like our normal edible cocoyam (Colocasia esculenta) but can be differentiated by the red and white blotches found on the leaves. They are tuberous, heart-shaped leaves may vary in size from 15 cm to 60 cm in length and features leaves with randomly white and red blotches. C. bicolor is a non- human edible plant that grows along river banks, lakes, streams, brooks, oases and shady areas in Nigeria. Also here in Nigeria, they grow indiscriminately, without being cultivated. Unlike the 6 edible cocoyam, it has a prominent and non-hardy cocoyam corm. It is self sustaining as it has broader leaves, which enable it to suppress other weeds under and around it (Umoren, 2005). All parts should not be ingested and may irritate sensitive skin. Caladium contains oxalate crystals which can cause illness and swelling of the mouth and throat (Iwu, 1993).
The use of this biomass (C. bicolor) for bio-ethanol production will help solve the problem of the diminishing fossil fuel reserve and at the same time not lead to food insecurity that can come about while using staple food like cassava, yam, sweet potato, and cocoyam.
(a) C. bicolor Plant (b) C. bicolor Corm
(c) C. bicolor Leaf (d) C. bicolor Stalk
Figure 1: Photographs Showing the Image of the Different Parts of “Caladium bicolor” Plant Studied.
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1.2.3 Raw Materials for Ethanol Production
The most used raw materials for bio-ethanol production can be classified into three main types: sugars, starches and cellulose materials (Liu and Shen, 2008).
1.2.3.1 Ethanol from Sugars
Sugar crops deemed suitable as raw material for bio-ethanol production include sugarcane, sugar beet, and sorghum to mention a few (Ogbonna et al., 2001). One advantage of the sugar crop is that it does not require any step of hydrolysis; it needs only a milling process for the extraction of sugars to fermentation (Icoz et al., 2009). Sugar cane in form of either cane juice or molasses (by-product of sugar mills) is the main feedstock for ethanol production in Brazil. About 79% of ethanol in Brazil is produced from fresh sugar cane juice and the remaining percentage from cane molasses (Wilkie et al., 2000). Some distilleries use only juice, while others use only molasses, but the mixture is considered to be a better substrate, since the juice has some nutritional deficiencies, whereas molasses has inhibitory compounds for yeast fermentation (Basso et al., 2011).
1.2.3.2 Ethanol from Starches
Starch is another renewable agricultural substrate available and can be easily hydrolysed to fermentable sugars using amylolytic enzymes. These enzymes could be economically produced by microorganisms (Nigam and Singh, 1995).
Starch, chemical formula (C6H10O5)n is a polysaccharide carbohydrate consisting of a large number of glucose units joined together by glycosidic bonds. It consists of amylose (linear chain of glucose) and amylopectin (branched chain of glucose). Starches are found in a large number of plants as the major carbohydrates reserve and provide an essential source of energy. The most common sources of food starch are corn, potatoes, wheat, cassava/tapioca and rice (Akpa and Dagde, 2012).
Amylose consists of anhydroglucose units that are linked by α-D-1, 4 glucoside bonds to form linear chains (Figure 2). Amylose molecules are typically made up from 200-2000 anhydroglucose units. The level of amylose and its molecular weight vary between different 8 starch types. Aqueous solutions of amylose are unstable due to intermolecular attraction and association of neighboring amylose molecules. This leads to viscosity increase, retrogradation and precipitation of amylose particles (Hedley, 2002).
Figure 2: Chemical Structure of Amylose (Reis et al., 2002).
Amylopectin has a polymeric, branched structure (Figure 3). It consists of anhydroglucose units that are linked by α -D-1,6 bonds that occur every 20-30 anhydroglucose units. In addition, it also has α-D-1,4 bonds that are present in amylose. The level of amylopectin varies between different starches types. Aqueous solutions of amylopectin are characterized by high viscosity, clarity, stability and resistance to gelling (Hedley, 2002).
Figure 3: Chemical Structure of Amylopectin (Reis et al., 2002).
1.2.3.3 Ethanol from Lignocelluloses
Lignocellulosic biomass is the most abundant material on Earth. Its sources, including raw materials like, agricultural residues (e.g. corn stover and wheat straw), forestry residues (e.g. sawdust and mill wastes), portions of municipal solid waste (e.g. waste paper) and various industrial wastes have a great potential to be used in the industrial processes. A considerable 9 amount of such materials as waste byproducts are being generated through agricultural practices mainly from various agro based industries (Pérez et al., 2002). Sadly, much of the lignocellulosic biomass is often disposed of by burning, which is not restricted to developing countries alone. Recently, lignocellulosic biomasses have gained increasing research interests and special importance because of their renewable nature (Asgher et al., 2013; Ofori-Boateng and Lee, 2013). Therefore, the huge amounts of lignocellulosic biomass can potentially be converted into different high value products including bio-fuels, value added fine chemicals, and cheap energy sources for microbial fermentation and enzyme production (Isroi et al., 2011; Asgher et al., 2013, Iqbal et al., 2013, Irshad et al., 2013).
Lignocellulosic substrates consist of mainly three types of polymers: cellulose, hemicelluloses and lignin (Hendriks and Zeeman, 2009) along with smaller amounts of ash, pectins, proteins and soluble sugars (Jorgensen et al., 2003). In plants, linear cellulose chains contribute to tensile strength, while hydrophobic amorphous lignin is responsible for chemical resistance, in particular protection against water. Hemicellulose provides bonding between cellulose and lignin. Thus, two major obstacles hinder the hydrolysis of cellulose in lignocellulosic material. They are, the recalcitrance of crystalline cellulose itself (emerging from the linear cellulose chain structure tightly bound in microfibrils), and the highly protective lignin surrounding it, and acting as a physical barrier for microorganisms (i.e. enzymatic attack).
A. Cellulose
Cellulose with the chemical formula (C6H10O5)n is a major component of lignocellulose. It has always been referred to as the most abundant renewable polymer on earth (Ding and Himmel, 2009) and constitutes a major fraction of plant cell wall. Chemically, it is a simple molecule composed of linear β-1,4-linked D glucopyranose chains (also called glucose or glucan chains). While β-1,4-linked glucose is the chemical repeating unit, the structural repeat is β-cellobiose (Figure 4) (Varrot et al., 2003). In cellulose, glucose chains are tightly bound to each other by van der Waals forces and hydrogen bonds into crystalline structures called elementary fibril (consisting of around 40 glucan chains), about 40 Å wide, 30 Å tick and 100 Å long (Bidlack et al., 1992). Aggregates of these elementary fibrils are called microfibrils order i.e., crystalline and amorphous regions (Taherzadeh and Karimi, 2008; Iqbal et al., 2011). Regions within the 10 microfibrils with high order are termed crystalline, and less ordered regions are termed amorphous.
Figure 4: Chemical structure of cellulose. Linear β-1,4-linked glucose is the chemical repeating unit, while the structural repeat is β-cellobiose, and consequently each glucoside is oriented at 180° in respect to its neighbors. From Varrot et al., 2003.
B. Hemicellulose
Hemicelluloses are the second most abundant renewable organic material, next to cellulose, on Earth. Contrary to cellulose, hemicelluloses are heterogeneous, amorphous and branched polymers, consisting of diverse monosaccharides i.e. mixture of pentose sugars (xylose and arabinose) and hexose sugars (glucose, mannose and galactose). Hemicellulose also consists of sugar acids such as glucuronic acid, methylglucuronic acid and galacturonic acid (Taherzadeh et al., 2013). The degree of polymerisation in hemicellulose is about 200 sugar units, which is far less than that of cellulose. They are easily hydrolysed to their monomeric sugars by acids. Hemicellulose’s backbone polymer could either be heteropolymer or homopolymer linked with β-1,4- glycosidic bonds and occassionally with β-1,3- glycosidic bonds (Kuhad et al., 1997). Softwood hemicellulose is dominated by glucomannans while the hemicelluloses of hardwood and agricultural residues are dominated by xylan. However, degree of acetylation in hardwood hemicelluloses is higher than in softwood hemicelluloses (Saha, 2003; Sixta 2006).
Due to their branched structure, hemicelluloses are more soluble than cellulose and they can be isolated from wood by extraction. Hemicelluloses are easily hydrolyzed by strong acid leaving 11 cellulose and lignin intact (Liu and Wyman 2005; Lloyd and Wyman, 2005), or by strong base
(Fan et al., 1982). In many cases diluted acid (0.5-1.0 % H2SO4) pretreatment under elevated temperatures (140-190 °C) will degrade most of the hemicellulose to soluble pentose and hexose sugars (Lloyd and Wyman, 2005). Even though this treatment is not particularly targeted towards solubilization of lignin, the lignin structure is disturbed and redistributed leading to much more favorable (pretreated) substrate for enzymatic hydrolysis (Yang and Wyman, 2004).
C. Lignin
Lignin is probably the most complex and the least characterized molecular group among the wood components. Its primary purpose is to give strength and water permeability to plants, and also to protect plants from pathogen infections. It has a long-chain, heterogeneous polymer composed largely of phenyl-propane units most commonly linked by ether bonds. Lignin acts like a glue by filling the gap between and around the cellulose and hemicellulose complexion with the polymers. It is present in all plant biomass; therefore, it is considered byproduct or as a residue in bio-ethanol production process. Lignin is comprised of complex and large polymer of phenyl-propane, methoxy groups and non-carbohydrate poly phenolic substance, which bind cell walls component together (Hamelinck et al., 2005). Phenyl-propanes (3 carbons attached with 6 carbon atom rings) are main block of lignin. These phenyl-propanes denoted as 0, I, II methoxyl groups attached to rings give special structure I, II and III. These groups depend on the plant source which they are obtained. Structure I exist in plants (grasses) and structure II found in the wood (conifers) while structure III present in deciduous wood. Generally, softwoods have higher lignin content than hardwood and agricultural residues (Kumar et al., 2009; Pérez et al., 2002). Lignin can be used as fuel for heat and electricity generation in the plant; other industrial applications for lignin exist such as adhesives and biocomposites production (Lora and Glasser, 2002; Macfariane et al., 2014; Ghaffar and Fan, 2014).
Other lignocelluloses components are extractives, which are non-cell wall components usually, found in foliage and bark. Common extractives are resins, terpenes and sterols, and they constitute approximately 1–5% of the wood. Non-extractive components such as ash, silica, proteins and pectins are also constituents of lignocellulosic materials (Klinke, 2004; Kostamo et al., 2004; Leiviskä et al., 2009). 12
1.2.4 Pretreatment of Lignocellulose Biomass
In order to produce ethanol from lignocellulosic materials, the bundles of lignocelluloses needs to be opened in order to access the polymer chains of cellulose and hemicelluloses by a process of pretreatment; hydrolyze the polymers in order to achieve monomer sugar solutions; ferment the sugars to ethanol solution (mash) by microorganisms; and purify ethanol from mash by e.g distillation and dehydration (Taherzadeh and Karimi, 2007). Without any pretreatment, the conversion of native cellulose to sugar is extremely slow, since cellulose is well protected by the matrix of lignin and hemicellulose in macrofibrils. Therefore, pretreatment of these materials is necessary to increase the rate of hydrolysis of cellulose to fermentable sugars (Galbe and Zacchi, 2002).
Pretreatment of lignocellulosic biomass has been an active field of research for several decades, and a wide variety of thermal, mechanical, chemical and biological pretreatment approaches (and combinations thereof) have been investigated and reported in the scientific literature (McMillan, 1994). It is important that the selected pretreatment technology includes number of requirements;
Should be able to improve the enzymatic accessibility of the lignocellulosic compound. Result in the minimum loss of the potential sugars. Prevent the formation of molecules which are inhibitory to microbial degradation or enzymatic action. Pretreatment technology should be economically sound in order to make the overall process i.e. conversion of biomass to simple sugar a feasible technology.
To make monomeric sugar utilization from lignocellulosic biomass a viable option, there are several methods introduced for pretreatment of lignocellulosic materials, which are summarized in Table 1. The methods have been classified into three major categories: physical, chemical, and biological methods. These methods can also be combined, for example, as physicochemical pretreatment for improvement of digestibility and consequently, improved ethanol yield. In some cases, a method is used to increase the efficiency of another method. For instance, milling could be applied to create a better steam explosion by reducing the chip size. Furthermore, it should be noticed that the selection of pretreatment method should be compatible with the selection of hydrolysis. For example, if acid hydrolysis is to be applied, a pretreatment with alkali may not be 13 beneficial (Taherzadeh and Niklasson, 2004). The pretreatment methods were reviewed by McMillan (1994), Wyman (1996), Sun and Cheng (2002), and Mosier et al. (2005b) (Table 1).
Table 1: Pretreatment Methods of Lignocellulosic for Enzymatic Hydrolysis.
Method Processes Mechanism of Changes on Biomass Physical -Ball-milling -Increase in accessible surface area and size Pretreatments -Two-roll milling of pores. -Hammer milling -Decrease of the cellulose crystallinity and -Colloid milling its degrees of polymerization. -Vibro energy milling -Partial hydrolysis of hemicelluloses. -Hydrothermal -Partial depolymerization of lignin. -High pressure steaming -Extrusion -Expansion -Pyrolysis -Gamma-ray irradiation -Electron- beam irradiation -Microwave irradiation Physicochemical Explosion: -Delignification. & Chemical -Steam explosion -Decrease of the cellulose crystallinity and Pretreatments -Ammonia fiber explosion its degrees of polymerization. (AFEX) -Partial or complete hydrolysis of -CO2 explosion hemicelluloses. -SO2 explosion Alkaline: -Sodium hydroxide -Ammonia -Ammonium sulfite Gas: -Chlorine dioxide -Nitrogen dioxide Acid: -Sulfuric acid -Hydrochloric acid -Phosphoric acid -Sulfur dioxide Oxidizing agents: -Hydrogen perioxide -Wet oxidation 14
-Ozone Cellulose solvents: -Cadoxen -CMCS Solvent extraction of lignin: -Ethanol-water extraction -Benzene-water extraction -Ethylene glycol extraction -Butanol-water extraction -Swelling agents
Biological -Actinomycetes -Delignification. Pretreatments -Fungi -Reduction in degree of polymerization of hemicelluloses and cellulose.
1.2.4.1 Physical Pretreatments
Physical pretreatments include mechanical (milling, grinding, chipping…), irradiation (gamma- ray, electron-beam, and microwave irradiation), high pressure steaming, extrusion, expansion, pyrolysis and liquid hot water (LHW) also known as hydrothermal pretreatment. Physical pretreatment functions primarily as a means of size reduction for the lignocellulosic material either physically or mechanically. It increases the surface area and porosity of the material; it decreases the cellulose crystallinity and depolymerises the cellulose fibre (Taherzadeh and Karimi, 2008); partially hydrolyzes hemicelluloses and partially depolymerizes lignin. Therefore the organic matter availability to enzymes or microorganisms is favoured (Hemery et al., 2009; Dumas et al., 2010; Ghizzi et al., 2010).
Mechanical and thermal methods exist to treat agro-industrial residues, but these methods tend to require a high energy input which can increase the processing cost considerably. Product separation for fermentation purposes can also make physical pretreatments expensive.
During steam explosion, lignocellulosic biomass is heated rapidly to a high temperature (160- 260°C) with sufficient pressure (1-7 MPa) to enable water molecules to penetrate the substrate structure for a few minutes. The pressure is then suddenly released to allow the water molecules to escape in an explosive manner. Steam pretreatment can be improved by using an acid catalyst, -1 such as H2SO4 or SO2 (0.3-3 gacid 100g ), which increases the recovery of cellulose and 15 hemicelluloses sugars (Ballesteros et al., 2000; Galbe and Zacchi, 2007). This pretreatment opens up the plant cells, increases surface area and enhances the digestibility of biomass
(Ballesteros et al., 2000). Piccolo et al. (2010) have shown that SO2 steam explosion at 190°C during 2 min increase the accessible surface area of wheat straw from 1.1 m2 g-1 (untreated wheat straw) to 1.9 m2 g-1. Limitations of steam explosion are the incomplete disruption of the lignin- carbohydrate matrix and the formation of hemicelluloses and cellulose degradation byproducts as water acts as an acid at high temperature (e.g. furfural and hydroxymethylfurfural) (Kumar et al., 2009).
Hydrothermal pretreatment or cooking of lignocellulosic materials in liquid hot water (LHW) is one of the old methods applied for pretreatment of cellulosic materials. Autohydrolysis plays an important role in this process, where no chemical is added. It results in dissolution of hemicelluloses mostly as liquid-soluble oligosaccharides and separates them from insoluble cellulosic fractions. The pH, processing temperature, and time should be controlled in LHW pretreatment in order to optimize the enzymatic digestibility of lignocellulosic materials (Mosier et al., 2005a; Mosier et al., 2005c; Wyman, 1996). LHW pretreatment of corn fiber at 160oC and a pH above 4.0 dissolved 50% of the fibre in 20 min (Mosier et al., 2005c). The results showed that the pretreatment enabled the subsequent complete enzymatic hydrolysis of the remaining polysaccharides to the corresponding monomers. The carbohydrates dissolved by the LHW pretreatment were 80% soluble oligosaccharides and 20% monosaccharides with less than 1% of the carbohydrate lost to degradation products. LHW causes ultrastructural changes and formation of micron-sized pores that enlarged accessible and susceptible surface area and make the cellulose more accessible to hydrolytic enzymes (Zeng et al., 2007).Three types of reactor can be used for liquid hot water pretreatment: co-current (biomass and water are heated together for a certain residence time), counter-current (water and lignocelluloses move in opposite directions), and flow-through (hot water passes over a stationary bed of lignocelluloses) (Liu and Wyman, 2005; Mosier et al., 2005a). In general, liquid hot water pretreatments are attractive for their cost-savings potential: no catalyst requirement and low-cost reactor construction due to low- corrosion potential. However, water and energetic requirement remain higher (Alvira et al., 2010). 16
Hydrothermal processing of agro-industrial residues causes a variety of effects including extractive removal, hemicellulose hydrolysis and alteration of the properties of both cellulose and lignin. Water treatments provide an interesting alternative for the chemical utilization of lignocellulose owing to the following reasons: i. No chemicals different from water are necessary, the whole process being environment- friendly. ii. Hemicelluloses can be converted into hemicellulosic sugars at good yields with low byproduct generation (Lamptey et al., 1985), leading to solutions of sugar oligomers and/or sugars that can be utilized for a variety of practical purposes (Modler, 1994; Saska and Ozer, 1995; Aoyama et al., 1995; Aoyama, 1996; LoÂpez-Alegret, 1996). iii. In comparison with acid pre-hydrolysis, no problems derived from equipment corrosion are expected owing to the mild pH of the reaction media. iv. Stages of sludge handling and acid recycling are avoided, resulting in a simplified process structure. v. The physico-chemical alteration provoked by treatments on lignin and cellulose facilitates the further separation and processing of these fractions. vi. Economic estimates (Schaffeld, 1994; Kubikova et al., 1996) showed advantages for water treatments over alternative technologies.
Without any pretreatment, corn stover with sizes of 53-75 µm was 1.5 times more susceptible to enzymatic hydrolysis than the larger stover particles of 425-710 µm. However, this difference was eliminated when the stover was pretreated with liquid hot water (LHW) also known as hydrothermal pretreatment at 190 oC for 15min, at a pH between 4.3 and 6.2 (Zeng et al., 2007). Laser et al. (2002) compared the performance of LHW and steam pretreatments of sugarcane bagasse in production of ethanol by Separate Hydrolysis and Fermentation. They used a 25-l reactor, temperature 170-230 oC, residence time 1-46 min and 1% to 8% solids concentration. Both of the methods generated reactive fibers, but LHW resulted in much better xylan recovery than steam pretreatment. It was concluded that LHW pretreatment produces results comparable with dilute-acid pretreatment processes.
17
1.2.4.2 Chemical Pretreatments
Chemical pretreatment employs the use of acids such as phosphoric acid, sulphuric acid and hydrochloric acid; gases, namely sulphur dioxide; and alkali e.g. ammonia for fractionation of lignocellulosic biomass and enhancement of cellulose digestibility (Zhang et al., 2007). Often, chemical pretreatment is usually combined with physical processes commonly referred to as physicochemical pretreatment such as steam explosion with the addition of SO2 or H2SO4, ammonia fibre explosion (AFEX), as well as CO2 explosion (Eklund et al., 1995; Kim and Hong, 2001; Alizadeh et al., 2005). Physicochemical processes have been reported to be effective and promising for large scale industrial processes due to its fast rate of lignocellulose degradation and complete hemicellulose fractionation (Taherzadeh and Karimi, 2008). The most commonly used chemical pretreatments include: acid and alkali based hydrolysis approaches. The acidic reaction can be divided into dilute or concentrated acid hydrolysis.
Dilute-acid hydrolysis is probably the most commonly applied method among the chemical hydrolysis methods. It is a method that can be used either as a pretreatment preceding enzymatic hydrolysis, or as the actual method of hydrolyzing lignocelluloses to the sugars (Taherzadeh and Karimi, 2007). The dilute-acid pretreatment can achieve high reaction rates and significantly improve cellulose hydrolysis. Different aspects of dilute-acid hydrolysis have recently been reviewed (Taherzadeh and Karimi, 2007). One of the main advantages of dilute-acid hydrolysis is achieving high xylan to xylose conversion yields, which is necessary to achieve favourable overall process economics in ethanol production from lignocelluloses (Sun and Cheng, 2002). On the other hand, a main disadvantage of this pretreatment method is the necessity of neutralization of pH for the downstream enzymatic hydrolysis. Furthermore, different chemical inhibitors might be produced during the acid pretreatment with reduced cellulase activity, and therefore, water wash is necessary for the pretreated biomass before enzymatic hydrolysis (Mes- Hartree and Saddler, 1983; Sun and Cheng, 2002). Badshah et al. (2012) have recently shown the application of dilute-acid pretreatment on sugarcane bagasse at 121°C during 15 min and -1 0.02 H2SO4 g L led to an increase of 166% compared to untreated bagasse. Dilute acid pretreatment using HCl was also found efficient to enhance the methane potentials of 21% and 48%, respectively for sunflower stalks and sunflower oil cakes (Monlau et al., 2012a; Monlau et 18 al., 2012b). On the contrary, dilute-acid pretreatment on maize silage was found inefficient, which can be explained by the nature of the substrate.
AFEX, or ammonia fibre explosion, is one of the physiochemical pretreatment methods in which lignocellulosic materials are exposed to liquid ammonia at high temperature (e.g. 90-100 oC) for a period of time (such as 30 min), and then the pressure is swiftly reduced. There are many adjustable parameters in the AFEX process: ammonia loading, water loading, temperature, time, blowdown pressure, and number of treatments (Holtzapple et al., 1991). AFEX, with a concept similar to steam explosion, can significantly improve the enzymatic hydrolysis. The optimal conditions for pretreatment of switchgrass with AFEX were reported to be about 100oC, ammonia loading of 1:1 kg of ammonia per kg of dry matter, and 5 min retention time (Alizadeh et al., 2005). Enzymatic hydrolysis of AFEX-treated and untreated samples showed 93% vs. 16% glutan conversion, respectively. An advantage of AFEX pretreatment is no formation of some types of inhibitory by-products, which are produced during other pretreatment methods, such as furans in dilute-acid pretreatment. However, cleaved lignin phenolic fragments and other cell wall extractives may remain on the biomass surface, which can easily be removed by washing with water (Chundawat et al., 2007). Although AFEX enhances hydrolysis of (hemi) cellulose from glass, the effect on biomass that contains more lignin (soft and hardwood) is meager. Furthermore, the AFEX pretreatment does not significantly solubilize hemicelluloses, compared to dilute-acid pretreatment. On the other hand, to reduce the cost and protect the environment, ammonia must be recycled after the pretreatment (Wyman, 1996; Sun and Cheng, 2002; Eggeman and Elander, 2005).
1.2.4.3 Biological Pretreatments
Biological method of pretreatment is a process whereby microorganisms are used for altering the structure of lignocelluloses, causing delignification by breaking down lignin into smaller structures through their extracellular enzymes. Biological pretreatment employs wood degrading microorganisms, including white-rot fungi, brown-rot fungi or soft-rot fungi, and bacteria e.g. proteobacteria and actinobacteria (Kurakeke et al., 2007; Adney et al., 2009; Ray et al., 2010) to modify the chemical composition and/or structure of the lignocellulosic biomass. Bio- delignification is useful for pretreatment purposes because it replaces or supplements the 19 chemical-based pretreatments, which include mechanical treatment with acid, alkali, and steam explosion (Iqbal et al., 2013). In spite of this, biological pretreatments are more effective, economical, eco-friendly and less health hazardous as compared to the physicochemical or chemical-based pretreatment approaches (Asgher et al., 2012); however, slow rate of action and long pretreatment times are its challenges (Sun and Cheng, 2002). In addition to this, most of the lignolytic microorganisms solubilize/consume not only lignin but also hemicellulose and cellulose. Because of these drawbacks/limitations the biological pretreatment faces techno- economic barriers and therefore is less attractive commercially (Eggeman & Elander, 2005). White-rot fungi have been widely investigated for bio-fuel production due to the substrate specificity of its extracellular lignolytic enzymes, namely, Lignin peroxidase (LiP), Manganese peroxidase (MnP) and Laccase (Lac). These enzymes can effectively degrade lignin and decrease the crystallinity of cellulose (Isroi et al., 2011). Biological pretreatment can also be combined with physical or chemical pretreatments for improved digestibility and ethanol yields (Isroi et al., 2012; Ma et al., 2010).
Ghosh and Bhattacharrya, (1999) studied the effect of white-rot fungi and brown-rot fungi on rice straw. Increases in methane of 32% and 46% were observed, respectively for rice straw pretreated with brown- and white-rot fungi compared to untreated straw.
1.2.4.4 Combined Pretreatments
These treatments are a combination of physical, chemical, biological and/or enzymatic treatments used in order to get the best of all treatments hence minimizing the disadvantages of individual treatments.
1.2.5 Enzymatic Hydrolysis
In the early 1970’s, it was considered that plant and animal materials were the best sources of enzymes, but nowadays, microbial enzymes are becoming increasingly important owing to their technical and economical advantages (Cherry et al., 2004). A number of microorganisms including fungal and bacterial species are associated with the production of amylase and cellulase. Extracellular amylase (, and glucoamylase) and cellulase (exoglucanase, endoglucanase and –glucosidase) have been reported to be produced by various species of 20 bacteria such as Bacillus subtilis, B. licheniformis, Escherichia species, Clostridium species and fungi such as Trichoderma, Aspergillus, Rhizopus, Penicillium and Candida species (Syu and Chen, 1997; Pothiraj et al., 2006).
Amylase are starch (a heterogenous polysaccharide composed of a polymer of glucose residues) degrading enzymes while cellulases are cellulose (a polysaccharide consisting of a linear chain of several hundred to many thousands of Beta 1,4 linked D-glucose) degrading enzyme. These enzymes act by hydrolyzing the –1,4 and –1,4 glycosidic bonds present in starch and cellulose respectively to yield a lower molecular weight sugar (Prasanna, 2005). Substrate concentration, accessible surface area, enzyme dosage, pH and temperature are important factors that should be optimised for efficient enzymatic hydrolysis process (Aslanzadeh et al., 2014).
1.2.5.1 Amylolytic Enzymes and their Producer Organisms
Starch represents one of the most abundant storage polysaccharides in nature and the most popular ingredient in food. Because of the complex structure of starch, cells require an appropriate combination of enzymes for its depolymerization to oligosaccharides and smaller sugars, such as glucose and maltose. Amylolytic enzymes play an important role in the degradation of starch and are produced in bulk from microorganisms representing about 25–33% of the world enzyme market. Microbial enzymes are preferred for their stability over plant and animal enzymes which increase their spectrum of industrial applications. They also have the advantages of cost effectiveness, consistency, less time and space required for production and ease of process modification and optimization. The amylolytic enzymes find a wide spectrum of applications in food industry for production of glucose syrups, crystalline glucose, high fructose corn syrups, maltose syrups, reduction of viscosity of sugar syrups, reduction of haze formation in juices, solubilization and saccharification of starch for alcohol fermentation in brewing industries, retardation of staling in baking industry, in detergent industry used as an additive to remove starch based dirts, in paper industry for the reduction of viscosity of starch for appropriate coating of paper, in textile industry for warp sizing of textile fibers and in pharmaceutical industry they are used as a digestive aid (Sivaramakrishnan et al., 2006).
The degree of hydrolysis of native starch depends on the factors such as the substrate concentration, type and concentration of enzyme used, and on the applied process conditions 21 such as pH, temperature and the mixing rate (Mojovic et al., 2006). There are two processes in enzymatic hydrolysis; liquefaction and saccharification. The breakdown of large particles drastically reduces the viscosity of gelatinized starch solution, resulting in a process called liquefaction. The final stages of depolymerization are mainly the formation of mono-, di- , and tri-saccharides. This process is called saccharification (Barsby et al., 2003).
Amylolytic enzymes of microbial origin are divided into exo-acting, endo-acting, debranching and cyclodextrin producing enzymes.
A. Exo Acting Amylases – Glucoamylases and β-Amylases
Glucoamylases (1,4-α-D-glucan glucohydrolase, EC 3.2.1.3) catalyse hydrolysis of α -1,4 and α - 1,6 glucosidic linkages to release β-D-glucose from the non-reducing ends of starch and related poly- and oligosaccharides. They have widely been reported to occur in a large number of microbes, including bacteria, yeast and fungi.
Filamentous fungi, however, constitute the major source among all microorganisms and strains of genera Aspergillus and Rhizopus are mainly used for commercial production (Pandey, 1995). β -amylases are known to be produced only by plants and certain bacteria mostly by several species of the genus Bacillus, including B. polymyxa, B. cereus, B. megaterium, and also by Clostridium thermosulfurogenes (Selvakumar et al., 1998). They hydrolyze α -1,4 bonds but cannot bypass α -1,6 linkages in amylopectin and glycogen and they produce maltose from amylose and maltose and a β -limit dextrin from amylopectin and glycogen.
B. Endo Acting Amylases
α -amylases (E.C. 3.2.1.1.) hydrolyse α -1,4 bonds and bypass α-1,6 linkages in amylopectin and glycogen. In spite of the wide distribution of amylases in microbes, animals and plants, microbial sources, namely fungal and bacterial amylases are preferred in industries. Among bacteria, Bacillus sp. is widely used for thermostable α -amylase production while fungi belonging to the genus Aspergillus are most common (Sivaramakrishnan et al., 2006). They are classified in two categories depending on the extent to which they hydrolyze starch. Liquefying α -amylases hydrolyze 30 to 40% of starch and saccharifying α -amylases hydrolyze 50 to 60%. 22
C. Debranching Amylases
Isoamylases and pullulanases are debranching enzymes that hydrolyze only α -1,6 linkages. On the basis of substrate specificity and product pattern, pullulanase (pullulan α -glucano-hydrolase; EC 3.2.1.41) have been classified into two groups: type I and type II. As they hydrolyze the α - glucosidase-resistant α -1,6 linkages in dextrins, they improve the starch saccharification rate and yield when used in combination with α -glucosidases. Many mesophilic (Aerobacter aerogenes, B. macerans, B. acidopullulyticus and Bacillus sp), thermophilic and hyperthermophilic bacteria and archae (B.stearothermophilus, Clostridium thermosulfurogenes, Pyrococcus and Thermococcus genus) have been reported to produce pullulanase (Gomes et al., 2003; Kunamneni and Singh, 2006).
D. Cyclodextrinases
Cyclodextrin glycosyltransferase (α -1,4-D-glucan, α -4-D-( α − 1,4-D-glucano)- transferase, EC 2.4.1.19) produces a series of non-reducing cyclic dextrins (α -, β- and γ-cyclodextrins) from starch, amylose, and other polysaccharides. α -, β- and γ-cyclodextrins contain six, seven and eight glucose units, respectively, that are linked by α -1,4-bonds. Thermococcus sp., B. coagulans, C. thermohydrosulfuricum 39E, B. sphaericus and alkalophilic Bacillus sp. are the most reported microorganisms producing these enzymes.
Amylase Producing Microorganisms
Amylases are produced by a wide spectrum of organisms, although each source produces biochemical phenotypes that significantly differ in parameters like pH and temperature optima as well as metal ion requirements (Rameshkumar and Sivasudha, 2011). Microorganisms are chosen preferentially for amylase production due to the relative ease of handling, availability, favorable growth conditions, and cheap nutrient requirement compared to other producers like plant and animal. Evidences of amylase in yeast, bacteria and moulds have been reported and their properties documented (Adebiyi and Akinyanju, 1998; Akpan et al., 1999; Buzzini and Martini, 2002). Among the microorganisms, many fungi had been found to be good sources of amylolytic enzymes (Omemu et al., 2004). Amylase of fungal origin was found to be more stable than the bacterial enzymes on a commercial scale. The Major advantage of using fungi for the 23 production of amylases is economical bulk production capacity and ease of manipulation (Aiyer, 2005). Many attempts have been made to optimize culture conditions and suitable strains of fungi (Abu et al., 2005). Studies on fungal amylase especially in the developing countries have concentrated mainly on Rhizopus sp. and Aspergillus niger probably because of the ubiquitous nature and non fastidious nutritional requirements of these organisms (Abe et al., 1988; Abu et al., 2005; Pandey et al., 2005; Gupta et al., 2008; Irfan et al., 2012). Aiyer, (2005) also reported that although many microorganisms produce this enzyme, the most commonly used for their industrial application are Bacillus licheniformis, Bacillus amyloliquifaciens and Aspergillus niger.
Other microorganisms producing appreciable amount of different amount of amylase enzyme are Escherichia sp., Micrococcus sp., Pseudomonas spp., Proteus sp., Serratia sp., Candida, Cephalosporium, Mucor, Penicillium Neurospora e.t.c.
1.2.5.2 Cellulolytic Enzymes and their Producer Organisms
Enzymatic hydrolysis is an effective and economical method to achieve fermentable sugars under mild and eco-friendly reaction conditions from the pretreated cellulosic biomass (Wyman et al., 2005). It is generally preferred over the acid hydrolysis due to its advantages such as: milder processing conditions (pH at 4.5–5.0, temperature at 40–50°C), possibility of achieving complete cellulose conversion to glucose, no formation of inhibitory by-products and no corrosive conditions associated with it (Ogier et al., 1999; Taherzadeh and Karimi, 2007). However, it does require 1–4 days of hydrolysis during enzymatic processing compared to only a few minutes for acid hydrolysis process.
Enzymatic hydrolysis of cellulose and hemicelluloses can be carried out by highly specific cellulase and hemicellulase enzymes (glycosylhydrolases). This group includes at least 15 protein families and some subfamilies (Rabinovich et al., 2002). The enzymes are extremely specific catalyst such as cellulase or cellulytic enzymes, which are produced by some microorganisms (Wyman, 1996). Cellulose hydrolysis to glucose is carried out by interaction or synergistic effect of three major groups of enzymes, which are: endo-1,4- β-d-glucanases, exo- 1,4-β-d-glucanases and β-d-glucosidases (Wood, 1985; Wyman,1996). The endo-glucanases cleaves the cellulose at the amorphous and less crystalline region and creates free chain ends; the 24 exo-glucanases cleaves the cellobiose units from the created free chain ends while β- glucosidase finally cleaves the cellobiose to glucose (Taherzadeh and Karimi, 2007).
Cellulase yields appear to depend upon a complex relationship involving a variety of factors like inoculums size, pH value, temperature, presence of inducers, medium additives, aeration, growth time, and so forth (Immanuel et al., 2006).
Cellulase Producing Microorganisms
Cellulolytic microbes are primarily cellulose degraders but generally do not utilize lipids or protein as energy source (Lynd et al., 2002). Many of them can utilize other carbohydrates in addition to cellulose but few anaerobic cellulolytic species have restricted carbohydrate range, limited to cellulose and their hydrolyzed product. Although a large number of microorganisms are capable of degrading cellulose, only a few of these microorganisms produce significant quantities of cell-free enzymes capable of completely hydrolysing crystalline cellulose in vitro. Fungi are the main cellulase-producing microorganisms, though a few bacteria and actinomycetes have also been reported to yield cellulase activity. Major cellulose producers have been listed (Table 2).
Microorganisms of the genera Trichoderma and Aspergillus are thought to be cellulase producers, and crude enzymes produced by these microorganisms are commercially available for agricultural use. Among the cellulases produced by different microorganisms, cellulases of Trichoderma reesei or T. viride have been the most broadly studied and best characterized. Trichoderma reesei has been reported to be capable of hydrolyzing native cellulose (Reczey et al., 1996; Singhania et al., 2006; Singhania et al., 2007) but inspite of being a prolific natural producer of extracellular cellulases, it may not be the most effective cellulase system for use in biomass conversion processes that essentially require complete hydrolysis of the feedstock for economic viability. A full complement production of cellulose, stability under the enzymatic hydrolysis conditions and resistance of the enzyme to chemical inhibitors are the advantages of the cellulose produced by Trichoderma. Microorganisms of the genus Trichoderma produce relatively large quantities of endo-ß-glucanase and exo-ß-glucanase but the main disadvantages of Trichoderma cellulose are the suboptimal levels and low activity of B-glucosidases. On the other hand, Aspergilli are very efficient endo-ß-glucanase and ß-glucosidase producers, with low 25 levels of exo-ß- glucanase production. In several studies, Trichoderma cellulase was supplemented with extra B-glucosidases and showed good improvement (Hari Krishna et al., 2001; Ortega et al., 2001; Tengorg et al., 2001; Itoh et al., 2003). The major microorganisms employed in cellulose production were reviewed by Rajeev et. al. (2005) (Table 2).
Table 2: Major Microorganisms Employed in Cellulase Production.
Major groups Genus Species Fungi Aspergillus A. niger A. nidulans A. oryzae A. aculeatus
Fusarium F. solani F. fusosporium
Humicola H. insolens H. griesa
Melanocarpus M. albomyces
Neorospora N. crassa
Phanerochaete P. chrysosporium
Penicillium P. brasilianum P. occitanis P. decumbans P. purpurogenum P. janthinellum
Talaromyces T. emersonii
Trichoderma T. reesei T. harzianum T. longibrachiatum
Bacteria Acidothermus A. cellulolyticus
26
Bacillus Bacillus sp. B. subtilis
Clostridium C. thermocellum C. acitobutylicum C. cellulovorans Pseudomonas P. cellulose
Rhodothermus R. marinus
Cellulomonas C. fim C. uda
Actinomycetes Streptomyces S. drodzowiczii S. lividans
Thermomonospora T. fusca T. curvata
1.2.6 Yeast and Its Fermentative Ability
Yeast is a fungus in which the unicellular form is the most predominant and which replicates by budding (or fission) (Walker, 2009). Yeasts are considerably larger in size than bacteria. In terms of quantity and in economics, is the most important group of microorganism exploited for industrial purposes (Russell, 2003). Yeast cells vary from oval to round and in size from roughly 5 to 10 µm length to a breadth of 5 to 7µm, but there are exceptions ranging from 2.5 to 21µm. Chemically the yeast cell consists of 75% water and 25% dry matter. The dry matter consist of carbohydrate (18-44%), protein (36-60%), nucleic acid (4-8%), lipids (4-7%), phosphorus (1- 3%), potassium (1-3%), sulfur (0.4%) and vitamins (trace amounts). Yeast can be classified as aerobic, facultative or respiro-fermentative or fermentative depending on their status viz a viz oxygen. An important fact to remember about yeast is the suppression of respiration by high glucose levels where the cells continue to ferment despite the availability of oxygen (called the Crabtree effect) (Walker, 2009). Yeast has been used for thousands of years to produce alcohol. The use of yeast to produce ethanol is conceptually straightforward. Using a large tank, the yeast 27 cells are simply fed a dilute solution of sugars and nutrients, allowed to grow and to ferment, and then the ethanol which they excrete are separated and purified.
There are many environmental conditions that affect yeast cell growth and the kinetics of chemical reactions within living cells. These include the availability of major and minor nutrients, the temperature, pH, and dissolved oxygen concentration, and the possible presence of competing organisms.
Yeast cells finely tune their growth and behavior in accordance with available nutrients. They can adjust their growth rate in response to their nutritional environment by altering the length of their cell cycle over at least a 10-fold range (Brauer et al., 2008). They can adapt to nutritional depletion by engaging one of a number of alternative developmental programs depending on the particular nutritional circumstances. These programs can range from rapid mitotic growth in rich media, to filamentous growth allowing foraging under limiting nutrient conditions, to various distinct quiescent states that reversibly shut down the cell in response to starvation for a single nutrient, to the extreme state of biological stasis following sporulation upon severe starvation.
Yeast cells grow on a wide variety of compounds as sources of energy and as carbon-containing precursors of anabolic metabolism and biomass accumulation (Johnston and Carlson, 1992). However, yeast cells consume glucose or fructose in preference to other mono-, di-, and trisaccharides, such as sucrose, raffinose, or trehalose, and prefer any fermentable carbon source over any source, such as glycerol, ethanol, or acetate, that has to be catabolized by oxidative phosphorylation.
The Saccharomyces family of yeast can best ferment in temperatures from 25-26°C. Ideally, the yeast reacts best in a slightly acidic environment (pH of 4.5) (Lin, 2006). S. cerevisiae has the capabilities of withstanding high concentrations of ethanol as well as producing high ethanol yields from glucose.
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1.2.7 Hydrolysis and Fermentation Strategies
1.2.7.1 Separate Hydrolysis and Fermentation (SHF)
In this process, starches and pretreated lignocelluloses are hydrolysed to glucose and subsequently fermented to ethanol in separate units. The major advantage of this method is that it is possible to carry out the starch and/or cellulose hydrolysis and fermentation at each of their optimum conditions. Among the fungi, most amylase production studies have been done within temperature range of 25-37 oC (Francis et al., 2003; Ramachandran et al., 2004); Bacterial alpha amylases are produced at a much wider range of temperature. Bacillus amyloliquefaciens, B. subtilis, B. licheniformis and B. stearothermophilus are among the most commonly used Bacillus sp. reported to produce alpha amylase at temperatures 37-60 oC (Mielenz, 1983; Syu and Chen, 1997; Mendu et al., 2005; Mishra et al., 2005); while the optimum temperature for cellulose is usually between 45 and 50 oC, depending on the cellulose-producing microorganism (Wingren et al., 2003; SÖderstrÖm et al., 2003; Saha et al., 2005; Olsson et al., 2006). However, the optimum temperature for most of the ethanol-producing microorganisms is between 30 and 37 oC.
Inhibition of amylase and cellulase activity by the released sugars is the main drawback of SHF. Another possible problem in SHF is that of contaminations and the process is rather long (Taherzadeh and Karimi, 2007). However, since fermentation and hydrolysis usually have different optimum temperatures, separate enzymatic hydrolysis and fermentation (SHF) is still considered as a choice.
1.2.7.2 Simultaneous Saccharification and Fermentation (SSF)
One of the most successful methods for ethanol production from starch and/or lignocellulosic materials is combination of the enzymatic hydrolysis (of starch and/or pretreated lignocellulose) and fermentation in one step, termed SSF. In this process, the glucose produced by the hydrolyzed enzymes is consumed immediately by the fermenting microorganism present in the culture. This is a great advantage for SSF compared to SHF, since the inhibition effects of cellobiose and glucose to the enzymes are minimized by keeping a low concentration of these sugars in the media. SSF gives higher reported ethanol yields than SHF and requires lower 29 amounts of enzyme (Eklund and Zacchi, 1995; McMillan et al., 1999; Sun and Cheng, 2002; Karimi et al., 2006). The risk of contamination in SSF is lower than in the SHF process, since the presence of ethanol reduces the possibility of contamination. Furthermore, the number of vessels required for SSF is reduced in comparison to SHF, resulting in lower capital cost of the process.
An important strategy in SSF is to have the optimum conditions for the enzymatic hydrolysis and fermentation as close as possible, particularly with respect to pH and temperature. However, the difference between optimum temperatures of the cellulotic enzymes and fermenting microorganisms is still a drawback of SSF. The optimum temperatures for cellulases is usually between 45 and 50 oC, whereas Saccharomyces cerevisiae has an optimum temperature between 30 and 35 oC and practically inactive at more than 40 oC.
Inhibition of amylase and/or cellulase by produced ethanol might be also a problem in SSF. It was reported that 30 g/l ethanol reduces the enzyme activity by 25% (Wyman, 1996). Ethanol inhibition may be a limiting factor in producing high ethanol concentration.
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CHAPTER TWO
MATERIALS AND METHODS
2.1 Collection of Sample (Substrate)
The wild cocoyam plant used in this study was harvested from a farm in Oboloafor in Nsukka, Enugu State and transported to the Microbiology Laboratory, University of Nigeria Nsukka for analysis.
2.2 Identification of Sample
Sample was taken to Botany Department, University of Nigeria, Nsukka for proper identification. The wild cocoyam plant (sample) was identified to be Caladium bicolor “Florida Clown”.
2.3 Proximate Analysis of Sample
Part of the fresh and healthy corms, leaves and stalks of the C. bicolor were taken to Crop Science Laboratory, University of Nigeria Nsukka, for proximate analysis of the corm (both the fleshy part and the peel), leaf and stalk.
2.4 Processing of Sample
The fresh and healthy unpeeled corms, leaves and stalks were sorted and washed with water to remove soil contamination. They were then spread out on clean surface and allowed enough time to air-dry under shade at room temperature. Using a pre-sterilized knife, the corms were peeled and sliced into smaller pieces. The leaves and stalks were also sliced. They were dried to a constant weight and ground separately to fine powders using manual grinder to obtain stock samples. The ground samples were screened through a fine mesh sieve to remove any large particle and to obtain the same size range.
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2.5 Isolation of Microorganisms
2.5.1 Samples
The organisms used for this study were isolated from three sources: (1) Ten grams sample of sliced rotten corms of C. bicolor was taken and put into 250 mL sterile conical flask containing 100 mL sterile distilled water. This was plugged with a sterile cotton wool, properly shaken and served as stock; (2) Ten grams sample of sliced healthy corms of C. bicolor were steeped for 3 days in a 250 mL sterile conical flask containing 100 mL sterile distilled water, plugged with a sterile cotton wool and this served as stock for mould and yeast isolation; (3) Palm wine samples were purchased from wine tappers at Obukpa and Ogige market both in Nsukka, Enugu State. A 100ml fresh and raw undiluted palm wine sample also served as stock for yeast isolation.
2.5.2 Media for Isolation
Potato Dextrose Agar (PDA) and Sabouraud Dextrose Agar (SDA) were used as growth media in this research. They were prepared according to manufacturers specifications thus, 39 g of powdered PDA medium and 65 g of powdered SDA medium were separately dissolved in 1 L distilled water, both supplemented with 0.05 g/L chloramphenicol to inhibit bacterial growth, homogenized and sterilized separately by autoclaving at 121oC for 15 min. They were allowed to cool before carefully dispensing approximately 20 mL of each medium into the Petri dishes. The Petri dishes containing the different media (PDA and SDA) were allowed to solidify and were incubated over night at room temperature (28 ± 2°C)) to check for sterility before using them for isolation.
2.5.3 Isolation Procedure
A ten-fold serial dilution was performed for each of the stock and aliquots of 0.1 mL of 10-4 to 10-7 dilutions were aseptically plated in duplicate onto the respective agar plates with the aid of automatic micro-pipette with sterile tips using spread plate method. The inoculum was spread on the agar medium with sterile glass spreader (the glass spreader was sterilized by dipping into ethanol, flaming, and then cooling). The culture plates were now incubated at room temperature (28 ± 2°C) for 2-7 days and then examined daily for development of fungal growth. 32
2.6 Purification / Preservation of Isolates
Colonies differing in size, shape and colour were selected from different agar plates and sub- cultured on PDA and SDA by the spread plate and streak plate technique. A distinct colony was picked from the culture plates with a sterile wire loop to obtain pure cultures. The agar plates were incubated at room temperature (28 ± 2°C) for 2-7 days. After successive sub-culturing, the subsequent pure cultures were maintained on agar slants and were preserved for use in refrigerator at 4oC. In subsequent experiment, the working cultures were propagated once in PDA agar plate, and then sub-cultured also on PDA agar plates to check for purity of the cells before use.
2.7 Screening of the Isolated Microorganisms
2.7.2 Qualitative Screening of Isolates for Amylase Production
The preliminary screening for amylase production was carried out by starch agar plate assay on standard media (Abe et al., 1988; Akpan et al., 1999; Fogarty, 1983). The constituents of media were 1.5 g yeast extract, 2.0 g soluble starch, 0.5 g peptone, 1.5 g NaCl and 15.0 g agar dissolved in 1.0 L distilled water and supplemented with 0.05 g/L chloramphenicol. The plates were point inoculated with spores of different fungal isolates. After 72 h of incubation, the inoculated plates were stained with Gram's iodine reagent, allowed for 15 min and washed with spring warm water to remove the excess color. Development of purple/blue-black colour indicates the absence of starch degradation while presence of clear zone around the colonies indicates secretion of amylase and hydrolysis of starch. Un-inoculated starch agar plate served as control.
2.7.2 Qualitative Screening of Isolates for Cellulase Production
Point inoculation of the spores of the different isolates was grown on PDA plates supplemented with carboxylmethylcellulase (CMC, 2% w/v) medium. After inoculation, the plates were incubated at room temperature (28 ± 2°C) for 72 h after which it was stained with 1% Congo red solution for 15 min. Excess dye was removed by washing with 1 M NaCl. The production of extracellular cellulase by the organism was indicated by a zone of clearance around the fungal colonies on the plate. Un-inoculated PDA agar plate supplemented with 2% CMC served as control (Bisaria and Ghose, 1987). 33
2.7.3 Quantitative Screening of Isolates for Ability to Generate Reducing Sugars from C. bicolor (Glucose was used as Standard)
The mineral salts medium (MSM) of Adeyemo and Sani, (2013) was used with slight modification for the cultivation of fungal isolates. It was prepared using the following compositions (g/L). KH2PO4, 5 g; (NH4)2SO4, 5 g; MgSO4.7H2O, 0.3 g; CaCl2, 0.5 g; FeSO4,
0.013 g; MnSO4.H2O, 0.04 g; ZnSO4.7H2O, 0.04 g; Yeast extract, 0.5 g; Carbon source, 40 g. The carbon sources used were the fleshy part of corm, peel of corm, leaf, and stalk, all from C. bicolor plant. Thirty milliliter of each medium was dispensed into 100 mL conical flask, plugged with cotton wool and sterilized in the autoclave at 121oC for 15 min. Spores of 3-4 days old cultures were harvested by washing PDA slants with 10 mL of sterile saline water. An aliquot of 1.2 mL (4%) of the spore suspensions was aseptically used to inoculate 30 mL of each of the liquid medium, containing any of the carbon sources. The culture media were incubated at room temperature (28 ± 2°C) under static motion with intermittent manual shaking (to enhance mass- transfer) for 8 days. The experiment was performed under aerobic condition. Two conical flasks were removed at 2 days intervals from each preparation. 10 mL of each cultured sample was withdrawn from culture into a centrifuge tube and the supernatant that resulted following centrifugation at 4000 rpm for 15 min to remove solids was used for analysis. The amylase/cellulase activity of each culture filtrate was measured by determining the amount of reducing sugar liberated using dinitrosalicylic acid method (DNSA) of Miller (1959). The absorbance of the tubes was measured at 540nm using 772S Spectrophotometer (B-Bran Scientific and Instrument Company England). This was done for all the eighteen (18) isolates.
2.7.4 Screening Yeast Isolates for Ethanol Production
Isolated yeasts were screened for ability to produce ethanol according to the method of Brooks (2008). A yeast suspension was prepared by aseptically transferring 2 wire loops full from each slant culture to 5 mL sterile Glucose peptone yeast (GPY) broth in Bijou bottle, comprising of 8 g of glucose, 1 g of peptone and 1 g of yeast extract in 100 mL. It was mixed by gentle manual agitation and incubated for 24 h at room temperature (28 ± 2°C). Then 1 mL of the pre-culture was transferred to sterile 10 mL GPY broth and incubated for 24 h at room temperature (28 ± 2°C). Eight yeast isolates were screened for fermentative ability by inoculating 2 mL of each 34 yeast suspension into 8 mL of GPY broth contained in test tubes carrying inverted Durham’s tubes and incubated for 30 h. Based on the volume of gas in the Durham’s tube, four ethanol productive strains were selected, two isolated from healthy corms of C. bicolor and the other two from palm wine.
2.8 Characterization/Identification of Selected Isolates
2.8.1 Characterization/Identification of Mould
The entire mould isolates were characterized and identified macroscopically and microscopically using conventional microbiological methods.
2.8.1.1 Macroscopic Identification of Mould
The macroscopic identification was based on the morphological, physiological and cultural characteristics as described by Ellis et al., (2007).
2.8.1.2 Microscopic Identification of Mould
Slide culture technique was used in this microscopic identification of moulds. The procedure is as follows:
Step 1: Preparing the Culture
Petri dishes containing ‘V’ shaped glass rod were sterilized. Also glass slides, cover slips and SDA medium were sterilized along with the Petri dishes. Sterile slide was aseptically placed on top of the ‘V’ shaped glass rod using a forceps, after which sterile scalpel blade was used to cut out the solidified SDA agar block about 1 cm square and the agar block placed on top of the slide using sterile scalpel blade. The fungus under investigation was inoculated at the four sides of the agar block and using an instant sterilized forceps, the sterile cover slip was picked and placed gently on the inoculated agar block. About 2 mL of sterile distilled water was dispensed aseptically into the plate to provide a humid environment suitable for fungal growth. The plates were incubated at room temperature (28 ± 2°C) for 5-7 days to allow sporulation to occur undisturbed. 35
Step 2: Mounting of Growth from Slide Culture Preparations
After about 5-7 days, the cover slip was gently removed with forceps and the side with growth was turned to face upwards. The agar block was gently removed with inoculating needle without disturbing the growth on the slide and the block discarded into a disinfectant jar to avoid spreading the spores on the block inside the laboratory. A new clean slide and a new cover slip were used for mounting the growth in the cover slip and slide, respectively using a drop of lacto- phenol blue. Water bubbles on the slide and cover slip within the areas of fungal growth were removed by dropping 70% ethanol to dry them up and excess mounting fluid were also removed. The growth on the slide as well as on the cover slip was observed under microscope to obtain two conventional slide mounts. The funguses were viewed with the microscope using x10 and x40 objective lens.
The microscopic identification was with the use of Photographic Atlas for the Microbiology Laboratory (Michael et al., 2011), and Descriptions of Medical Fungi (Ellis et al., 2007).
2.8.2 Characterization/Identification of Yeast Strains
The yeast strains selected were characterized and identified on the basis of macroscopic and microscopic morphological description, nitrogen assimilation test and sugar fermentation test as described by Yarrow (1998), Queshi et al. (2007) and Kurtzman et al. (2011).
2.8.2.1 Colonial Morphology of Yeast Isolates
Colonies of each yeast strain on GPY agar medium were observed and described on the basis of colour, texture and shape.
2.8.2.2 Microscopy
An aliquot of 48 h old culture was picked with wire loop and placed on a clean grease free glass slide. Then a drop of lacto-phenol blue was added and mixed using the wire loop. A cover slip was then placed on the slide and viewed under the microscope using x10 and x40 objective lens.
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2.8.2.3 Assimilation of Nitrogen Compound
This physiological test was carried out to check the ability of the yeast strain to grow on a medium containing sodium nitrate as the sole source of nitrogen. A basal agar medium was prepared by dissolving 20 g of glucose, 1 g of potassium dihydrogen phosphate, 0.5 g of magnesium sulphate (heptahydrate), 20 g of agar in 1 L sterile distilled water. The prepared sterile agar plates were seeded with yeast suspension by dispensing 0.1 mL of 10-4 dilutions of yeast suspensions and spread using a glass spreader, and then it was allowed to dry for about 20 min. The basal medium lacked nitrogen source but contained glucose as carbon source. A sodium nitrate source was introduced by placing a filter paper soaked in saturated solution of sodium nitrate on the surface of the seeded plate. The plates were incubated at room temperature (28 ± 2°C) and examined 2 days interval for 4 days for zones of growth around the sites of the nitrogen source.
2.8.2.4 Sugar Fermentation Test
The sugar fermentation test was carried out using the following sugars: glucose, galactose, maltose, fructose, sucrose, raffinose, xylose and lactose. The ability to ferment sugar was assayed in test tubes containing 8 mL of basal medium containing inverted Durham tube to enable detection of gas production. The basal medium was prepared by dissolving 2 g of sugar (except in the case of raffinose where 4 g was used), 0.4 5g of yeast extract, 0.75 g peptone and 4 mL of bromothymol blue (indicator) in 100 mL distilled water. The medium was sterilized at 121oC for 15 min. A 2 mL of the yeast suspension [prepared by aseptically transferring 2 wire loops full from each slant culture to 5 mL GPY broth in Bijou bottle and mixed by gentle manual agitation and incubated for 24 h at room temperature (28 ± 2°C), then 1 mL of pre-culture transferred to 10 mL GPY broth and incubated for 24 h at room temperature (28 ± 2°C)] was dispensed into 8 mL basal medium and incubated for 24 h at room temperature (28 ± 2°C) and examined 2 days interval for 4 days for acid and gas production.
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2.9 Preparation/Standardization of Inoculum
2.9.1 Preparation of Mould Inoculum
The mould isolates were propagated from the stock isolate to PDA agar plate to check for purity of the cells before use. The mould isolates were then subcultured from the pure PDA plate to sterile PDA agar slants. Spores of 4 days old cultures were harvested by washing PDA slants with 10 mL of sterile normal saline.
2.9.2 Preparation of Yeast Inoculum
From the stock culture, yeast strain selected as potential starter culture was subcultured in a PDA plate to check for purity before use. A 100 mL liquid medium was prepared comprising of 2 g glucose, 0.5 g yeast extract, 0.5 g (NH4)2SO4, 0.2 g KH2PO4, 0.1 g MgSO4.7H2O in 250 mL conical flask. The flask was autoclaved at 121oC for 15 min. The medium was allowed to cool and one loopful of the isolated yeast cell inoculated into the liquid medium. The flask was incubated for 24 h at room temperature (28 ± 2°C). Ten milliliter of the pre-culture was taken and inoculated into another 250 mL of the above medium. This was also incubated for 24 h at room temperature (28 ± 2°C) (Ogbonna et al., 2011). The yeast cells were harvested by dispensing the liquid culture medium in 10ml volume sterile centrifuge tubes and spinned at 4000rpm for 5 min using centrifuge (Andreas Hettich D72 centrifuge, Tuttlingen Germany). After centrifugation, the supernatant was discarded and the pellets washed by adding 10 mL of normal saline solution to each tube and spinned at 4000 rpm for 5 min and the supernatant discarded. The pellets were re-suspended in 10ml volume of saline solution contained in a Macarthney bottle (Moonja et al., 2003; Zheng et al., 2012). To form the working stock, the cells were diluted using normal saline solution to a final optical density (A540nm) reading of 0.5 absorbance.
2.9.3 Inoculum Standardization
To standardize the inoculums using haemocytometer (Neubauer Chamber), Superior Marienfeld, Made in Germany, four steps were involved. 38
Step 1: Sample Preparation: A ten-fold serial dilution was prepared from the inoculum suspensions of the mould spores.
Step 2: Introducing the Sample into the Neubauer Chamber: A haemocytometer was setup and 0.1 mL of the dilution previously prepared (Step 1) was withdrawn using a disposable micropipette tip. The spore suspension was dispensed/ introduced to the edge of the glass slide placed on the grid of the haemocytomer. The cells took up space and filled the grid by capillary action.
Step 3: Microscope Set up and Focus: The haemocytometer was placed on the stage of a light microscope and fixed with the clamp of the microscope. The light of the microscope was turned on and the microscope focused by adjusting the stage to observe a sharp image of the spores using objective lens of x40 magnification. The spores in four extreme big squares were counted (each extreme square is subdivided into 16 sub-squares).
Step 4: Concentration Calculation: The spore concentration was calculated using the formula:
Concentration (spores/mL)= Number of spores x 10,000
Number of squares x Dilution
The spore suspension was then used as inoculum after measuring the strength (1.0 x 108 spores/mL) using a hemocytometer.
The whole process (steps 1 - 4) was repeated for the yeast cell suspension to get a final cell concentration (4.0 x 107 cells/mL).
2.10 Preparation of Standard Graphs
2.10.1 Preparation/Standardization of Glucose Concentration
A glucose stock solution was prepared by weighing out 0.5 g of glucose in 100 mL of sterile distilled water. This was properly mixed and five successive transfers (in triplicate) was made in an arithmetic progression (0.2, 0.4, 0.6, 0.8, & 1.0) from the stock glucose solution into test tubes 39 and a final volume of 1 mL was made up in each test tube with sterile distilled water using pipette. One milliliter of dinitrosalicylic acid (DNSA) was added into each test tube and placed in boiling water for 10 min. The test tubes were allowed to cool, and 10 mL of sterile distilled water was added to all the tubes. The absorbance of the tubes were measured at 540nm using 772S Spectrophotometer (B-Bran Scientific and Instrument Company England) against a blank containing 1 mL of distilled water and 1 mL of DNSA to which 10 mL of distilled water was added after boiling. The standard graph was a plot of the absorbance readings of the dilution tubes against their respective glucose concentration (Miller, 1959) (Graph in appendix I).
2.10.2 Preparation/Standardization of Starch Concentration
Starch stock solution was prepared by weighing out 0.5 g of soluble starch, which was slowly poured and mixed into approximately 75 mL of sterile distilled water that had just been boiled. This solution was brought to 100 ml volume by further addition of water to form a stock of starch. Five successive transfers (in triplicate) was made in an arithmetic progression (0.2, 0.4, 0.6, 0.8, & 1.0) from the starch stock solution into test tubes and a final volume of 3 mL was made up in each test tube with distilled water using pipette. Five milliliter of iodine solution was added into each test tube and this was properly mixed. The final volume of all the tubes was made up to 15 mL by addition of 7 mL distilled water. The absorbance of the tubes were measured at 550nm using 772S Spectrophotometer (B-Bran Scientific and Instrument Company England) against a blank containing 5 mL of iodine solution and 10 mL of distilled water. The standard graph was a plot of the absorbance readings of the dilution tubes against their respective starch concentration (Astolfi-Filfo, 1986) (Graph in appendix II).
2.10.3 Preparation/Standardization of Cellulose Concentration
Cellulose stock solution was prepared by weighing 100 mg of micro-crystalline cellulose into a test tube; 10 mL of 67% sulphuric acid was added and allowed to stand for 1 h. One milliliter of the above solution was diluted to 100 mL with distilled water. Five successive transfers (in triplicate) was made in an arithmetic progression (0.15, 0.35, 0.55, 0.75, & 0.95) from the cellulose stock solution into test tubes and a final volume of 1 mL was made up in each test tube using pipette. To all the test tubes, 10 mL of anthrone reagent was added and mixed well. A blank was set up with 1 mL distilled water and 10 mL anthrone reagent. The tubes were placed 40 in a boiling water-bath for 10 min. The tubes were allowed to cool and the colour measured at 630nm using 772S Spectrophotometer (B-Bran Scientific and Instrument Company England). The standard graph was a plot of the absorbance readings of the dilution tubes against their respective cellulose concentrations (Updegroff, 1969) (Graph in appendix III).
2.11 Enzymatic Hydrolysis of Caladium bicolor
The mineral salts media (MSM) for the enzymatic hydrolysis of C. bicolor was prepared with the following compositions (g/L): KH2PO4, 5 g; (NH4)2SO4, 5 g; MgSO4.7H2O, 0.3 g; CaCl2, 0.5 g;
FeSO4, 0.013 g; MnSO4.H2O, 0.04 g; ZnSO4.7H2O, 0.04 g; Yeast extract, 0.5 g; Carbon source, 40 g (slightly modified method of Adeyemo and Sani, 2013). The carbon sources used were the fleshy part of corm, peel of corm, leaf, and stalk, all from C. bicolor plant. Thirty milliliter of each medium was dispensed into 100 mL conical flask, plugged with cotton wool and sterilized in the autoclave at 121oC for 15 min. Each flask containing any of the carbon sources/substrates were aseptically inoculated with 1.2 mL (1.0 x 108 spores/mL) spore suspension of Aspergillus sp. (Org 2). The culture media were incubated at room temperature (28 ± 2°C) under static motion with intermittent manual shaking (to enhance mass-transfer) for 7 days. Two conical flasks were removed daily from each preparation. 10 mL of each cultured sample were withdrawn from culture into a centrifuge tube and the supernatant that resulted following centrifugation at 4000 rpm for 15 min to remove solids were used for analysis.
2.12 Analytical Methods in Enzymatic Hydrolysis of Caladium bicolor
The cell free extract obtained by centrifugation of the enzyme hydrolyzed broth at 4000rpm for 15 min was used for the analysis below:
2.12.1 Determination of Reducing Sugar Concentration
The cell free extract obtained was analyzed for total reducing sugars by Dinitrosalicylic acid method (Miller 1959). The DNS reagent was prepared by dissolving 1 g of 3, 5-dinitrosalicylic acid in 50 mL of distilled water and 30 g of sodium potassium tartrate tetrahydrate,
(KNaC4H4O64H2O) was slowly added. Twenty milliliters of 2 N NaOH (1.6 g NaOH in 20 mL of distilled water) was added. The preparation was diluted to a final volume of 100 mL with distilled water. 41
Glucose content of samples were analysed daily by transferring 1 mL of the DNSA reagent into a test tube containing 1 mL of the sample supernatant and placed in boiling water for 10 min. Thereafter, the tubes were allowed to cool, and 10 mL of sterile distilled water added. The colour intensity of the tubes were measured at 540nm using 772S Spectrophotometer (B-Bran Scientific and Instrument Company England) against a blank containing 1 mL of distilled water and 1 mL of DNSA to which 10 mL of distilled water was added after boiling. Absorbance readings were converted to glucose concentration using standard curve prepared.
2.12.2 Determination of the Starch Content of Caladium bicolor (the fleshy part of corm)
The determination of the starch content was carried out according to Astolfi-Filfo (1986). Iodine solution, 5 mL (0.5% KI and 0.15% I2) and 3 mL of the samples were mixed. The final volume was made up to 15 mL by addition of 7 mL distilled water. The absorbance was measured at 550 nm against a blank containing 5 mL of iodine solution and 10 mL of distilled water using 772S Spectrophotometer (B-Bran Scientific and Instrument Company England). Absorbance readings were converted to starch concentration using standard curve prepared.
2.12.3 Determination of the Cellulose Content of Caladium bicolor
Cellulose undergoes acetolysis with acetic/nitric reagent forming acetylated cellodextrins which get dissolved and hydrolyzed to form glucose molecules on treatment with 67% H2SO4. This glucose molecule is dehydrated to form hydroxymethyl furfural which forms green coloured product with anthrone and the colour intensity is measured at 630 nm.
In the determination of cellulose concentration, the reagents used include Acetic/ Nitric reagent (got by mixing 150ml of 80% acetic acid and 15ml of concentrated nitric acid); Anthrone reagent (got by dissolving 200mg anthrone in 100ml concentrated sulphuric acid, anthrone reagent is prepared fresh and chilled for 2 h before use) and 67% sulphuric acid.
The cellulose content of C. bicolor (the peel, leaf and stalk) was measured using Acetic/Nitric acid Anthrone reagent method of cellulose estimation as proposed by Updegroff (1969). Three millilitre acetic/nitric reagent was added to 1 g of the sample in a test tube and mixed in a vortex mixer. The tube was placed in a water-bath at 100°C for 30 min, allowed to cool and then centrifuged for 20 min. The supernatant was discarded and then the residue washed with distilled 42 water. Ten milliliters of 67% sulphuric acid was added and allowed to stand for 1 h. One milliliter of the above solution was diluted to 100 mL. To 1 mL of this diluted solution, 10 mL of anthrone reagent was added and mixed well. A blank was set with I mL distilled water and 10 mL anthrone reagent. The tubes were placed in a boiling water-bath for 10 min. Thereafter, the tubes were allowed to cool and the colour intensity measured at 630nm using 772S Spectrophotometer (B-Bran Scientific and Instrument Company England). Absorbance readings were converted to cellulose concentration using standard curve prepared.
2.13 Determination of Optimal Conditions on Hydrolysis of Caladium bicolor using the Selected Test Organism Aspergillus sp. (Org 2)
For optimization of the medium, the traditional method of “one variable at a time” approach was used by changing one component at a time while keeping the others at their original level. Original level of hydrolysis conditions: Time: 7 days, pH: 5.0, substrate concentration: 4% (w/v), 8 nitrogen source: 0.5% (NH4)2SO4 and 0.05% Yeast extract, and inoculum size: 4% (1.0 x 10 spores/ml).
2.13.1 Effect of Incubation Time
The rate of hydrolysis by the fungal isolate was studied by measuring daily reducing sugar and residual starch/cellulose concentration for a period of 7 days and the period of maximum reducing sugar production was determined.
2.13.2 Effect of Varying Initial pH of the Medium
The influence of initial pH on reducing sugar production was studied by adjusting pH of the culture medium before sterilization to the various pH (pH 3, 5, 7, and 9) with 0.1 M NaOH and 0.1 M HCL using a digital pH meter (Hanna Instrument- H196107, pHep pH Tester, Italy). The pH value that gave the highest reducing sugar production was determined.
2.13.3 Effect of Varying Substrate Concentration
Different concentrations of each of the substrates ranging from 1.0% to 5.0% were used in the preparation of the hydrolyzing medium with concentration increments of one. The substrate concentration with the highest reducing sugar production was determined. 43
2.13.4 Effect of Different Nitrogen Sources
The following nitrogen sources 5g/L were used to evaluate the influence of organic and inorganic nitrogen sources on hydrolysis of C. bicolor. The nitrogen sources used include (NH₄)₂SO₄, NaNO₃, Yeast extract, Peptone, and Soya-bean meal. The respective nitrogen sources were added as a sole source of nitrogen in the culture medium in place of (NH₄)₂SO₄ and yeast extract employed earlier in the mineral salt medium. The nitrogen source with the highest reducing sugar production was determined.
2.13.5 Effect of Varying Inoculum Size
Each of the culture medium was fed with varying inoculum size of the organism. The inoculum size was varied from 1% to 5% with concentration increments of one. The inoculum size concentration with the highest reducing sugar production was determined.
2.13.6 Time Course of Enzymatic Hydrolysis of Caladium bicolor
The various optimal conditions for starch and cellulose hydrolysis that produced maximal reducing sugar from all the earlier experiments were combined. The effect of time of incubation on the hydrolysis of the different substrates were studied under the determined optimal conditions for a period of 7 days and the period of maximum reducing sugar production was determined.
2.14 Fermentation
Separate Hydrolysis and Fermentation (SHF) was carried out in 100 mL conical flasks each containing 30 mL of medium, prepared using the various determined optimal conditions for starch and cellulose hydrolysis of C. bicolor. The medium was cotton plugged, sterilized, allowed to cool and aseptically inoculated with 1.0 x 108 spores/mL Aspergillus sp. The conical flasks were incubated under aerobic condition for 4-5 days depending on the day of maximum reducing sugar production for each of the substrates. Following hydrolysis, 60 mL each of the hydrolyzed culture medium in 100 mL conical flask was supplemented with 0.5% yeast extract. Aseptically, 1.2 mL (4.0 x 107 cells/mL) cell suspension of Saccharomyces sp. (Org 19) was seeded into the hydrolyzed flask. It was covered with cotton wool wrapped in aluminum foil and 44 masked to create microaerophilic environment. The fermentation media were incubated at room temperature (28 ± 2°C) under static motion with intermittent manual shaking (to enhance mass- transfer) for 7 days. Two conical flasks were removed daily from each preparation. 30 mL of each cultured sample was withdrawn from culture and centrifuged. The supernatant that resulted following centrifugation at 4000 rpm for 15 min to remove solids were used for analysis. The following analyses were done daily: determination of residual sugar and determination of ethanol produced. Residual starch and cellulose were determined on the first and last day of fermentation.
2.15 Measurement of Ethanol Concentration
Ethanol concentration was determined by the boiling iodometric method (Gwarr, 1987). The apparatus used are: retort stand (2), tripod stand, Bunsen burner, conical or flat bottom flask, cork, delivery tube, receiving tube (large test-tube),wire guaze, 100 mL measuring cylinder, and burette. While the reagents used for measurement of ethanol concentration were 10% potassium sulphate (K2SO4), 10 mL; deionized water or sample i.e. fermented broth containing ethanol, 10 mL; 42.5 g/L potassium dichromate (K2Cr2O7), 10 mL; 1:1 dilution of concentrated sulphuric acid (H2SO4), 10 mL; 10% potassium iodide (KI), 5 mL; 0.4% soluble starch, 2 mL and I M solution of sodium thiosulphate (Na2S2O3.5H2O), 24.8 g/100 mL.
Distillation
In a 50 mL flat bottom flask labeled “A”, 10 mL of 10% K2SO4 was added, and then 10 ml of deionized water for blank or 10 mL sample for test (fermented broth containing ethanol) was added to the flask. To the receiving tube labeled “B”, 10 mL K2Cr2O7 (42.5 g/L) was pippeted into a big test tube and then 10 mL of 1:1 dilution of conc. H2SO4 was added. Flask “A” was mounted on a retort (tripod) stand. Flask “A” was covered with a rubber bung carrying a delivery glass tube joined by a teflon tubing to another piece of glass tubing leading to the receiving tube “B” held in a position by a clamp. A low flame was ignited under flask “A” and adjusted so that the liquid in flask “A” boils with little frothing to a boiling point. The ethanol in the sample was distilled into the receiving tube “B” and was absorbed by the K2Cr2O7 and H2SO4. This made the colour change from yellow to greenish. Distillation was done until about half of the content of flask “A” has distilled over into tube “B”. Boiling was stopped and contents of tube “B” were 45 transferred to a 100 mL measuring cylinder and rinsed out with deionized water to make a total volume of 60 mL. The 60 mL was then transferred into a 500 mL flat bottom flask labeled “C” and 5 mL of 10% KI solution was added. The flask was swirled to mix and then allowed to cool to room temperature. After cooling, 2 mL of 0.4% soluble starch was added and mixed.
Titration
1 M solution of Na2S2O3.5H2O was put in a burette and initial reading taken. The mixture in the flask labeled “C” {containing 10 mL K2Cr2O7 (42.5 g/L), 10 mL 1:1 dilute conc. H2SO4, 5 mL 10% KI solution and 2 mL 0.4% starch solution} was titrated with the 1 M solution of
Na2S2O3.5H2O put in a burette until a blue colour appears. The final reading was then taken and subtracted from the initial reading to get a titer value. A value between 8.0 to 9.0 was regarded as a good blank titer value. The whole test was done in duplicate and the average titre obtained.
Ethanol content of the sample was read out by reference to a standard table of results (Appendix
IV); where a = blank titre and b = test titre. The blank titer is subtracted from the test titer and the value was then read out from the table under the blank value to get the ethanol concentration in the fermentation medium. The level of ethanol from table was multiplied by the dilution factor to get the percentage (v/v) ethanol, in the fermentation medium.
2.16 Statistical Analysis
Data obtained in this study were analyzed statistically using Microsoft Excel 2007 at 95% level of significance.
46
CHAPTER THREE
RESULTS
3.1 Proximate Analysis of Caladium bicolor (Wild Cocoyam)
Starch and cellulose are complex materials, it is essential to know the initial levels of these constituent in the starting material. Different parts of dried C. bicolor (the fleshy part of corm, peel of corm, leaf, and stalk) were analyzed for moisture, ash, fiber, fats, protein and total carbohydrate contents as shown in Table 3. Caladium bicolor is a potential material for producing ethanol due to its carbohydrate rich content with the peel of the corm having the highest total carbohydrate content (85.932% ± 0.014), followed by the fleshy part of corm (81.788% ± 0.015), then the stalk (73.883% ± 0.014), and finally the leaf having the least carbohydrate content (64.803% ± 0.012). The total carbohydrate was by difference according to AOAC (1995). Preliminary analysis of the samples showed that there was little or no starch content in the peel of corm, leaf and stalk (Table 3).
47
Table 3: Proximate Analysis of Caladium bicolor (Wild Cocoyam)
Parameters Proximate Composition (%) Analysed Fleshy Part of Peel of Corm Leaf Stalk Corm Moisture 9.246 ± 0.003 6.894 ± 0.003 9.534 ± 0.005 12.488 ± 0.007 Ash 2.275 ± 0.005 2.402 ± 0.009 2.937 ± 0.004 1.872 ± 0.005 Fibre 1.163 ± 0.003 1.441 ± 0.003 5.594 ± 0.004 5.189 ± 0.004 Fats 2.862 ± 0.003 1.918 ± 0.002 2.337 ± 0.008 1.485 ± 0.005 Protein 2.666 ± 0.005 1.413 ± 0.003 14.795 ± 0.003 5.083 ± 0.004 Total Carbohydrate 81.788 ± 0.015 85.932 ± 0.014 64.803 ± 0.012 73.883 ± 0.014 Starch 43.379 ± 1.489 ------Cellulose --- 76.768 ± 0.002 42.938 ± 0.003 69.931 ± 0.002
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3.2 Screening of the Isolated Microorganisms
3.2.1 Qualitative Screening of Isolates for Enzyme Production
A total of 18 isolates of both mould and yeast species were isolated from rotten corms of C. bicolor and fresh palm wine. The result showed that nine (9) isolates with carbohydrate degrading ability were obtained. Six of the isolates were able to produce both amylase and cellulase while the other three isolates (designated Org 6, Org 8, and Org 11) produced only amylase as shown in Table 4.
49
Table 4: Qualitative Screening of Isolates for Enzyme Production
Source of Isolate Isolates Culture Type Zone of Clearance
Amylase Cellulase
Rotten C. bicolor Org 1 Mould + + Rotten C. bicolor Org 2 Mould + + Rotten C. bicolor Org 3 Mould + + Rotten C. bicolor Org 4 Mould - - Palm Wine Org 5 Yeast - - Rotten C. bicolor Org 6 Mould + - Rotten C. bicolor Org 7 Mould + + Rotten C. bicolor Org 8 Mould + - Rotten C. bicolor Org 9 Mould - - Palm Wine Org 10 Yeast - - Rotten C. bicolor Org 11 Mould + - Rotten C. bicolor Org 12 Mould + + Rotten C. bicolor Org 13 Mould - - Rotten C. bicolor Org 14 Mould + + Palm Wine Org 15 Yeast - - Palm Wine Org 16 Yeast - - Palm Wine Org 17 Yeast - - Palm Wine Org 18 Yeast - -
Key: (+) Zone of Clearance; (-) No Zone of Clearance.
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3.2.2 Quantitative Screening of Isolates for Ability to Generate Reducing Sugars from C. bicolor (Glucose was used as Standard)
Quantitative screening of isolates for hydrolytic ability was carried out in a submerged culture condition.
Figure 5 shows the result of the quantitative screening of the fungal isolates using the fleshy part of corm as substrate. The highest reducing sugar released was found with Org 2 (20.063 mg/mL), followed by Org 14 (19.094 mg/mL), and the least Org 11 (6.634 mg/mL).
Figure 6 shows the result of the quantitative screening of the fungal isolates using the peel of corm as substrate. The highest reducing sugar released was found with Org 2 (10.357 mg/mL), followed by Org 14 (10.193 mg/mL), and the least Org 6 (1.457 mg/mL).
Figure 7 shows the result of the quantitative screening of the fungal isolates using the leaf as substrate, the highest reducing sugar released was found with Org 2 (7.120 mg/mL), followed by Org 1 (6.310 mg/mL). Org 6 and Org 8 produced the least amount of reducing sugar with a value of 2.427 mg/mL.
Figure 8 shows the result of the quantitative screening of the fungal isolates using the stalk as substrate, the highest reducing sugar released was found with Org 14 (12.460 mg/mL), followed by Org 2 (11.167 mg/mL), and the least Org 6 (0.647 mg/mL).
Figure 9 shows the comparison of the reducing sugar released from the various parts of C. bicolor during the quantitative screening by different fungal organisms. Altogether, Org 2 yielded the highest reducing sugar with the fleshy part of corm (20.063 mg/mL), peel of corm (10.357 mg/mL), and the leaf (7.120 mg/mL). Org 14 produced the highest reducing sugar with the stalk (12.460 mg/mL), followed by Org 2 (11.167 mg/mL). Based on the amount of reducing sugar released from the various C. bicolor parts by each of the isolates, Org 2 was selected as the most suitable isolate for the hydrolysis of the various parts of C. bicolor.
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Figure 5: Screening of the Fungal Isolates for Hydrolytic Ability using Fleshy Part of Corm as Substrate.
52
Figure 6: Screening of the Fungal Isolates for Hydrolytic Ability using Peel of Corm as Substrate.
53
Figure 7: Screening of the Fungal Isolates for Hydrolytic Ability using Leaf as Substrate.
54
Figure 8: Screening of the Fungal Isolates for Hydrolytic Ability using Stalk as Substrate.
55
Fleshy Part Peel of Corm Leaf Stalk Of Corm
Figure 9: Comparison of the Hydrolytic Ability of the Various Organisms.
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3.2.3 Screening Yeast Isolates for Ethanol Production
In the screening of nine (9) yeast isolates for fermentative ability, four (4) alcohol productive strains were selected based on the amount of gas produced. Two from healthy corms of C. bicolor and two from palm wine (Table 5).
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Table 5: Screening Yeast Strains for Ethanol Production
Source of Isolate Isolates Degree of Gas Production Degree of Gas Production
24 h 48 h
Palm Wine Org 5 ++ ++
Palm Wine Org 10 - +
Palm Wine Org 15 +++ ++++
Palm Wine Org 16 - -
Palm Wine Org 17 ++ +
Palm Wine Org 18 - +
Palm Wine Org 19 +++++ +++++
Healthy C. bicolor CB 1 ++++ ++++
Healthy C. bicolor CB 2 +++++ ++++
KEYS: (+++++) Very High Gas Production; (++++) High Gas Production; (+++) Moderate Gas Production; (++) Low Gas Production; (+) Very Low Gas Production; (-) No Gas Production.
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3.3 Characterization/Identification of Selected Isolates
3.3.1 Characterization/Identification of Mould Isolates
The entire mould isolate were characterized and identified macroscopically and microscopically using conventional microbiological method as described by Michael et al. (2011) and Ellis et al. (2007) as shown in Table 6.
3.3.2 Characterization/Identification of Yeast Strains
The four selected yeast isolates Org 15, Org 19, CB 1, and CB 2, were tentatively identified to belong to the genus Saccharomyces using the method of Yarrow, 1998; Kurtzman et al., 1999; and Queshi et al., 2007 (Table 7). The four yeast strains showed similar cultural morphology. Fermentation trial was also conducted to be able to select the best yeast isolate to work with. From the result obtained, Org 19 did best and was selected and the other isolates screened out on the bases of insignificant CO2 production compared to Org 19.
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Table 6: Characterization/Identification of Mould Isolates
Isolates Macroscopy Microscopy Probable Organism Org 1 Colonies are granular, flat, Hyphae are septate (branched septate), Aspergillus often with radial grooves, conidial heads are typically radiate, grow rapidly, produce biseriate but having some heads with yellow-green conidia, and phialides borne directly on the vesicle reverse side yellow. (uniseriate). Conidiophore stipes are hyaline and coarsely roughened. Conidia are globose, pale green and conspicuously echinulate.
Org 2 Colonies grows rapidly, Hyphae are septate (branched septate), Aspergillus producing dark-brown to conidial heads are large, globose, dark black conidial heads. The brown and radiates, conidiophores stipes reverse side is without are smooth-walled, and hyaline. Conidial colour. head is biseriate with the phialides borne on brown, septate metulae. Conidia are globose, dark brown and rough-walled. Org 3 Colonies show typical Hyphae are septate (branched septate), Aspergillus blue-green surface conidial heads are typically columnar and pigmentation with a suede- uniseriate. Conidiophore stipes are short, like surface consisting of a smooth-walled and have conical-shaped dense felt of terminal vesicles which support a single conidiophores. Grows row of phialides on the upper two thirds rapidly and the reverse is of the vesicle. Conidia are produced in pale yellowish. basipetal succession forming long chains and are subglobose, green and rough- walled to echinulate. Org 6 Colonies are fast growing Produces septate, hyaline hyphae, Penicillium and become fully mature in conidiophores branched, biverticillate about 5 days. The surface (phialides extend from branches which appearance is velvety. The extend from the hyphae), Phialides colony colour is blue-green ampule shaped producing smooth, single with a white edge. The celled conidia which extend as basipetal reverse is reddish-brown. chains and conidia shape is ovoid.
Org 7 Colonies are typically plain Conidial heads are short columnar and Aspergillus green in colour with dark biseriate. Conidiophore stipes are usually red-brown cleistothecia short, brownish and smooth walled. developing within and Conidia are globose and rough-walled. upon the conidial layer. Reverse side is olive. Org 8 Fast growing, colony Chains of single-celled conidia are Penicillium colour blue-green, with produced in basipetal succession from a 60
white apron, consisting of a specialised conidiogenous cell called dense felt of a phialide. Phialides are produced from conidiophores. branched metulae, giving a brush-like appearance. The branching pattern is two- stage branched (biverticillate asymmetrical). Conidiophores are hyaline and smooth-walled. Phialides are flask- shaped, consisting of a cylindrical basal part and a distinct neck. Conidia are in long chains, in columns, globose, hyaline, and smooth-walled.
Org 11 Colonies are typically plain Conidial heads are short columnar and Aspergillus green in colour with dark biseriate. Conidiophore stipes are usually red-brown cleistothecia short, brownish and smooth walled. developing within and Conidia are globose and rough-walled. upon the conidial layer. Reverse side is olive. Org 12 Colonies are rapidly Sporangiophores are borne from a Rhizopus growing with some sporangium located on the collumella at tendency to collapse and the apical end of the sporangiophore, are fills the petri dish with smooth-walled and non-septate. Sporangia fluffy, cotton-candy like are globose, with a flattened base, greyish growth in less than 5 days. black, powdery in appearance, and many Growth is whitish in spored. Columellae and apophysis colour, turns brown with together are globose, soon collapsing to an age. umbrella-like form after spore release. Sporangiospores are angular, subglobose, with ridges on the surface. Org 14 Colonies grows rapidly, Hyphae are septate (branched septate), Aspergillus producing black conidial conidial heads are large, globose, dark heads. The reverse side is brown and radiates; conidiophores stipes without colour. are smooth-walled, and hyaline. Conidial head is biseriate with the phialides borne on brown, septate metulae. Conidia are globose, black and rough-walled.
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Table 7: Cell Morphology and Physiological Characterizations of Yeast Strains
Isolates Cultural Probable
Morphology Genus
Glucose Galactose Maltose Fructose Sucrose Raffinose Xylose Lactose Nitrate Org 15 Smooth, A/G A A/G A A/G A - A NG Saccharomyces
circular, &
creamy
Org 19 Smooth,circular A/G A/G A/G A/G A/G A/G - A NG Saccharomyces
, & creamy
white
CB 1 Smooth, A/G A/G A/G A/G A/G A/G - A NG Saccharomyces
circular, &
creamy
CB 2 Smooth, A/G A/G A/G A/G A/G A/G - A NG Saccharomyces
circular, &
creamy
KEYS: (A/G) Acid and Gas Production; (A) Acid Production Only; (-) No Acid and No Gas Production; (NG) No Growth.
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3.4 Determination of Optimal Conditions on Hydrolysis of Various Parts of Caladium bicolor using the Selected Test Fungus Aspergillus sp. (Org 2)
3.4.1 Effect of Incubation Time
The rate of hydrolysis by Aspergillus sp. (Org 2) was studied by measuring daily reducing sugar concentration and residual carbohydrate (starch and cellulose) for a period of 7 days. The result showed that maximum reducing sugar production occurred on day six when the fleshy part of corm was used as substrate, and day five when peel of corm, leaf and stalk of C. bicolor were used as substrate. For all the substrates, there was a continuous decrease in carbohydrate concentration (Figures 10-13).
Reducing sugar produced by the test organism (Aspergillus Org 2) was compared in composite media containing the various plant parts as substrate (Figure 14). The result showed that the fungus exhibited significantly higher productivity of reducing sugar in liquid medium containing the fleshy part of corm compared to the others while the leaf of C. bicolor had the lowest reducing sugar production.
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Figure 10: Effect of Incubation Time on Hydrolysis of the Fleshy Part of Corm.
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Figure 11: Effect of Incubation Time on Hydrolysis of the Peel of Corm.
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Figure 12: Effect of Incubation Time on Hydrolysis of the Leaf.
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Figure 13: Effect of Incubation Time on Hydrolysis of the Stalk.
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Fleshy Part of Corm
Peel of Corm
Leaf
Stalk
Figure 14: Comparison of the Reducing Sugar Produced from the Various Substrates with Effect to Incubation Time.
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3.4.2 Effect of Varying Initial pH of the Medium
In this study, the influence of the varying initial pH between the ranges of pH 3 to 9 on hydrolysis of the various C. bicolor parts was assayed. Reducing sugar was produced for all the pH evaluated and with all the substrates (Figures 15-18). Figure 19 is a comparison of the reducing sugar released from the various substrates by the test organism which shows that pH 5 was the optimum initial pH for the production of reducing sugar from all the substrates.
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Figure 15: Effect of Varying Initial pH on Hydrolysis of the Fleshy Part of Corm.
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Figure 16: Effect of Varying Initial pH on Hydrolysis of the Peel of Corm.
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Figure 17: Effect of Varying Initial pH on Hydrolysis of the Leaf.
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Figure 18: Effect of Varying Initial pH on Hydrolysis of the Stalk.
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Fleshy Part Peel of Corm Leaf Stalk Of Corm
Figure 19: Comparison of the Effect of Varying Initial pH on Hydrolysis of the Different Plant Parts.
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3.4.3 Effect of Varying Substrate/Carbon Concentration
Reducing sugar production by the test organism was compared among media containing different substrates at different concentrations ranging from 1% - 5% w/v (Figures: 20-24). The result shows that increase in substrate/carbon concentration also resulted to an increase in reducing sugar concentration, and increase in substrate concentration beyond the level that gave the optimum reducing sugar did not result in proportionate increase in reducing sugar yield.
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Fleshy Part of Corm
Fleshy Part of Corm
Fleshy Part of Corm
Fleshy Part of Corm
Fleshy Part of Corm
Figure 20: Effect of Varying Substrate/Carbon Concentration on Hydrolysis of the Fleshy Part of Corm.
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Peel of Corm
Peel of Corm
Peel of Corm
Peel of Corm
Peel of Corm
Figure 21: Effect of Varying Substrate/Carbon Concentration on Hydrolysis of the Peel of Corm.
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Figure 22: Effect of Varying Substrate/Carbon Concentration on Hydrolysis of the Leaf.
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Figure 23: Effect of Varying Substrate/Carbon Concentration on Hydrolysis of the Stalk.
79
Fleshy Part Peel of Corm Leaf Stalk of Corm
Figure 24: Comparison of the Effect of Varying Substrate/Carbon Concentration on Hydrolysis of the Different Plant Parts.
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3.4.4 Effect of Different Nitrogen Sources
The effect of various organic and inorganic nitrogen sources (ammonium sulphate, sodium nitrate, yeast extract, peptone and soya bean) on the hydrolysis of the various substrates was determined. Figures 25-29 depicts the effect of the various nitrogen sources with the various plant parts as carbon source on production of reducing sugar using the test organism. Soya bean showed the best result in enhancing the hydrolysis of the fleshy part of the corm and the peel of the corm, while yeast extract and ammonium sulphate were the best in the hydrolysis of the leaf and stalk, respectively.
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Figure 25: Effect of Different Nitrogen Sources on Hydrolysis of the Fleshy Part of Corm.
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Figure 26: Effect of Different Nitrogen Sources on Hydrolysis of the Peel of Corm.
83
Figure 27: Effect of Different Nitrogen Sources on Hydrolysis of the Leaf.
84
Figure 28: Effect of Different Nitrogen Sources on Hydrolysis of the Stalk.
85
Fleshy Part Peel of Corm Leaf Stalk of Corm
Figure 29: Comparison of the Effect of Different Nitrogen Sources on Hydrolysis of the Different Plant Parts.
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3.4.5 Effect of Varying Inoculum Size (1.0 x 108 spores/ml)
Effect of inoculum size on the hydrolysis of substrates was assayed using 1%(v/v), 2%(v/v), 3%(v/v), 4%(v/v) and 5%(v/v) inoculum size (Figures 30-34). At 2% inoculum size, maximum reducing sugar was produced with the fleshy part of corm, peel of corm, and stalk with values 12.484 mg/mL, 15.418 mg/mL, and 11.024 mg/mL respectively (Figures 30, 31, & 33). Maximum reducing sugar (4.831 mg/mL) was produced at inoculum size of 5%(v/v) for the leaf (Figure 32). Figure 34 shows a comparison of the effect of varying inoculum size on hydrolysis of the various parts of C. bicolor.
87
Figure 30: Effect of Varying Inoculum Size on Hydrolysis of the Fleshy Part of Corm.
88
Figure 31: Effect of Varying Inoculum Size on Hydrolysis of the Peel of Corm.
89
Figure 32: Effect of Varying Inoculum Size on Hydrolysis of the Leaf.
90
Figure 33: Effect of Varying Inoculum Size on Hydrolysis of the Stalk.
91
Fleshy Part Peel of Corm Leaf Stalk of Corm
Figure 34: Comparison of the Effect of Varying Inoculum Size on Hydrolysis of the Different Plant Parts.
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3.4.6 Time Course of Enzymatic Hydrolysis of Various Parts of Caladium bicolor
The optimal conditions were combined in single fermentation for each substrate and reducing sugar released measured.
Figure 35 shows the result of the reducing sugar released from the fleshy part of corm of C. bicolor under the following culture conditions: pH 5, 5% substrate concentration, 0.5% soya bean as nitrogen source and 2% inoculum concentration. The maximum reducing sugar produced was on Day 5 (23.297 mg/mL).
Figure 36 shows the result of the reducing sugar released from the peel of corm of C. bicolor under the following culture conditions: pH 5, 5% substrate concentration, 0.5% soya bean as nitrogen source and 2% inoculum concentration. The maximum reducing sugar produced was on Day 4 (18.013 mg/mL).
Figure 37 shows the result of the reducing sugar released from the leaf of C. bicolor under the following culture conditions: pH 5, 4% substrate concentration, 0.5% yeast extract as nitrogen source and 5% inoculum concentration. The maximum reducing sugar produced was on Day 4 (6.667 mg/mL).
Figure 38 shows the result of the reducing sugar released from the stalk of C. bicolor under the following culture conditions: pH 5, 5% substrate concentration, 0.5% ammonium sulphate as nitrogen source and 2% inoculum concentration. The maximum reducing sugar produced was on Day 5 (15.320 mg/mL).
Figure 39 is a comparison of the sugar yield during time course of enzymatic hydrolysis of various parts of C. bicolor. In all, the highest reducing sugar produced was from fleshy part of corm (23.297 mg/mL), followed by the peel of corm (18.013 mg/mL), then the stalk (15.320 mg/mL), and lastly the leaf (6.667 mg/mL).
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Figure 35: Time Course of Hydrolysis of the Fleshy Part of Corm.
94
Figure 36: Time Course of Hydrolysis of the Peel of Corm.
95
Figure 37: Time Course of Hydrolysis of the Leaf.
96
Figure 38: Time Course of Hydrolysis of the Stalk.
97
Fleshy Part Peel of Corm Leaf Stalk of Corm
Figure 39: Comparison of the Sugar Yield from Different Plant Parts during Time Course of Hydrolysis.
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3.5 Fermentation
Ethanol was produced from the hydrolysate of the fleshy part of corm, peel of corm, leaf and stalk (the different plant parts) on inoculation with Saccharomyces (Org 19) at 2% inoculum concentration (v/v) containing 4.0 x 107 cells/mL. From the result, the medium with the hydrolysate from “stalk of C. bicolor” fermented fastest (3rd day) compared to the hydrolysate from the other substrates with ethanol yield 0.280% (Figure 43). The hydrolysates from the fleshy part of corm, peel of the corm, and the leaf produced their maximum ethanol on the 5th day with the following concentrations 0.485%, 0.210%, and 0.380% respectively (Figures 40- 42). A mixed substrate hydrolysate was also fermented containing 1.2% w/v each, of the various substrates (fleshy part of corm, peel of corm, leaf and stalk) and produced maximum ethanol on the 6th day with value 0.280% (Figure 44).
The data represented in Figure 45 shows the percentage ethanol yield from the different substrate hydrolysates when fermented by Saccharomyces sp. (Org 19). Fleshy part of corm gave the highest ethanol yield (0.485%).
99
Figure 40: Ethanol Production from Hydrolysate of Fleshy Part of Corm and Reducing Sugar Utilization.
100
Figure 41: Ethanol Production from Hydrolysate of Peel of Corm and Reducing Sugar Utilization.
101
Figure 42: Ethanol Production from Hydrolysate of Leaf and Reducing Sugar Utilization.
102
Figure 43: Ethanol Production from Hydrolysate of Stalk and Reducing Sugar Utilization.
103
Figure 44: Ethanol Production from Hydrolysate of Mixed Substrate (Various Plant Parts) and Reducing Sugar Utilization.
104
Fleshy Part Peel of Corm Leaf Stalk Mixed of Corm Substrate
Figure 45: Percentage Ethanol Yield of the Different Substrates.
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CHAPTER FOUR
DISCUSSION AND CONCLUSION
4.1 Screening of Isolates for Enzyme Production
The use of starch agar plate and iodine for detecting amylase (hydrolytic enzyme) producing microorganisms has been reported by Akpan et al. (1999); Mishra and Behera, (2008); Suganthi et al. (2011) and Geetha et al. (2011). This procedure was employed in the screening of eighteen fungal isolates in this work. Results obtained showed starch hydrolytic ability for nine fungal isolates (Table 4). This is in line with the work of Geetha et al. (2011) who screened sixteen fungal isolates including Rhizopus sp., Aspergillus flavus., Penicillium sp., Trichoderma viride and Trichoderma sp. Among the isolates, Aspergillus sp. exhibited higher amylolytic activity in starch agar medium. The clear zone observed indicated that the organisms synthesized amylases for the hydrolysis of starch in their environment. The areas containing un-hydrolysed starch exhibited a blue black colour.
Six out of eighteen fungal isolates (Table 4) tested positive for cellulose hydrolyzing ability. This confirms the earlier reports of Desvaux et al. (2000) and Khokhar et al. (2012) that only a few fungi possess the ability to degrade cellulose. All the isolates selected from the PDA-Congo red stained plates produced detectable quantities of cellulase during hydrolysis of the various Caladium bicolor parts, hence the use of PDA-Congo red stained plates present a simple and easy way of screening for cellulolytic microbes.
Both fungi and bacteria have been heavily exploited for their abilities to produce a wide variety of cellulase and amylase. However, emphasis has been placed on the use of fungi because of their ability to produce profuse amount of extracellular cellulases and amylases which are easily extracted and purified. The data presented in this study shows that the fungal isolate Aspergillus sp. (Org 2) had a higher amylase and cellulase activity compared to other fungal isolates. Compared to other fungi, .Aspergillus spp. has been used mostly for the simultaneous production of cellulase and amylase (Ray et al., 1993; Abo-state et al., 2010; Khan and Yadav, 2011; Adejuwon et al., 2012). The current findings suggest that the Aspergillus sp. isolated in this work has a multi-enzyme system. Multi-enzyme systems have been reported by Syu and Chen (1997) 106 and Pothiraj et al. (2006). According to the authors, bacteria (Bacillus subtilis, B. licheniformis, Escherichia species, and Clostridium species) and fungi (Trichoderma, Aspergillus, Rhizopus, Penicillium and Candida species) possess multi-amylase enzyme (, and glucoamylase) and cellulase (exoglucanase, endoglucanase and –glucosidase) enzyme systems. Also Nwagu et al. (2012), working on the amylase and cellulase production ability of some microorganisms, screened fungal isolates Aspergillus spp., Mucor sp., Penicillium sp., Rhizopus sp., Enterobacter sp., Escherichia sp., Bacillus spp. and Pseudomonas sp. Out of the ten isolates with carbohydrate degrading ability, all including fungi and bacteria showed multi-enzyme system except Escherichia sp. that produced only amylase.
Pothiraj et al. (2006) reported that Rhizopus stolonifer cultivated on cassava waste gave a higher cellulase activity than Aspergillus niger cultivated on the same cassava waste. This is contrary to our observation as Aspergillus sp. (Org 2) elicited the highest cellulose activity among other fungi screened, quantified based on the amount of reducing sugar produced. Our observation is in conformity with the work of Panda et al. (2012) who reported that Aspergillus species (A. niger, A. terreus, A. stellatus, A. flavus, and A. fumigatus) showed highest pectinolytic and cellulolytic activity amongst twenty five fungal isolates screened. Also, Brindha et al. (2011) reported that out of the three isolates (Aspergillus sp., Penicillium sp. and Trichoderma sp.) screened for amylase and cellulase production in submerged fermentation, Aspergillus sp. had the highest amylase activity and moderate cellulase activity. The higher amylase and cellulase activity of Aspergillus sp. (Org 2) used in this study might be due to the good growth of its mycelial biomass on its source (rotten C. bicolor corm) which led to higher glucose production. Also appropriate pH, aeration and temperature condition could have influenced the results.
4.2 Optimal Production Studies (Hydrolysis of Starch and Cellulose)
The cultural conditions were optimized for the increased production of amylase and cellulase. For initial medium optimization, the traditional method of “one variable at a time” approach was used by changing one component at a time while keeping the others at their original level.
Time course study revealed that the test organism was capable of producing amylase and cellulase as evidenced by the reduction of starch and cellulose in the fermentation medium, followed by the increased production of reducing sugar (Figures 10-14). The various parts of C. 107 bicolor was rapidly hydrolyzed to produce maximum reducing sugar by the 6th day for fleshy part of corm and 5th day for the peel of corm, leaf and stalk. Gupta et al. (2008) and Chimata et al. (2010) reported that the maximum production of alpha-amylase by Aspergillus spp. was achieved after 5 days of incubation while Omolasola et al. 2008 reported maximum cellulase production on day 5 of hydrolysis. In all the substrates, the concentration of starch/cellulose decreased gradually as the fermentation period increased. It was observed that starch was more rapidly degraded compared to cellulose. While the residual starch content for the fleshy part of corm was 0.047mg/mL, residual cellulose content for the peel of corm, leaf and stalk was 2.601mg/mL, 2.698mg/mL, and 2.601mg/mL respectively. Cellulose is well protected by the matrix of lignin and hemicellulose in macrofibrils thereby limiting the access of cellulolytic enzymes and resulting in the slow rate of its conversion to sugar. Due to this, pretreatment of cellulolytic materials is widely embraced to increase the rate of hydrolysis of cellulose to fermentable sugars (Galbe and Zacchi, 2002). Pretreatments may be chemically based. A main disadvantage of this pretreatment method is different chemical inhibitors might be produced during the pretreatment with reduced cellulase activity (Sun and Cheng, 2002). Biological pretreatments are more effective, economical, eco-friendly and less health hazardous as compared to the physicochemical or chemical-based pretreatment approaches (Asgher et al., 2012); however, slow rate of action and long pretreatment times are its challenges (Sun and Cheng, 2002). In addition to this, most of the lignolytic microorganisms solubilize/consume not only lignin but also hemicellulose and cellulose. Because of these drawbacks/limitations the biological pretreatment faces techno-economic barriers and therefore is less attractive commercially (Eggeman & Elander, 2005).
Enzymes have an optimal pH range within which they are most effective and therefore substrate pH affects fungal growth and productivity (Przybylowicz and Donoghue, 1990). From the result, the optimal initial pH was pH 5 for both starch and cellulose degradation by the Aspergillus sp. (Figures 15-19). Low reducing sugar was observed in the fermentation media at initial pH 3, as pH increased there was gradual increase in reducing sugar production up to pH 5, after which a continuous decrease in fermentable sugar was observed for all the substrates. Most of fungal cultures prefer a slightly acidic pH in the medium for growth and enzyme biosynthesis (Haltrich, 1996), in agreement with the results obtained. In literature optimum production of alpha amylase was found to be best at pH 5.0 for both free and immobilized cells of Aspergillus niger. Below 108 and above this pH, production of alpha amylase was significantly lower (Gupta et al., 2010). Ogier et al. (1999) and Taherzadeh & Karimi (2007) reported that enzymatic hydrolysis of cellulosic materials is preferred over acid hydrolysis due to milder processing conditions such as pH at 4.5–5.0. This supports the finding of Lee et al. (2002) who reported that the pH optimum of β- glucosidase was between pH 5 and 6.
Maximum reducing sugar production was observed with 5% substrate concentration for fleshy part of the corm, peel of corm and stalk while for the leaf, the optimal substrate concentration was 4% (Figures 16-20). It was observed that increase in substrate concentration led to increase in reducing sugar production while further increase in substrate concentration beyond the level that gave the optimum glucose did not result in proportionate increase in glucose yield. This is in line with the report of Omemu et al. (2004) who reported that amylase activity increased with increase in the starch concentration from 1% to 3%. Beyond 3%, there was a decline in amylase activity. This can be explained with the report of Omojasola et al. (2008) that natural substrates containing different minerals apart from carbon may serve as nutrient supplements; hence increase in substrate concentration leads to increase in these nutrients which may adversely affect the cell growth and metabolite production. A decrease in production beyond optimum concentration may be as a result of the inhibitory effect of accumulated cellobiose and cellodextrins of low degree of polymerization to the growth medium (Gilkes et al., 1984). It might also be due to the specific binding of the enzymes with the substrates (Gilkes et al., 1984). This can also be due to the high viscosity of the medium, which decreases the oxygen supply to the cells and retards cell division resulting in low production of the desired metabolite (Fritsche, 1999).
Among the various organic and inorganic nitrogen sources, the maximum reducing sugar produced was obtained when soya bean was used as a sole nitrogen source for fleshy part of corm and peel of corm while yeast extract and (NH4)2SO4 was best for leaf and stalk, respectively (Figures 21-25). The high yield of reducing sugar in media containing soya bean as the sole nitrogen source when the fleshy part of corm and peel of corm were used as substrate might be as a result of its components. Osho and Dashiell (1998) reported that soya bean which has less purchase cost has about 40% protein, 30% carbohydrates, 20% oil and 10% mineral. The result is also in line with the work of Goyal et al. (2005) who reported that soybean meal was the 109 best nitrogen source for raw starch digesting thermostable α-amylase production by the Bacillus sp. I-3 strain. Among the inorganic nitrogen sources ammonium sulphate elicited the highest cellulase activity determined by the amount of reducing sugar produced when C. bicolor stalk was hydrolyzed and is in agreement with the report of Sethi et al. (2013) that among the various nitrogen sources tested, ammonium sulphate was found to be the best nitrogen source for cellulase production by E. coli, Bacillus, Pseudomonas and Serratia. Gottschalk et al. (2013) also reported maximal β-glucosidase production by A. awamori, using yeast extract.
Effect of inoculum concentration as shown in figures 26-30 showed that 2% inoculum concentration gave the highest reducing sugar for fleshy part of corm, peel of corm, and stalk while 5% inoculum concentration gave the highest reducing sugar production for leaf. There was a decrease in amounts of reducing sugar production at inoculum concentrations above the optimal. This result is consistent with the work of Omojasola et al. (2008) whose result showed a decrease in the concentration of fermentable sugar after the optimum using Aspergillus. The decrease in glucose production with further increase in inoculum might be due to clumping of cells which could have reduced sugar and oxygen uptake rate and also, enzyme release (Srivastava, et al., 1987). Amadi and Okolo (2012) also reported that clumped morphology resulted in low yield during the production of raw starch digesting amylase (RSDA) by Aspergillus carbonarius.
To achieve high enzyme yield, efforts are made to develop a suitable medium for proper growth and maximum secretion of enzyme, using an adequate combination of carbohydrates, nitrogen and minerals (Goyal et al., 2005; Sodhi et al., 2005). The above optimal conditions were combined in single fermentations for each substrate using Aspergillus sp. (Org 2), then amylase and cellulase activity were measured by the amount of reducing sugar released and also the amount of residual starch/cellulose (Figures 31 - 35). From the use of different carbohydrate sources in this study, fleshy part of corm proved to be the best substrate for reducing sugar production (Figure 35). In the presence of fleshy part of corm at concentration of 5% (w/v), the reducing sugar yield reached 23.297 mg/ml after 5days of fermentation, while in the presence of peel of corm, and stalk at the same concentration, reducing sugar yield was 18.013 mg/ml (Day 4) and 15.320 mg/ml (Day 5), respectively. At 4% concentration (w/v), the reducing sugar yield for leaf of C. bicolor was 6.667 mg/ml on Day 4. Caritas and Humphrey (2006); Narasimha et al. 110
(2006); Omojasola et al. (2008) and Amaeze et al. (2015) also gave similar time course reports of maximum glucose yield on 5th day of fermentation using A. niger. The current study showed that the susceptibility of the various parts of C. bicolor to the crude enzymes of Aspergillus sp. (Org 2) was significantly dependent on the carbohydrate source, incubation time and other fermentation conditions. This agrees with earlier reports by Okolo et al. (1995) that the susceptibility of starch granules to digestion by amylase is dependent on starch source and length of amylase treatment.
During the hydrolyzing process in this current study, decrease in reducing sugar concentration was observed after the optimal reducing sugar production. This can be explained by the report of Bull (1972) and Demain (1972) that most extracellular catabolite enzymes seem to be affected by catabolite or end-product feedback repression. Catabolite repression of enzyme synthesis has been reported in static submerged fermentations (SMF) of bacteria and fungi (Tomanaga, 1966; Heinekes and O’Connor, 1972).
4.3 Fermentation
Ethanol fermentation of the hydrolysates of the fleshy part of corm, peel of corm, leaf, stalk and mixed substrate of C. bicolor by Saccharomyces sp. (Org 19) produced 0.485%, 0.210%, 0.380%, 0.280%, and 0.280% ethanol yield, respectively. This result reveals a higher productivity with the hydrolysate of the fleshy part of corm than in the other hydrolysates. This is due to the presence of more reducing sugar in fleshy part of corm which could be fermented to ethanol than in the other hydrolysates. Agu et al. (2014) while studying ethanol production from local tubers also reported higher ethanol yields with fleshy part of tuber of Dioscorea bulbifera than with the peel of tuber. Studies on high substrate fermentation have attested to the fact that higher substrate concentration results in higher ethanol concentrations (Laopaiboon et al., 2008 & 2009).
4.4 CONCLUSION
Fuel derivation from non-food starchy plants and cellulosic biomass are essential in order to overcome our excessive dependence on petroleum for liquid fuels and also address the build-up of greenhouse gases that cause global climate change. This study revealed that Caladium bicolor, 111 a non-human edible plant which does not find any significant commercial use in Nigeria can serve as ideal substrate for ethanol production instead of being left behind for natural degradation or burning. The use of this biomass (C. bicolor) for bio-ethanol production will help solve the problem of the diminishing fossil fuel reserve and at the same time not lead to food insecurity issues that can come about while using staple foods. It will also help to reduce the dumping and burning of these agricultural waste materials therefore resulting in a cleaner environment with fewer environmental hazards and concurrent increase in economic gain.
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APPENDIX I
Glucose Standard Curve (DNSA Method)
137
APPENDIX II
Starch Standard Curve (Starch-Iodine Method)
138
APPENDIX III
Cellulose Standard Curve (Acetic/Nitric Reagent and Anthrone Reagent Method)
139
APPENDIX IV
REFERENCE STANDARD TABLE FOR ETHANOL DETERMINATION (Boiling Iodometric Method). a –b a
8.0 8.1 8.2 8.3 8.4 8.5 8.6 8.7 8.8 8.9 9.0
0.1 0.02 0.02 0.02 0.02 0.01 0.01 0.01 0.01 0.01 0.01 0.01
0.2 0.03 0.03 0.03 0.03 0.03 0.03 0.03 0.03 0.03 0.03 0.03
0.3 0.05 0.05 0.05 0.05 0.04 0.04 0.04 0.04 0.04 0.04 0.4
0.4 0.06 0.06 0.06 0.06 0.06 0.06 0.06 0.06 0.06 0.06 0.06
0.5 0.08 0.08 0.08 0.08 0.07 0.07 0.07 0.07 0.07 0.07 0.07
0.6 0.09 0.09 0.09 0.09 0.09 0.09 0.09 0.09 0.09 0.08 0.08
0.7 0.11 0.11 0.11 0.11 0.10 0.10 0.10 0.10 0.10 0.10 0.10
0.8 0.13 0.12 0.12 0.12 0.12 0.12 0.12 0.11 0.11 0.11 0.11
0.9 0.14 0.14 0.14 0.14 0.13 0.13 0.13 0.13 0.13 0.13 0.13
1.0 0.16 0.15 0.15 0.15 0.15 0.15 0.13 0.14 0.14 0.14 0.14
1.1 0.17 0.17 0.17 0.17 0.16 0.16 0.16 0.16 0.16 0.15 0.15
1.2 0.19 0.18 0.18 0.18 0.18 0.17 0.17 0.17 0.17 0.17 0.17
1.3 0.20 0.20 0.20 0.20 0.19 0.19 0.19 0.19 0.19 0.18 0.18
1.4 0.22 0.22 0.21 0.21 0.21 0.20 0.20 0.20 0.20 0.20 0.19 140
1.5 0.23 0.23 0.23 0.23 0.22 0.22 0.22 0.22 0.22 0.21 0.21
1.6 0.25 0.25 0.24 0.24 0.24 0.24 0.23 0.23 0.23 0.23 0.22
1.7 0.27 0.26 0.26 0.26 0.25 0.25 0.25 0.24 0.24 0.24 0.24
1.8 0.28 0.28 0.27 0.27 0.27 0.27 0.26 0.26 0.26 0.25 0.25
1.9 0.30 0.29 0.29 0.29 0.28 0.28 0.28 0.27 0.27 0.27 0.26
2.0 0.31 0.31 0.30 0.30 0.30 0.29 0.29 0.29 0.28 0.28 0.28
2.2 0.34 0.34 0.34 0.33 0.33 0.32 0.32 0.32 0.31 0.31 0.31
2.3 0.38 0.37 0.37 0.36 0.36 0.35 0.35 0.34 0.34 0.34 0.33
2.5 0.39 0.38 0.38 0.38 0.37 0.37 0.36 0.36 0.35 0.35 0.35
2.6 0.41 0.40 0.39 0.39 0.38 0.38 0.38 0.38 0.37 0.37 0.36
2.8 0.44 0.43 0.43 0.42 0.42 0.41 0.41 0.40 0.40 0.39 0.39
3.0 0.47 0.46 0.45 0.45 0.45 0.44 0.44 0.43 0.43 0.42 0.42
3.5 0.55 0.54 0.53 0.52 0.52 0.52 0.51 0.50 0.50 0.49 0.49
4.0 0.63 0.62 0.61 0.60 0. 60 0.59 0.58 0.58 0.57 0.56 0.56
4.2 0.66 0.65 0.64 0.63 0.63 0.6 0.61 0.60 0.60 0.59 0.59
4.4 0.69 0.68 0.67 0.66 0.65 0.64 0.63 0.62 0.62 0.62 0.61
4.5 70 0.69 0.69 0.68 0.67 0.66 0.65 0.65 0.64 0.63 0.63
4.6 0.72 0.71 0.70 0.69 0.68 0.68 0.67 0.66 0.65 0.65 0.64
4.8 0.75 0.74 0.73 0.72 0.71 0.71 0.70 0.69 0.68 0.66 0.66 141
5.0 0.78 0.77 0.76 0.75 0.74 0.74 0.73 0.72 0.71 0.70 0.69
5.2 0.81 0.80 0.79 0.78 0.77 0.77 0.76 0.75 0.74 0.73 0.72
5.4 0.84 0.83 0.82 0.81 0.80 0.79 0.78 0.78 0.77 0.76 0.75
5.5 0.86 0.85 0.85 0.83 0.82 0.81 0.80 0.79 0.78 0.77 0.76
5.6 0.87 0.86 0.85 0.84 0.83 0.82 0.81 0.81 0.80 0.79 0.78
5.8 0.91 0.90 0.88 0.87 0.86 0.85 0.84 0.83 0.82 0.82 0.82
6.0 0.94 0.93 0.92 0.90 0.89 0.88 0.87 0.86 0.85 0.84 0.83
6.2 0.97 0.96 0.95 0.93 0.92 0.91 0.90 0.89 0.88 0.87 0.86
6.4 1.00 0.99 0.98 0.96 0.95 0.94 0.93 0.92 0.91 0.90 0.89
6.5 1.02 1.00 0.99 0.98 0.97 0.96 0.94 0.93 0.92 0.91 0.90
6.6 1.03 1.02 1.01 0.99 0.98 0.97 0.96 0.95 0.94 0.93 0.92
6.8 1.06 1.05 1.04 1.02 1.01 1.00 0.99 0.98 0.97 0.96 0.95
7.0 1.09 1.08 1.07 1.05 1.04 1.03 1.02 1.02 1.01 0.99 0.97
7.5 1.18 1.16 1.14 1.13 1.12 1.10 1.09 1.08 1.07 1.05 1.04