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Enhanced Production of Biofuel from Sugar

Industry Waste

By

Muzna Hashmi

PhD Thesis

Department of Microbiology

Faculty of Biological Sciences

Quaid-i-Azam University

Islamabad, Pakistan

2016

Enhanced Production of Biofuel from Sugar

Industry Waste

A thesis submitted in the partial fulfillment of the requirements for the

degree of

Docter of Philosophy

In

Microbiology

By

Muzna Hashmi

Department of Microbiology

Faculty of Biological Sciences

Quaid-i-Azam University, Islamabad, Pakistan

2016

Enhanced production of biofuel from sugar industry waste Page iii

DECLARATION

The material presented in this thesis is my original work and this work has

never been previously presented for any other degree.

Muzna Hashmi

Enhanced production of biofuel from sugar industry waste Page iv

CERTIFICATE

The Department of Microbiology, Quid-i-Azam University, Islamabad,

accepts this thesis by Muzna Hashmi in its present form as satisfying the

thesis requirement for degree of Doctor of Philosophy in Microbiology.

Supervisor: ________________________

Dr. Aamer Ali Shah

External Examiner: _____________________

_

External Examiner: ______________________

Chairperson: _______________________

Dr. Fariha Hasan

Dated:

Enhanced production of biofuel from sugar industry waste Page v

CONTENTS

Sr. No Titles Page. No.

i. List of Tables iv

ii. List of Figures viii

iii. List of Appendices xi

iv. List of Abbreviation xv

v. Acknowledgment xvi

vi. Abstract xviii

1. Introduction 1

2. Literature Review 9

3. Materials and Methods 40

4. Results 62

5. Discussion 113

6. Conclusions 131

7. Future Prospects 132

8. References 133

8. Appendices 160

Enhanced production of biofuel from sugar industry waste Page vi

List of Tables

No. Tables Page No

4.1. Physicochemical Properties of Molasses 64

4.2.a. Screening of isolated yeast strains against 10%

ethanol tolerance

65

4.2.b. Screening of isolated yeast strains against 15%

ethanol tolerance

66

4.3. Enhanced production of bioethanol by various

yeast strains using different concentration of

sugar

71

4.4. Enhancement of fermentation efficiency due to

optimization

82

4.5. The effect of specific gravity and feeding rate

of molasses on fermentation efficiency and

process completion time of fed batch

fermentation using Lalvin EC-1118

87

4.6. The effect of specific gravity and feeding rate

of molasses on fermentation efficiency and

process completion time of fed batch

fermentation using MZ-4

88

4.7. a. Severity factors for different conditions of

autohydrolysis

91

4.7.b. Severity factors for IL pretreatment conditions 91

4.8.a. Lignin Determination of Autohydrolyzed

Sugarcane bagasse

92

4.8.b. Lignin determination of IL pretreated sugarcane

bagasse

93

4.9.a. Carbohydrate determination of autohydrolyzed

sugarcane bagasse

95

Enhanced production of biofuel from sugar industry waste Page vii

4.9.b. Carbohydrate determination of IL pretreated

sugarcane bagasse

96

4.10. Assignments of FTIR-ATR absorption bands

for bagasse

96

4.11. Crystallinity measurements of autohydrolyzed

and IL pretreated bagasse

101

Enhanced production of biofuel from sugar industry waste Page viii

List of Figures

No Figures Page No.

2.1. Chemical composition of lignocellulosic biomass 23

2.2. Cellulose structure: Crystalline (red); Para-crystalline

(green) and amorphous (blue) chains

25

2.3. Separation of cellulose, hemicelluloses and lignin after

pretreatment of lignocellulosic biomass

35

2.4. Simplistic overview of some factors limiting efficient

hydrolysis of cellulose

37

4.1. Phylogenetic tree of isolated Saccharomyces cerevisiae

strain MZ-4

68

4.2. HPLC chromatogram for ethanol detection 72

4.3. Effect of pH on enhanced production of bioethanol by

Lalvin EC-1118 and MZ-4

74

4.4. Effect of temperature on enhanced production of

bioethanol by Lalvin EC-1118 and MZ-4

74

4.5. Effect of inoculum size on enhanced production of


76

4.6. Effect of inoculum age on enhanced production of


76

4.7. Effect of nitrogen source on enhanced production of


79

4.8. Effect of chelating agents on enhanced production of


81

4.9. The effect of specific gravity and feeding rate for

enhanced production of bioethanol during fed-batch

fermentation using strains Lalvin EC-1118 and MZ-4

86

4.10 Drying and milling of sugarcane bagasse through sieve

size 40 Mesh

90

4.11.a. Sugarcane bagasse autohydrolyzed at 190°C for 10 90

Enhanced production of biofuel from sugar industry waste Page ix

min; 205°C for 6 min and untreated bagasse control

4.11.b. Sugacane bagasse, IL pretreated at 110°C for 30 min;

water treated at 110°C for 30 min (control) and

untreated bagasse (control)

90

4.12. Chromatogram showing peaks of glucose, xylose,

arabinose and mannose standard

94

4.13.a. FTIR spectra of untreated, autohydrolysis and IL

pretreated bagasse from 1800-600 cm-1

region

98

4.13.b. FTIR spectra of untreated, autohydrolysis and IL

pretreated bagasse from 4000 to 1800 cm-1

region

98

4.14. XRD analysis of untreated, autohydrolyzed and IL

pretreated bagasse

100

4.15.a. Enzyme loading optimization for autohydrolyzed

samples

103

4.15.b. Enzyme loading optimization for IL pretreated samples 103

4.16. Chromatogram of sugarcane bagasse hydrolysate

obtained after enzymatic hydrolysis

104

4.17.a. Glucose concentration released from autohydrolyzed

samples during enzymatic hydrolysis

106

4.17.b. Xylose concentration released from autohydrolyzed


106

4.18.a. Cellulose digestibility from autohydrolyzed samples

during enzymatic hydrolysis

107

4.18.b. Xylan digestibility from autohydrolyzed samples


107

4.19.a Glucose concentration released from IL pretreated


109

4.19.b. Xylose concentration released from IL pretreated

samples during enzymatic

109

4.20.a. Cellulose digestibility from autohydrolyzed samples


110

Enhanced production of biofuel from sugar industry waste Page x

4.20.b. Xylan digestibility from autohydrolyzed samples during

enzymatic hydrolysis

110

4.21. Production of Bioethanol from untreated, autohydrolyzed

and IL pretreated sugarcane bagasse

112

Enhanced production of biofuel from sugar industry waste Page xi

List of Appendices

No Appendices Page. No.

A Composition of Wallsteiner (WLN) media 161

B Sequence for Strain MZ-4 162

C Standard calibration curves for quantification of sugars and

ethanol

163

C1 Calibration curve for reducing sugar analysis by DNS method 163

C2 Calibration curve for ethanol determination by HPLC method 164

C3 Calibration curve for glucose determination by HPLC method 164

C4 Calibration curve for xylose determination by HPLC method 165

C5 Calibration curve for mannose determination by HPLC method 165

C6 Calibration curve for arabinose determination by HPLC

method

165

D Bioethanol production from sugarcane molasses 166

D.1. Effect of pH on enhanced production of bioethanol by using


167

D.2. Effect of temperature on enhanced production of bioethanol by

using Lalvin EC-1118 and MZ-4

167

D.3. Effect of inoculum size on enhanced production of bioethanol

by using Lalvin EC-1118 and MZ-4

168

D.4. Effect of inoculum age on enhanced production of bioethanol


168

D.5. Effect of nitrogen source on enhanced production of

bioethanol by using Lalvin EC-1118 and MZ-4

169

D.6. Effect of chelating agents on enhanced production of


169

E Ethanol Production from sugarcane bagasse 171

E.1.1. Glucose concentration released from untreated and

autohydrolyzed samples during enzymatic hydrolysis

171

E.1.2. Cellulose digestibility of untreated and autohydrolyzed 171

Enhanced production of biofuel from sugar industry waste Page xii


E.1.3. Xylose concentration released from untreated and

autohydrolyzed samples during enzymatic hydrolysis

172

E.1.4. Xylan digestibility of untreated and autohydrolyzed samples


172

E.2.1 Glucose concentration released from untreated control, water

treated control and ionic liquid pretreated samples during


173

E.2.2. Cellulose digestibility of untreated control, water treated

control and ionic liquid pretreated samples during enzymatic

hydrolysis

173

E.2.3. Xylose concentration released from untreated control, water

treated control and ionic liquid pretreated samples during


174

E.2.4. Xylan digestibility of untreated control, water treated control

and ionic liquid pretreated samples during enzymatic

hydrolysis

174

E.3.1. Bioethanol production from untreated bagasse by using

various yeast strains

175

E.3.2. Bioethanol production from bagasse autohydrolyzed at 190°C

by using various yeast strains

175



175



176

E.3.5. Bioethanol production from IL pretreated bagasse at 110°C by

using various yeast strains

176

F. Statistical analysis for the production of bioethanol from

sugarcane molasses

177

F.1. Analysis of variance for the effect of pH on enhanced

production of bioethanol by using Lalvin EC-1118 and MZ-4

177

Enhanced production of biofuel from sugar industry waste Page xiii

F.2. Analysis of variance for the effect of temperature on enhanced

production of bioethanol by using Lalvin EC-1118 and MZ-4

177

F.3. Analysis of variance for the effect of inoculum size on

enhanced production of bioethanol by using Lalvin EC-1118

and MZ-4

178

F.4. Analysis of variance for the effect of inoculum age on


and MZ-4

178

F.5. Analysis of variance for the effect of nitrogen source on


179

F.6. Analysis of variance for the effect of nitrogen source on

enhanced production of bioethanol by using MZ-4

179

F.7. Analysis of variance for the effect of chelating agents on


181

F.8. Analysis of variance for the effect of chelating agents on

enhanced production of bioethanol by using MZ-4

182

F.9. Analysis of variance for the effect of fed batch fermentation on


183

F.10. Analysis of variance for the effect of fed batch fermentation on


183

G. Statistical analysis for the production of bioethanol from

sugarcane bagasse

184

G.1.1. Analysis of variance for the glucose concentration released

from untreated and pretreated bagasse samples during


184

G.1.2. Tukey multiple comparisons for the glucose concentration

released from untreated and pretreated bagasse samples during


185

G.2.1. Analysis of variance for the xylose concentration released

from untreated and pretreated bagasse samples during


186

Enhanced production of biofuel from sugar industry waste Page xiv

G.2.2. Tukey multiple comparisons for the xylose concentration

released from untreated and pretreated bagasse samples during


187

G.3.1. Analysis of variance for the production of bioethanol from

pretreated bagasse by using various yeast strains

188

G.3.2. Tukey multiple comparisons for the production of bioethanol

from pretreated bagasse by using various yeast strains

189

Enhanced production of biofuel from sugar industry waste Page xv

List of Abbreviations

Abbreviation Description

% Percentage

C Celcius

conc. Concentration

CuSO4 Copper sulfate

DNS Dinitrosalycilic acid

DAP Di-ammonium phosphate

DAP Dilute acid pretreatment

EDTA Ethylene diamine-tetraacitic acid

FPU Filter paper assay unit

FTIR Fluorescence transform infrared

spectroscopy

G Gram

h hour

H2SO4 Sulfuric acid

HCl Hydrochloric acid

I Intensity

IL Ionic liquid

K4Fe(CN)6 Potassium ferrocyanide

LAP Laboratory Analytical Protocols

min Minutes

ml milliliter

NA.K. Tartrate Sodium potassium tartarate

NaCl Sodium chloride

NaOH Sodium hydroxide

NCBI National center of biological information

nm Nanometer

NREL National renewable energy lab

OD Optical density

PCR Polymerase chain reaction

RI Refractive index

Rpm rotation per minute

rRNA Ribosome ribonucleic acid

s seconds

Sp. grv. Specific gravity

S.F. Severity factors

Temp. Temperature

WLN Wallerstien

XRD X-Ray Diffraction Crystallography

YPD Yeast extract-peptone-dextrose

β Beta

ul microliter

Enhanced production of biofuel from sugar industry waste Page xvi

Acknowledgement

This thesis is more than just a culmination of a long academic and personal journey, it's a

celebration. This was a long road. But it was also brightly marked with an array of

growing moments that punctuated periods of bleak self-doubt. Only by learning to

humble myself through education, recognizing my weaknesses and limitations, and most

importantly, developing into a person who could effectively seek help from others did I

find the means to push myself and my boundaries. And so, I would like to thank those

who helped me reach the end of this journey, who helped me find strength through their

support, and who guided me onto the brilliant trajectory that I find myself on today.

It gives me great pleasure to express my profound gratitude, sincere thanks and sense of

obligation to my major supervisor, Dr. Aamer ali shah, Associate Professor, Department

of Microbiology, Faculty of biological Sciences, Quaid-i-Azam University (QAU)

Islamabad for his guidance, suggestions, support and encouragement in the completion of

this thesis.

I wish to extend my greatest appreciation to Prof. Dr. Abdul Hameed, Professor,

Department of Microbiology, Faculty of biological Sciences, Quaid-i-Azam University

(QAU) Islamabad for his valuable suggestions and the amazing mentorship and

confidence, who have been integral to the conception, progress and completion of this

research. I am very grateful to Dr. Fariha Hasan, Chairperson Department of

Microbiology for providing all the existing research facilities of the department to

accomplish this work.

I wish to extend my greatest appreciation to Prof. Dr. Safia Ahmed, Prof. Dr. Aftab

Iqbal Shafi and Dr. Malik Badshah, for providing their valuable suggestions, guidance

and encouragement for the completion of this work. . I also want to thank all my teachers

Dr. Rani Faryal, Dr. Naeem Ali, Dr. M. Imran, Dr. Ishtiaq Ali, Dr. Asif Jamal, Dr.

Rubab, Dr. Javaid Dasti and Dr. Samiullah, for their supportive attitude throughout

my research.

I would also like to acknowledge Higher Education Commission of Pakistan and

International Research support Initiative Program (IRSIP) for providing me funds to

complete my research at University of Tennessee, Knoxville, USA, where I worked

under the guidance of Prof. Arthur J. Ragauskas, Professor/Governor’s Chair in

Biorefining, Oak Ridge National Laboratory, Department of Chemical and Biomolecular

Engineering, The University of Tennessee, Knoxville, USA.

Enhanced production of biofuel from sugar industry waste Page xvii

I would like to extend my thanks to Dr. Nicole Labbe’, and Dr. Jingming Tao, Center

of Renewable Carbon, University of Tennessee, Knoxville, USA to provide me analytical

tools and help me for paper write up.

Special thanks go to Director, Murree Brewery for providing me molasses and for all

the beneficial knowledge regarding my research work. I would also like to extend my

thanks to Muhammad Naeem and Ather Hashmi, Technical Experts, Kamstec

International, and Asgher Khan, scientist SB pharma, to provide me HPLC operational

training.

A great depth of loving thanks to my friends Mishal Subhan, Sadia Satti, Leena,

Maria, Sara Shahid, Zimbeel Firdous, Mehreen Zaka, Tehmeena, Sumia Sahar for

their joyous company, lots of love, prayers, encouragement and helping me throughout

my studies. I would like to thank my lab fellows Saima, Aisha siddique, Hira, Afshan

Hina, Maliha Ahmed, Ramla, Anum, Nida Kanwal Nazia, Fozia, Muhammad Rafiq,

Muhammad Irfan, Muttiullah Khatak, Haleem, Waseem, Sahibzada and all others

for their enjoyable company and encouragement during my work.

I would like to thanks my lab fellows Tais, Naijia, Qining, Thomas and Romina

Stoffel for all their cooperation and friendly behavior in University of Tennessee,

Knoxville, USA. A special thank goes to Dr. Tyrone wells for his help, suggestions and

encouragement during my research work and thesis completion.

A non-payable debt to my mother (Masarrat Hashmi), whose love, care and prayers

mean a lot to me. I am also been fortunate to have a great father (Prof. Tufail Hashmi)

who always encouraged me for higher studies and did all his best for my career. Sweet

thanks to my brother Usama, Brother in law (Muhyudin Hashmi), Sister in law (Uzma),

sister (Rashida) and her children (Sidra , Awab, Zuha and Taha) for their love, care,

encouragement, prayers, and patience in bearing me during the tough times of my study.

There are many others around my life who have been helpful to me in many different

ways. There is no way to list all these individuals. I will simply say thank you to my

department, my university and all my fellows around my life that made this work

possible.

God Bless Them ALL.

Muzna Hashmi

Enhanced production of biofuel from sugar industry waste Page xviii

Abstract

The continuous upturn in the cost of petroleum and increasing energy crises has directed

the world’s interest to focus on alternative renewable energy resources. Recently,

bioethanol is emerging as an alternative fuel to substitute gasoline, which is petroleum

derived source of conventional energy. A significant variety of feedstocks can be used for

the production of bioethanol; however, sugar industry waste is considered as the best

option to evade food vs. fuel debate. In this study, two industrial wastes i.e. sugarcane

molasses and bagasse were converted to bioethanol using different microbial strains and

pretreatment strategies. To improve bioethanol production, different yeast strains were

isolated from numerous sources, and MZ-4 labeled strain was selected on the basis of its

maximum ethanol tolerance i.e. 15% (v/v). MZ-4 strain was then identified as

Saccharomyces cerevisiae by 18SrRNA sequencing, and later compared with a

comparatively better commercially available strain Lalvin EC-1118 strain, which was

maximally tolerant to 18% (v/v) ethanol. The physicochemical parameters were

optimized for both strains independently. During batch fermentation by strain MZ-4, the

maximum ethanol yield was determined as 11.1% (v/v) with 69.3% fermentation

efficiency, when pH 5 was adjusted for molasses dilution containing 25% (w/v) sugar

concentration with 10% inoculum before incubation at 33°C for 72 h. However, Lalvin

EC-1118 strain showed comparatively less ethanol yield of 10.9% (v/v) with

fermentation efficiency of 68.1% under its optimal conditions i.e. pH 4.5; inoculum size

of 7.5% and incubation at 30°C for 72 h. Additionally, the study on effect of various

nitrogen sources showed that, MZ-4 produced more ethanol when 0.1% (w/v) NH4Cl was

added; whereas, Lalvin EC-1118 demonstrated better production after the addition of

0.1% (w/v) (NH4)2HPO4. Moreover, it was also observed that MZ-4 and Lalvin EC-1118

exhibited better yields when 0.01 and 0.04% (w/v) of K4Fe(CN)6 was used respectively,

as a chelating agent. During the fed batch fermentation, Lalvin EC-1118 produced a

greater ethanol yield of 13.9% with fermentation efficiency of 81.1%, when 1.090

specific gravity of molasses dilution was adjusted and fed after every 12 h. However, the

strain MZ-4 showed better fermentation efficiency of 83.2% with comparatively less

Enhanced production of biofuel from sugar industry waste Page xix

ethanol yield i.e. 13.5% (v/v) by using molasses dilution of same specific gravity and 24

h feeding interval.

Meanwhile, one of the main challenges for bioethanol production from lignocellulosic

material such as sugarcane bagasse is the recalcitrance of the biomass. A second study

evaluated the efficiency of an ionic liquid (IL) i.e. 1- butyl-3-methyl imidazolium acetate

([C4mim][OAc]) pretreatment at 110°C for 30 min, and compared it with high

temperature autohydrolysis pretreatment (i.e. 110°C for 30 min, 190°C for 10 min and

205°C for 6 min). It was found that sugarcane bagasse exhibited a considerable decrease

in lignin content, reduced cellulose crystallinity, and enhanced cellulose and xylan

digestibility, when subjected to IL pretreatment. Pretreated samples were also

characterized by Fourier transform infrared spectroscopy to verify these findings.

Altogether, cellulose and xylan digestibility of IL pretreated bagasse was determined as

97.4 and 98.6% after 72 h of enzymatic hydrolysis, respectively. In the case of

autohydrolysis, the maximum of cellulose and xylan digestibility was determined after 72

h as 62.1 and 5.7% from bagasse pretreated at 205°C for 6 min, respectively. X-ray

diffraction analysis also showed a significant reduction in crystallinity of IL pretreated

bagasse samples. During fermentation process, IL pretreated and autohydrolyzed bagasse

(205°C for 6 min) exhibited maximum ethanol production of 78.8 and 70.9 mg/g

substrate after 24 h of fermentation, respectively. Comparatively, the fermentation of

bagasse autohydrolyzed at 190°C for 10 min and 110°C for 30 min yielded maximum

ethanol of 66.0 and 28.4 mg/g substrate by using S. cerevisiae Lalvin EC-1118,

respectively. Thus it can be concluded that, fed batch fermentation is employed for the

maximum ethanol yield from sugarcane molasses using Lalvin EC-1118 strain, while IL

pretreated bagasse gives maximum yield when fermented with strain MZ-4.

PhD Thesis

Enhanced production of biofuel from sugar industry waste Page 1

Chapter 1

Introduction

PhD Thesis


Depletion of fossil fuel resources, limited global supply of oil, energy crises and

increasing CO2 emission has increased the worldwide interest to substitute fossil fuels

by some alternative fuel (Huang and Ragauskas, 2012). Today, the ecofriendly biofuel

utilization as a substituent for the petroleum based products, has attracted worldwide

interest for its production at large scale because it can be used in current unmodified

engines by blending it with fossil fuels in different proportions (Macedo, 1998;

Hansen et al., 2005). In order to create more sustainable and economically viable

system, it is more important to emphasize on cheaper ways to produce biofuel to

make it more favorable as compared to petroleum based products (Zabed et al., 2014).

Currently, most popular biofuels being used at different countries are bioethanol,

biodiesel and biogas. The production of biofuel mainly depends on the availability of

substrate and the ease of its formation. Biodiesel are more commonly produced in

Europe from oil containing seeds and plants. However, the production of biogas is

mainly being investigated in Sweden and Germany from the combination of cattle

manure and different agricultural feedstock (Held et al., 2008). In contrast, those

substrates which are rich in sugars are preferably converted in to bioethanol through a

simple process of fermentation (Balat and Balat, 2009). Ethanol can be used in

substituent to gasoline and provide an environmentally safe alternative to fossil fuels

(Macedo, 1998). Currently, ethanol producing industries are utilizing two main types

of feedstock that are sugars and starch containing crops (Wilkie et al., 2000; Mojović

et al., 2006; Balat and Balat, 2009). More than 60% of ethanol among the world is

being produced from sugar crops like sugarcane and rest of 40% is being produced

from starchy grains (Salassi, 2007).

Sugarcane has widely been recognized one of the main biofuel crops in last 10-15

years; because it only requires a simple process of fermentation for the production of

bioethanol from sugarcane juice and molasses (Hartemink, 2008). Sugarcane is

actually a crop of tropical area, which is being cultivated in more than 200 countries,

worldwide. Brazil, ranked second among world’s bioethanol producer, is mainly

utilizing sugarcane juice and molasses for the production of bioethanol. More than

40% of their fuel demands are met from bioethanol (Agama Energy., 2003).

Sugarcane contains 30% more sugars than corn that is widely being used in USA for

the production of bioethanol. Brazil and USA supplies more than 65% of world’s total

ethanol. In Brazil more than 20% of total vehicles have flex fuel engines that are able

PhD Thesis


to consume all proportions of ethanol as fuel. Other vehicles with conventional

engines, can use only ethanol-gasoline blend up to E15 i.e. 15% of ethanol with 85%

of gasoline (Rosillo-Calle and Cortez, 1998).

There are two main types of the wastes generated by sugar industry i.e. molasses and

bagasse. In sugar industry, sugarcane juice is squeezed out under high pressure with

the help of heavy rollers. The juice is clarified, then heated and centrifuged multiple

times to crystallize and separate sugar crystals. Molasses is the non-crystallizable

residues that remain after purification of sucrose from sugarcane juice. This is

moderately economical, rapidly accessible substrate, which is usually utilized as a

feedstock for production of bioethanol. A typical sugar cane molasses normally has

17–25% of water substance, 45–60% of sugar content (sucrose, glucose, and fructose)

and 2–5% of polysaccharides (dextrin, pentose and polyuronic acids). It also contains

the non-sucrose substances, incorporate inorganic salts, kestose, raffinose, natural

acids etc. (Najafpour and Poishan, 2003; da Silva et al., 2012; de Andrade et al.,

2013).

Sugarcane industry is the second largest industry of Pakistan after textile industry.

Pakistan yields 63 million metric tons of sugarcane per anum, and ranks fifth for its

production, worldwide. In Pakistan, there are 83 sugar mills, which annually generate

2.0 million MT molasses that is later converted in to bioethanol (PBS, 2013). High

cost ethanol production from sugarcane molasses can be mainly attributed to low

ethanol content in fermentation media which requires more energy consumption for

distillation process (Zabed et al., 2014). Therefore, efforts are made to enhance

ethanol concentration in fermentation broth to reduce distillation cost (Bai et al.,

2004). Currently Pakistan is producing only 0.13 million MT of ethanol per year

which can be increased to three times, if the fermentation efficiency reaches up to

90%. Fermentation efficiency of the process in most of the distilleries in Pakistan is

less than 50% as estimated by annual sugar report (PBS, 2013). The main hurdle in

increasing the ethanol yield and fermentation efficiency is the selection of most potent

microbial strain for the process of fermentation. Moreover, the final ethanol yield and

fermentation efficiency is also affected by operating the process under unfavorable

physicochemical parameters. Different physicochemical parameters like sugar

concentration, temperature, pH and nutrients are needed to be optimized to determine

PhD Thesis


the best conditions at which maximum yield can be obtained (Wyman and Hinman,

1990).

There are several microorganisms that have the ability to ferment sugars into ethanol.

The most commonly used microorganisms are yeast especially Saccharomyces

cerevisiae (Zhu et al., 2012). S. cerevisiae has the ability to utilize both monomeric

sugars and sucrose, which makes it an efficient microbe to be used in variety of

substrate (Badotti et al., 2008; Canilha et al., 2012). Other advantages related to its

use are its highest resistance against high ethanol concentration, inhibitor resistance

and its ability to consume significant amount of substrate. Unfortunately, S. cerevisiae

lacks genes which could make it able to assimilate xylose; however, to obtain optimal

ethanol yields from sugarcane bagasse, conversion of hemicellulose fraction is also

essential (Canilha et al., 2012). There are only a few species which are capable of

converting xylose ethanol such as Scheffersomyces stipitis (pichia stipites), Candida

guilliermondii, Candida shehatae and Pachysolen tannophilus that can help to

convert xylose, i.e. the second most abundant component of bagasse, into bioethanol

(du Preez et al., 1986; Canilha et al., 2012).

The dried fibrous residue that remains after extraction of juice is termed as bagasse. It

is considered in Brazil that 1 ton of sugarcane will generate 280 kg of bagasse. Total

sugarcane production in 2015 was reported as 1877 million metric tons, worldwide.

Almost 50% of the bagasse is usually burnt in distilleries for power generation and the

remaining is stockpiled. The excess of this industrial waste has raised world’s interest

on biorefinery concept, and now latest researches are being done to convert sugarcane

bagasse into bioethanol (Rabelo et al., 2011).

The major problem face by industry for the production of bioethanol from sugarcane

bagasse is its lignocellulosic structure. Lignocellulosic biomass is a suitable resource

for renewable energy in terms of sustainability and ease of fermentation of

enzymatically released sugars that can be converted into bioethanol to substitute for

gasoline (Li et al., 2010). This resource is mainly composed of cellulose (30–45%),

hemicelluloses (20–30%), and lignin (5–20%) (Vallejos et al., 2012). Cellulose chains

are held together by van der Waals interactions and hydrogen bonding which makes it

a highly crystalline material (Qiu and Aita, 2013). The xylan layer is the most

PhD Thesis


prominent hemicelluloses in grasses and hardwoods and forms covalent linkages to

lignin in the cell wall. Xylan also has non-covalent interactions with cellulose, which

are believed to play a role in preventing enzymatic degradation (Ioelovich and Morag,

2012; Qiu and Aita, 2013). Lignin is a complex and branched aromatic structure that

is associated with hemicellulose and contributes to the recalcitrance of biomass (Qiu

and Aita, 2013; Pu et al., 2015). There are several stages involved in conversion of

recalcitrant lignocellulosics into ethanol that include physicochemical pretreatment,

enzymatic hydrolysis, fermentation, ethanol separation and effluent treatment.

Pretreatment is an important step in the overall process and is believed to break down

some of the carbohydrate-lignin complexes (Qiu and Aita, 2013; Yáñez‐S et al.,

2013).

Enzymatic hydrolysis of pretreated biomass is one of the most promising means of

releasing simple sugars from biomass (Batalha et al., 2015). Typically, enzymatic

hydrolysis of un-pretreated biomass is reported to produce less than 20% sugar yield

of theoretical value (Qiu et al., 2012). Different pretreatment methods have been

developed to reduce recalcitrance of lignocellulosic biomass but there are many

drawbacks associated with these procedures. For example biological pretreatments

(i.e. lignin degrading fungi) often require large residence times (Levin et al., 2008;

Dias et al., 2010), mechanical methods such as various grinding and milling

techniques are not appropriate due to their high capital costs and intensive energy

requirements (Naimi et al., 2006). Furthermore, various physicochemical techniques

(e.g. liquid hot water, autohydrolysis, supercritical fluids, steam explosion, dilute

acid, alkali) require high temperature and pressure along with specialized equipment

(Qiu and Aita, 2013; Yu et al., 2013; Batalha et al., 2015). Another drawback

associated with these pretreatments is release of inhibitors, which affect enzymatic

hydrolysis and subsequent fermentation process (Hongdan et al., 2013; Batalha et al.,

2015). These problems highlight the need for a more rapid; environment friendly, cost

effective and efficient method for lignocellulosic biomass conversion.

Despite being energy intensive, autohydrolysis is recommended as environmentally

benign and clean process (Lei et al., 2013) which doesn’t require any catalyst or

corrosive compounds (Hongdan et al., 2013). Biomass and water are heated from

130-230ºC for different time periods (from few seconds to several hours) to carry out

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this pretreatment (Batalha et al., 2015). At high temperature (~200˚C) water has an

acidic pH and acquires catalytic properties which eliminates the requirement of

catalyst to disrupt biomass (Mosier et al., 2005). Auto-ionization of water and

ionization of acidic species (uronic acid and formic acid) at high temperature generate

hydronium ions that catalyze the series of reactions and cause reduction in degree of

polymerization (DP) of hemicelluloses and celluloses by hydrolysis of selective

glycosidic bonds (Lee et al., 2009; Batalha et al., 2015). During autohydrolysis, acetyl

groups are released from substituted xylan chains (along with other organic acids)

which act as catalysts to assist in acid-catalyzed hydrolysis of hemicellulose fraction

of lignocellulosic biomass (Huang and Ragauskas, 2012; Sun et al., 2014; Batalha et

al., 2015; Pol et al., 2015). The main compositional changes observed after

autohydrolysis are lignin transformations and depolymerization of hemicellulose and

celluloses into oligomers and monomers due to very high severity conditions (Batalha

et al., 2015). These compositional changes during autohydrolysis pretreatment create

more number of structural changes including increasing the reducing ends of plant

polysaccharides for efficient exoglucanase activity and hence increased cellulose

digestibility (Huang and Ragauskas, 2012; Hongdan et al., 2013; Batalha et al., 2015).

The significant increase in lignin content after autohydrolysis might be attributed to

the removal of significant amount of hemicellulose while retaining most of the lignin.

The pseudo-lignin can also be generated from carbohydrate without significant

contribution from lignin, especially under high severity pretreatment conditions

(Sannigrahi et al., 2011).

Ionic liquid pretreatment (IL) is another method of reducing the recalcitrance of

biomass that has recently drawn a great deal of attention because of the unique

physical and chemical properties of ILs that are a very stable class of organic salts

with potential application as ―green solvents‖ (Qiu et al., 2012). The main advantages

of using ILs are related to their non-explosive, non-toxic, environment friendly, low

volatility, good recyclability and general stability under severe reaction conditions (Li

et al., 2010; da Silva et al., 2011; Qiu et al., 2012). For biomass pretreatment, three of

the most cited ILs are imidazoliums i.e. [C4mim][Cl] (1-butyl-3-methylimidazolium

chloride), [C2mim][Cl] (1-ethyl-3-methylimidazolium chloride) and [C2mim][OAc]

(1-ethyl-3-methylimidazolium acetate). All these alkylimidazolium salts have been

reported as most effective agents for lignocellulosics dissolution (Karatzos et al.,

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2012). It has been reported that the acetate ion in ILs are less viscous and can function

as a weak base to remove lignin and de-acetylate biomass (Mäki-Arvela et al., 2010;

Karatzos et al., 2012). Studies on these three alkylimidazolium salts reveal that

shorter alkyl chain of [C2mim]+ imparts greater extent of saccharification with faster

dissolution. However, higher dissolution extent of [C2mim]+ does not benefit the

overall process of pretreatment since losses in [C4mim]+-treated biomass were much

less as compared to [C2mim]+ pretreatment process. In terms of hemicellulose

saccharification yield, [C4mim]+ ILs perform better as hemicellulose is preserved in

its polymeric form and recovered form after pretreatment (Karatzos et al., 2012). Due

to these reasons, [C4mim][OAc] (1-butyl-3-methyl imidazolium acetate) pretreatment

was selected for this study. Previously, [C4mim][OAc] pretreatment of sugarcane

bagasse was reported by Silveria et al., (2015), who studied the effect of

[C4mim][OAc] pretreatment in combination with ethanol and supercritical CO2 (at

110, 145 and 180ºC for 2 h); however, Aver et al., (2013) used only [C4mim][OAc]

for the pretreatment at 120ºC for 24 h. Moreover, none of those studies discussed the

effect of [C4mim][OAc] pretreatment on crystallinity of sugarcane bagasse and the

efficiency of fermenting microbes for the production of bioethanol from

[C4mim][OAc] pretreated bagasse. In this study, [C4mim][OAc] was used alone for

pretreatment at comparatively less severe conditions (i.e. 110ºC for 30 min); and

compared with high temperature autohydrolysis to investigate the changes it imparts

to structure and composition of sugarcane bagasse and its potential to produce

bioethanol from sugarcane bagasse. Moreover, the efficiency of various commercially

available yeast strains was compared with a newly isolated strain to determine a better

fermenting strain for enhanced bioethanol production.

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Aim and Objectives

The aim of this study was to enhance the production of bioethanol from the waste

generated by sugar industry i.e. sugarcane molasses and bagasse. This aim of the

study was achieved by formulating following objectives:

Isolation, screening and molecular characterization of indigenous yeast strain

Comparison of indigenous yeast strain with commercially available strain for enhanced

production of bioethanol from sugarcane molasses

Effect of optimized physicochemical parameters on fermentation efficiency and final

ethanol yield from sugarcane molasses

Enhanced production of bioethanol from sugarcane molasses by using fed batch

fermentation

Optimization of feeding rate and substrate concentration during fed batch fermentation

Effect of autohydrolysis and IL pretreatments under different severity conditions on

compositional changes of sugarcane bagasse

Effect of autohydrolysis and IL pretreatments on structural changes of sugarcane bagasse

Effect of autohydrolysis and IL pretreatments on crystallinity of sugarcane bagasse

Effect of autohydrolysis and IL pretreatments on enhanced glucose and xylose release

from sugarcane bagasse during enzymatic hydrolysis

Effect of autohydrolysis and IL pretreatments of sugarcane bagasse on enhanced cellulose

and xylan digestibility during enzymatic hydrolysis

Comparison of indigenous and commercially available yeast strains for enhanced

production of bioethanol from pretreated sugarcane bagass

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Chapter 2

Literature Review

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The world’s energy sector is mainly dependent on non-renewable petroleum products.

In last few decades, an increase in population also raised the energy demands. It has

been estimated that the energy requirement has been increased 17 folds in previous

century (Demirbas, 2007). Furthermore, the emission of greenhouse gases i.e. CO2,

CO, NO2 and SO2, resulted in an increase in air pollution and led to global climate

change (Fuglestvedt et al., 2000). Today, the ecofriendly biofuel utilization as a

substituent for the petroleum based products, has attracted worldwide interest for its

production at large scale because it can be used in current unmodified engines by

blending it with fossil fuels in different proportions (Macedo, 1998; Hansen et al.,

2005). In order to create more sustainable and economically viable system, it is more

important to emphasize on cheaper ways to produce biofuel to make it more favorable

as compared to petroleum based products (Zabed et al., 2014). Various efforts are

being done in this regard to search a renewable source of energy. Recently, biofuels

are considered as an efficient renewable alternative energy source that can easily be

produced by various biological sources i.e. animal, plants, microorganisms etc.

(Aristidou and Penttilä, 2000; Zaldivar et al., 2001).

2.1. Types of Biofuels

Biofuels can be produced in variety of forms to fulfill various energy requirements

(like petroleum products). Some important types of biofuels are:

2.1.1. Biodiesel

Biodiesel consist of short chain alkyl ester, which are formed by transesterification

reaction of vegetable or animal fats (Stevens and Verhé, 2004). Edible oils are usually

not used as fuel; however, the low quality oil is converted into biodiesel that is later

processed and separated from water to be used in engines. The biodiesel can be used

in pure form (B100), or it can be blended with conventional petroleum based biodiesel

to be used in engines (Tickell and Tickell, 2003).

2.1.2. Bioalcohols

Bioethanol is the most commonly used bioalcohols, which can be used in substituent

to gasoline (Huang and Ragauskas, 2012), while other less common bioalcohols are

biomethanol, biopropanol and biobutanol (Minteer et al., 2011). Different types of

bioalcohols are mainly produced by variety of microorganisms, during the process of

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fermentation (Wyman and Hinman, 1990). Sugar rich feedstock i.e. sugarcane juice,

fruits juices, molasses are widely used for the production of bioethanol by the process

of alcoholic fermentation. However, starch containing plants (e.g. cassava, sweet

potato, corn) are first subjected to react with amylase enzymes for the conversion of

starch in to simple monomeric sugars, which are subsequently fermented for the

formation of bioalcohols (Ziska et al., 2009). The production of bioethanol from

lignocellulosic wastes i.e. bagasse, miscanthus, pinus, wheat stalks etc. requires

various pretreatments (physical, chemical or biological). The pretreated biomass then

undergoes enzymatic hydrolysis (by cellulases and hemicellulases) to breakdown

complex fibers to release simple monomeric sugars (i.e. glucose and xylose), which

are later fermented to produce bioethanol (Ragauskas, 2014).

2.1.3. Biogas

For the production of biogas, various energy crops and biodegradable waste (like

manure) is fed in to biodigester, and anaerobic digestion is carried out by using the

consortium of various anaerobic microbes (e.g. acetogens and methanogens). Methane

gas is recovered at the end of reaction from biodigester and used as biofuel. However,

the solid byproduct recovered at the end of process can be used as fertilizers

(Sreekrishnan et al., 2004; Amon et al., 2007; Taherzadeh and Karimi, 2008).

2.1.4. Syngas

Syngas is produced by combination of three processes i.e. pyrolysis, combustion and

gasification. The pyrolysis converts the biofuel into carbon monoxide. Little oxygen

is provided to support combustion. The gasification converts further substrate into

carbon monoxide and hydrogen. The syngas is a better fuel than combustion of

original biofuel because of more content of energy present in syngas (Basu, 2010;

Göransson et al., 2011).

2.2. Generations of Biofuels

On the basis of type of substrate, processing technology and their level of

development; biofuels are categorized into various generations (Nigam and Singh,

2011).

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2.2.1. First Generation biofuels

First generation biofuels are usually produced from sugar containing crop (i.e.

sugarcane juice, sugarbeet, molasses); starch containing crops (i.e. cereals, grains);

vegetable oils, animal fats etc. (Naik et al., 2010; Havlík et al., 2011).

2.2.2. Second Generation Biofuel

Second generation biofuel uses non-food substrate to produce biofuel. The substrate

utilized for second generation are stalks of wheat, corn, wood, sugarcane bagasse or

energy crops (miscanthus or bagasse). The second generation biofuel avoids food vs.

fuel debate by utilizing nonfood crops for the production of bioethanol. It usually

utilizes lignocellulosic materials which are degraded by various pretreatments and

degrading enzymes to convert complex structure into simple monomeric sugars,

which can be subsequently converted into variety of biofuels. Many second

generation biofuel i.e. biohydrogen, biomethane, biodiesel are under investigation

(Naik et al., 2010; Sims et al., 2010; Havlík et al., 2011).

2.2.3. Third Generation Biofuel

Algae are considered as low input high output feedstock to generate third generation

biofuels. In comparison to land crop like soybean, it is able to produce 30 times more

energy per acre. The most promising advantage of algae biofuel is its

biodegradability, which makes it environmentally safe option. Second and third

generation biofuels are termed as advanced biofuels (Dragone et al., 2010; Maity et

al., 2014).

2.2.4. Fourth Generation Biofuel

The fourth generation biofuel is attempting to convert vegetable oil and biodiesel into

gasoline. Another famous company ―synthetic genomics‖ is trying to produce biofuels

directly from carbon dioxide. Some researchers are trying to produce those genetically

modified crops that may able to consume more amount of carbon dioxide than

released by the biofuels thus creating an idea of carbon negative fuel (Demirbas,

2009; Lü et al., 2011).

2.3. Bioethanol as Fuel

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Ethanol can be used for variety of purposes like solvent, paints, perfumes and

beverage; however, recent investigations have revealed its importance as a biofuel

that can be used in substituent to gasoline (Wyman and Hinman, 1990; Alfenore et

al., 2004). Ethanol used in vehicles can either be a pure form (E100), or it can be

blended with various proportion of gasoline. The commonly used ethanol blends are

E5, E10, E25 and E85, where the letter ―E‖ describes the percentage of ethanol with

in ethanol-gasoline blend e.g. E5 contains 5% ethanol with 95% of gasoline. The

modern vehicles can use the ethanol-gasoline blend up to E15; however, there are

specialized vehicles which can run on any type of the ethanol blend and are known as

―Flex-fuel vehicles‖ (Suarez-Bertoa et al., 2015).

2.3.1. Production of Bioethanol

Bioethanol can be produced by variety of methods that include synthetic method or

biological method. Nowadays, the ethanol that is utilized as a solvent (or non-

beverage purposes) is usually produced by acid-catalyzed hydration of a

petrochemical feedstock i.e. ethylene. The ethanol produced by this method is termed

as ―synthetic‖ (Gnansounou and Dauriat, 2005)

C2H4 + H2O CH3CH2OH

Ethylene Water Ethanol

Ethanol is produced by biological method (fermentation), when it has to be used in

beverages or biofuel. One of the most common yeast being used in this process is S.

cerevisiae. The overall process was explained by scientist Gay-Lussac, who formed

the basis to calculate fermentation efficiency.

C6H12O6 2C2H5OH + 2CO2

Glucose Ethanol Carbon dioxide

(1 Kg) (0.511 kg) (0.489 kg)

During this process 1 kg of sugar is converted into 0.51kg of ethanol and 0.49kg of

CO2 (Gnansounou and Dauriat, 2005).

2.3.2. Role of Substrate

The nature and type of the substrate has immense importance for the production of

bioethanol (Prescott et al., 2002). Substrates are mainly categorized into three types

i.e. sugar containing substrate, starch containing substrate, and lignocellulosic waste.

There are varieties of feedstock which are rich in sugars like fruit juices, sugarcane

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juice, sugarcane molasses etc., and they can easily be converted into bioethanol by the

process of fermentation (Nigam, 1999). Other important substrates used for this

purpose are sweet sorghum (Bulawayo et al., 1996), sugar beet and beet molasses (El-

Diwany et al., 1992; Agrawal and Kumar, 1998).

Some other easily fermentable substrates are cheese whey and milk; however, those

microbial strains can be used to utilize these substrates which have the ability to

hydrolyze lactose (Ghaly and Ben-Hassan, 1995; Silva et al., 1995). Starch containing

substrates can also be used for the production of bioethanol. Most common among

them is corn, which is utilized by USA (the top ethanol producer) for the production

of bioethanol. Other starch containing materials are sweet potato and wheat, which are

also reported for the production of bioethanol by using various microbial strains

(Lindeman and Rocchiccioli, 1979; Maisch et al., 1979; Sree et al., 2000).

Recently, many researchers are working on the production of bioethanol from

lignocellulosic wastes, which may include woods, grasses, agricultural feedstock etc.

(Taherzadeh and Karimi, 2008). Different enzyme companies are trying to enhance

the production of various hydrolyzing enzymes i.e. cellulase and xylanases by

designing genetically modified organisms. The reduction in enzyme cost will reduce

the cost of overall process (Kaar and Holtzapple, 2000; Sun and Cheng, 2002; Yu and

Zhang, 2004). Pineapple, Cocoa and sugarcane bagasse are being tried to use for the

production of bioethanol, however this is very expensive process to convert them into

bioethanol (Samah et al., 1992). Many researches are being done to make this process

economically feasible to commercialize.

2.4. Bioethanol Production from Sugarcane Molasses

Molasses is actually a thick, dark brown; honey like material that can be obtained

from the sugarcane juice, after the sugar has been crystallized. Different types of the

sugars are present in sugarcane molasses e.g. glucose, sucrose, fructose that constitute

about 45-60% of the total sugar. Sugarcane molasses also contains nitrogen which is

important for the generation of amino acid and proteins in fermenting microbes (W

Borzani et al., 1993; Walter Borzani, 2001). Different investigations are being done

for the production of bioethanol by using free and immobilized microbial cells (Gikas

and Livingston, 1997; Yamada et al., 2002). As molasses contains easily fermentable

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sugar; therefore, no specific pretreatment is required to convert it into ethanol. Brazil,

the second major ethanol producer among the world, utilizes sugarcane juice and

molasses for the production of bioethanol (Bose and Ghose, 1973; Morimura et al.,

1997; Agrawal and Kumar, 1998). Similarly molasses is also utilized in India and

Pakistan for same purpose (Sharma and Tauro, 1986; Bulawayo et al., 1996).

Moreover, in India and Pakistan, sugarcane molasses is very cheap and plenty;

therefore, they prefer to utilize this easily fermentable substrate for the production of

bioethanol (Sharma and Tauro, 1986).

2.4.1. Role of Microorganisms

An extensive research has been carried out on various types of microorganisms i.e.

bacteria, fungi or yeast, which have been involved in process of alcoholic

fermentation (Bajaj et al., 2001). Among various microbes, S. cerevisiae has been

considered as the most efficient strain for the production of bioethanol. Other

important yeast strains that have been used in industry for the production of

bioethanol are Shizosaccharomyces pombe; Saccharomyces uvarum;

Zygosaccharomyces spp; Saccharomyces ellipsoideus; and Kluyveromyces (Walker,

1998; Canilha et al., 2012). Among the bacteria, the most promising specie that has

been studied for the enhanced production of bioethanol is Zymomonas mobilis.

Skotnicki et al., (1981) studied the ethanol yield by using 11 different strains of Z.

mobilis, and reported that some of these bacterial strains were tolerant against high

sugar and ethanol concentration. Moreover, these strains also showed stability at high

temperature condition. Bertolini et al., (1991) isolated a strain of S. cerevisiae and

allowed it to grow at 48% sugar concentration, and reported the fermentation

efficiency of 89 to 92%. However, in a comparative study, Bansal and Sing (2003)

reported that S. cerevisiae exhibited better ethanol production from sugarcane

molasses as compared to Z. mobilis. The main reason for different level of production

by using various strains was difference in metabolic pathways acquired by these

strains. The enzymatic study of these microbial strains showed the presence of

specialized enzymes (i.e. invertase and zymase) in these strains, for the production of

bioethanol. Invertase is involved in conversion of sucrose in to reducing sugars which

is subsequently fermented to ethanol with the help of zymase enzymes. Moreover, it

was observed that each microbe performs its best at specific physicochemical

conditions. At extreme conditions, the enzyme activities are reduced which adversely

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affects the efficiency of microbial strain for the production of bioethanol

(Gnansounou and Dauriat, 2005).

Genetically Modified Organisms

Two of the main enzymes responsible for the production of bioethanol are pyruvate

decarboxylase (PDC) and alcoholic dehydrogenase (ADH). These enzymes are found

in Z. mobilis as well as S. cerevisiae; however, Z. mobilis showed more affinity

towards substrate and S. cerevisiae has been shown more tolerance against ethanol

(Gunasekaran and Raj, 1999; Matthew et al., 2005). Therefore, the Plant

Biotechnology Unit of the Corporación para Investigaciones Biológicas (CIB)

genetically modified S. cerevisiae by inserting two main genes from Z. mobilis i.e.

pdc and adhII. As a result, the engineered strains exhibited better ethanol yield as

compared to parental strain i.e. CBS8066, when glucose was used as carbon source

(Vásquez et al., 2007; Peña-Serna et al., 2011).

2.4.2. Ethanol Tolerance

One of the main hurdles faced by fermenting microbes is their intolerance against

high concentration of ethanol, which reduces the final ethanol concentration. High

ethanol content denatures proteins and necessary enzymes, thus hinders the process of

fermentation. It has been observed that baker’s yeast can’t tolerate the ethanol more

than 5-6% (v/v); however, 12-15% (v/v) ethanol production is common in wine

industries. It has been reported that those strains which are used in alcohol industries

are tolerant up to 18% (v/v) ethanol (Balat and Balat, 2009).

2.4.3. Physicochemical Pretreatments

The main problems faced by ethanol industry are the lower ethanol yield and

fermentation efficiency, which are usually attributed to low ethanol tolerance among

fermenting microbe. Second major reason of these problem is operating the process

under non-favorable physicochemical parameters i.e. sugar concentration, pH,

temperature, inoculum etc. (Wyman and Hinman, 1990).

(a) Effect of Sugar Concentration

In ethanol industries, increase in sugar concentration is one of the best way to enhance

the production of bioethanol; however, too high concentration inhibits metabolic

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pathway of fermenting microbe (Jones et al., 1994). It has been observed that the

increase in amount of sugar concentration creates high osmotic pressure that is

difficult to be tolerated by microorganisms, thus affects fermentation efficiency.

Bertolini et al., (1991) isolated various strains from Brazilian ethanol industries and

studied their osmotic tolerance. Some of the yeast strains were able to utilize 30%

sugar concentration and produced various amount of ethanol in fermentation media.

Borzani et al., (1993) studied the logarithmic relationship between initial sugar

concentration and fermentation time and found that microbes withstand increase in

sugar concentration up to certain limit; further increase in sugar concentration

adversely affects fermentation process. Sree et al., (2000) used various sugar

concentrations i.e. 150, 200 and 250 (gm/l) at 30°C and studied that the final ethanol

yield obtained by these concentration was 72.5, 93 and 83 (gm/l), respectively at 30ºC

after 48 h. Periyasamy et al., (2009) revealed that under optimized condition, S.

cerevisiae strain produced 6.7% (v/v) of ethanol when 30% (w/v) of sugar was

present in fermentation medium. In another study maximum ethanol production was

determined as 7.7% (v/v) from 16% (w/v) sugar containing fermentation medium

(Arshad et al., 2008).

(b) Effect of pH

In order to obtain high ethanol yield from fermentation medium, the adjustment of pH

to the optimal value is quite important. The pH range 4-5 is considered as the optimal

range for most of the fermenting yeasts. The pH adjustment is important to avoid

bacterial growth by providing acidic environment because Lactobacilli, the main

contaminants of fermentation media prefer to grow at pH 5.4 to 5.6; furthermore,

fermenting yeast showed better growth at slightly acidic pH (Mathewson, 1980). The

studies showed that the growth of contaminants in a medium produces undesirable

compounds in fermentation medium, which makes the environment unfavorable for

other microbes (Yadav et al., 1997; Periyasamy et al., 2009). The optimum pH of

different yeast varieties was studied by many researchers and all reported same range

for optimum pH i.e. 4-5, when yeast was used for alcoholic fermenting (Lin and

Tanaka, 2006; Mariam et al., 2009; Maharjan et al., 2012). It was studied by Wang et

al., (2001) that the contamination of acetic acid bacteria increases at pH above 7.

Moreover, it also affected aldehyde dehydrogenase activity, which stops alcoholic

fermentation and enhanced glycerol production.

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(c) Effect of Temperature

In order to obtain the desired amount of product, it is important to monitor the

temperature, as it is one of the important factors that alters the rate of process and

directly affects the final yield. During the process of ethanol production, heat is

evolved from the fermentation process which increases the temperature of reactor;

due to these reasons, the fermenter should be cooled down frequently to maintain the

temperature at the optimum level. The increase in temperature adversely affects the

viability of microbial cells and metabolic process. It has been investigated that high

temperature alters the fatty acid composition in yeast cell membrane (Ohta et al.,

1988). The change in phospholipid content in cell membrane affects membrane

fluidity and cellular activities (Banat et al., 1998). At higher temperature, decrease in

ethanol yield might be attributed to protein denaturation which hinders enzyme’s

catalytic activity and cause death of yeast cells (Dhaliwal et al., 2011).

(d) Nutrient Requirement

It was previously believed that the ethanol tolerance is not affected by nutritional

requirement; however, with the advent of research this concept has been changed

(Casey et al., 1983). Now, the studies have been shown that the addition of

nitrogenous source like urea in fermentation medium not only enhances ethanol

tolerance, but also improves sugar utilization competences of fermenting microbes.

For better fermentation, urea was commonly added in fermentation media as nitrogen,

whereas DAP (Diammonium hydrogen phosphate) as phosphorus plus nitrogen

source. Nitrogen is important for amino acid synthesis, while phosphate has major

role in glycolytic pathway during fermentation and also involves in nucleic acid

synthesis, thus plays vital role in yeast replication (Mukhtar et al., 2010). Nofemele et

al., (2012) reported in his studies that 2g/l of urea addition has been shown to enhance

the ethanol concentration up to maximum level when fermentation was carried out at

35ºC. Other researchers has been shown the similar effect that the addition of urea and

DAP both played important role to enhance ethanol yield. Mukhtar et al., (2010)

studied effect of nitrogen on a commercial yeast i.e. Saf-instant and determined

similar increase in ethanol yield, when either urea or DAP was added. Maharjan et al.,

(2012) evaluated the effect of various nitrogen sources such as urea, ammonium

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sulfate, di-ammonium phosphate, ammonium chloride and ammonium nitrate to select

the best option for enhanced ethanol production by using yeast strain S2Y8.

(e) Effect of Chelating Agents

Molasses contains many metals and heavy metals like iron (Fe), aluminum (Al),

copper (Cu) etc., which can hinder the activity of fermenting microbes and reduces

final yield. Many scientists have studied the effect of various chelating agents to

remove the metals from fermentation media. Lee et al., (2012) studied the effect of

EDTA (ethylene diamine tetra acetic acid) and NTA (nitrile tri-acetic acid) for

adsorption of metal ions present in media and also studied the effect on enhanced

ethanol production by using L. japonica as fermenting strain. Benerji et al., (2010)

used EDTA and sodium potassium tartrate in different concentrations to study their

effects on ethanol production from muhua flower. Pandey et. al., (1993) studied the

concentration ranges from 50 mg/L to 2000 mg/L of various chelating agents i.e.

EDTA, potassium ferrocyanide (K4Fe(CN)6,) and sodium potassium tartrate on

production of ethanol from sugarcane molasses.

(f) Effect of Inoculum

In general large amount of cells at their exponential phase are required to make the

process of fermentation successful. Many scientists have previously studied the effect

of inoculum on enhanced production of bioethanol. Munene et al., (2002) reported

that the inoculum size of 7×106 viable count/ml yielded maximum ethanol with

minimum byproducts i.e. glycerol. Laopaiboon et al., (2007) reported that 1x108

cells/ml was the optimized inoculum size for maximum ethanol production. Later,

Perisyasmi et al., (2009) during his study on S. cerevisiae revealed that 2 g of yeast

inoculum exhibited maximum production of bioethanol. Benerji et al., (2010)

determined that 1.5% (v/v) of 48 h old inoculum was best for the maximum

production of bioethanol.

2.5. Types of Fermentation

2.5.1. Batch Fermentation

During batch fermentation, the fermenter is filled with substrate, then pH and

temperature of the system is adjusted according to the optimized conditions.

Moreover, nutrient supplements are added to meet the growth requirement of

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fermenting microbes. The substrate is steam sterilized before the addition of inoculum

in fermenter. The process of fermentation is completed after certain time and the

entire fermentation medium is removed for the product recovery. Melle-Boinot

process is also a type of batch fermentation, during which the substrate is first

sterilized and pH is adjusted by the addition of H2SO4. The dissolved solid content of

fermentation broth is adjusted between 14-22°brix. After completion of fermentation

process, the medium is centrifuged to separate ethanol and yeast. Yeast cells are then

recycled to the same fermenter for next cycle in order to obtain maximum

fermentation efficiency by maintaining elevated cell concentration (Kosaric and

Velikonja, 1995). One of the major limitations associated with batch fermentation is

that it requires higher concentration of sugar for higher ethanol yield; however, high

sugar concentration inhibits the process due to osmotic intolerance of most of the

yeast strains (Grubb and Mawson, 1993). Moreover, there is also accumulation of

ethanol at the end of process that adversely affects the growth of yeast, thus hinders

the process for further ethanol production (Lynd et al., 1991).

2.5.2. Fed-Batch Fermentation

Due to osmotic intolerance of fermenting strains, fed batch fermentation was

introduced. Fed batch fermentation is a semi-batch fermentation, in which substrate

and necessary nutrients are added either continuously or intermittently. The product is

recovered at the end of the fermentation process, either fully or partially. This process

can be repeated several times if the microbial cells are fully viable. During fed batch

fermentation the volume of fermentation medium increases during the course of

reaction. Fed batch fermentation avoids the problem of high sugar intolerance of

fermenting strain because it allows the stepwise addition of substrate in fermenter

(Yamanè et al., 1984). By manipulating the feeding rate, the nutrient addition can be

manipulated to remain constant or increased at predetermined optimal rate. This type

of fermentation is usually used in Brazil for the production of high concentration of

ethanol from sugarcane molasses (Minihane and Brown, 1986).

2.5.3. Continuos Fermentation

Continuous fermentation is the process that makes it possible to produce ethanol

continuously for unlimited period of time (Klapatch et al., 1994). The substrate is

continuously added to the fermenter and ethanol is continuously removed, which

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removes the problems created in fermentation medium because of high concentration

of substrate or product. The number of microbial cells is also adjusted to the constant

number in fermenter by continuous removal of worn out cells (Hack et al., 1994;

Banat et al., 1998).

The major limitation of the continuous fermentation for the ethanol production is the

supply of oxygen. A continuous supply of oxygen is required for the growth of cell

and biomass generation; however, the process of fermentation requires anaerobic

conditions. During ethanol production, lower availability of oxygen makes it difficult

to generate energy cells to replace the worn out old cells. Further research is still

required to overcome these limitations (Banat et al., 1998).

Continuous fermenter mainly consists of series of tanks: the first tank ―wort receiver‖

dilutes the wort to adjust specific gravity of fermentation medium, which opens into a

hold up vessel. The holdup vessel mixes new wort with yeast and the recycled wort

coming from the first fermenter. The holdup fermenter is followed by first fermenter

(residence time 30 h), second fermenter (residence times 12 h) for final tuning, and

yeast separator. The yeast separator separates the yeast from this system by

centrifugation (Boulton and Quain 2001).

2.6. Bioethanol Production from Sugarcane Bagasse

Sugarcane bagasse is one of the major byproducts of sugar industry, which is

lignocellulosic in nature and mainly consists of cellulose, hemicellulose and lignin.

This industrial waste is broken down to sugars (i.e. glucose and xylose) by various

pretreatment strategies, which is subsequently converted to various types of fuels i.e.

bioethanol, biogas, biobutanol etc. (Maitan-Alfenas et al., 2015)

2.6.1. Chemical Composition of Sugar cane Bagasse

Lignocellulosic material mainly consist of three types of polymers i.e. cellulose (30-

50%), hemicellulose (15-30%) and lignin (10-25%); however, the composition of

these three constituents vary with the type of plants (Monlau et al., 2013). In addition,

small amount of pectin, proteins and non-structural carbohydrates (i.e. sucrose,

glucose and fructose) are also present in lignocellulosic biomass (Jørgensen et al.,

2007).

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Fig. 2.1. Chemical composition of lignocellulosic biomass (Scheller and Ulvskov,

2010)

(a) Cellulose

Cellulose is considered as the main constituent of cell wall. It is linear polysaccharide

polymer of glucose that is linked together by β-(1→4) glycosidic bonds (Fengel and

Wegener, 1984; Fengel, 1992). The nature of these bindings allows the cellulose

polymer to arrange in linear chains. The chemical formula of cellulose is represented

as (C6H10O5)n, and the different chemical properties of cellulose depends on its degree

of polymerization that ranges from 500 to 15000 (Holtzapple et al., 1990). The inter

and intra-molecular hydrogen bonding helps in formation of various parallel chains

that are coalesced to form micro-fibrils, which are further united to constitute a fiber

(Faulon et al., 1994; Chandra et al., 2012). This highly organized structure makes

cellulose surface highly hydrophobic and tensile, which is resistant to organic solvents

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and enzymatic hydrolysis (Ward et al., 1989). The hydrophobicity of cellulose

molecule makes a thick layer of water that makes the diffusion of enzymes more

difficult (Matthews et al., 2006).

In plant cell walls, cellulose can be found as either crystalline, amorphous or both

forms. The intra and inter-molecular hydrogen bonding with in cellulose structure

constitutes 36 chains that aggregate to form crystalline structure (Matthews et al.,

2006). It is suggested that 36 glucan chains constitute elementary fiber of cellulose.

The most inner six chains are truly crystalline that are surrounded by 12 para-

crystalline chains. However, the outer shell of the cellulose fiber contains 18 chains

which are amorphous in nature (Ragauskas, 2014).

It is considered that, the enzymes can easily access amorphous regions of cellulose

and hydrolyze it to release glucose. However, the crystallinity of cellulose makes it

difficult to be degraded by enzymatic activity as the contact efficiency of crystalline

cellulose is decreased (Chang and Holtzapple, 2000). During autohydrolysis and

dilute acid pretreatment (DAP), it has been determined that the increase in the

crystallinity of cellulose had adverse effect on efficiency of pretreatment. Thusly, it is

considered that the strategies to remove crystallinity can enhance the digestibility

(Han et al., 1983). Other studies reported that, crystallinity had no effect on

digestibility; rather the increase in digestibility was attributed to increase in pore size,

reduction in degree of polymerization (DP) and particle size (Puri, 1984; Sinitsyn et

al., 1991). All these factors are interlinked that makes it difficult to analyze only one

factor separately. However, due to heterogeneous nature of biomass, crystallinity can

only be considered as one of the important factor that affects digestibility (Taherzadeh

and Karimi, 2008).

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Fig.2.2. Cellulose structure: Crystalline (red); Para-crystalline (green) and amorphous

(blue) chains (http://www.uky.edu/~dhild/biochem/11B/lect11B.html)

(b) Hemicelluloses

―Hemicelluloses‖ is a collective term used to represent various polysaccharides

present in cell wall of plants. These are considered as highly branched structures and

are associated with celluloses. Contrary to cellulose, hemicelluloses have low degree

of polymerization (less than 200) and are mostly amorphous in nature. Hemicelluloses

are mainly composed of pentoses (xylose and arabinose) and hexoses (mannose,

glucose and galactose). In hardwoods and agricultural residues, xylose (C5 sugar) is

the most abundant reducing sugar among hemicelluloses. Xylan has backbone of β-

(1,4)-linked xylosyl residues which is acetylated (Kuhad et al., 1997). Fengel and

Wegener (1984) reported that, the quantity of acetic acid was more in hardwood

feedstock as compared to softwood feed stock. In heteroxylans, residues of xylose are

replaced by other components, and are mostly reported in variety of plants. Grasses

are mainly composed of glucuronoarabinoxylans, which contains glucuronic acid and

arabinose associated with xylan (Carpita et al., 2001; Saha, 2003). Most of the sugar

component of hemicelluloses involves in formation of covalent linkages between

lignin and carbohydrate resulting in formation of lignin-carbohydrate complex (LCC).

Benzyl ester, benzyl ether and glycosidic linkages are the most common LCC

linkages reported in various biomasses. The benzyl ester linkage can be hydrolyzed

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by alkaline treatment; however, the other two remains stable during alkaline treatment

(Ragauskas, 2014).

(c) Lignin

After cellulose and hemicelluloses, lignin is the third most abundant constituent of

plant cell wall, which imparts resistance (against microbes), stability and

impermeability to the cell wall structure. Lignin-carbohydrate complex actually

imparts structural integrity, rigidity and prevent the swelling of biomass. Lignin

adopts various ways like sterical hindered of enzymes and fiber swelling, to make the

digestion of biomass difficult (Mooney et al., 1998). It is a highly branched

amorphous structure consists of coniferyl (guaiacyl propanol), coumaryl (p-

hydroxyphenyl propanol) and sinapyl (syringyl alcohol), which are phenyl propanoic

alcohols with different substituents (Robert, 2003). The abundance of the polyphenyl

propanoic acid depends upon type of specie, maturity and the locality of lignin with in

cell wall. Usually, low lignin content was reported in grasses; however, hardwood has

comparatively higher lignin content (Monlau et al., 2013). Previously, three main

groups of lignin was reported i.e. the lignin from softwood contains guaiacyl units, the

hardwood with guaiacyl and syringyl units and herbaceous plants have all of the

above three units present in varying fractions (Boerjan et al., 2003; Vanholme et al.,

2010). The greater amount of guaiacyl subunits in softwood makes it more

recalcitrant than hardwood which is mainly composed of guaiacyl and syringyl

subunits (Ramos et al., 1992).

It has been reported that lignin was less hydrophilic as compared to celluloses and

hemicelluloses that makes the water absorption and fiber swelling quite difficult

(Fengel and Wegener, 1984; Grabber, 2005; Akin, 2008). The water around 180°C

starts dissolving lignin under neutral conditions; however, the solubility of lignin

under acidic, basic or neutral conditions depends on precursors of lignin i.e. sinapyl,

coniferyl or courmaryl alcohol (Kubikova et al., 1996; Grabber, 2005). Alcohol,

dioxane, acetone, pyridine and dimethyl sulfoxide are usually used to dissolve the

lignin (Ragauskas, 2014). It is also believed that dissolved lignin inhibit cellulases and

xylanases that makes the digestion more complicated (Berlin et al., 2007).

2.7. Pretreatment Strategies

2.7.1. Physical Pretreatments

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(a) Milling

During the process of milling, the biomass is grounded to the final particle size up to

0.2 to 2 mm. The reduction in size makes the substrate more accessible to enzymes by

degrading lignin-carbohydrate complex and also plays an important role in reduction

of crystallinity (Mais et al., 2002). There are different types of millings i.e. ball

milling, hammer milling, two-roll milling, disk milling and colloid milling. The main

bottleneck of this process is the high energy consumption, which can be overcome by

wet disk milling. Da Silva et al., 2010 treated sugarcane bagasse to compare the

effectiveness of ball milling and wet disk milling, and reported 78.7% and 49.3%

glucose yield, respectively.

(b) Irradiation

The ultimate goal of this pretreatment is to enhance the enzymatic hydrolysis. The

irradiations can be either used alone or in combination with other pretreatments. The

irradiations directly affect cellulose component and breakdown the glycosidic bonds

thus generated delicate fibers and oligosaccharides. More specific microwave

irradiation disrupts cellulose by molecular collision and dielectric polarization

(Gabhane et al., 2011). Imai et al., (2004) studied the effect of irradiation on

carboxymethyl cellulose (CMC), and reported the increase in enzymatic hydrolysis of

cellulose by 200%. Kaumakura et al., (1983) reported the doubling of enzymatic

saccharification when bagasse was irradiated with acid pretreatment. Intanakul et al.,

(2003) irradiated the sugarcane bagasse by using water/glycerine as the medium of

action and recovered 50% of carbohydrate as reducing sugar. The major advantage of

this process is short processing time and selectivity; however, the major drawback is

its high cost and difficulty to operate at industrial level (Cheng et al., 2011).

(c) Pyrolysis

During this process, lignocellulosic biomass is heated to temperature above 300°C,

which cause rapid degradation of cellulose into gaseous component and residual char.

At lower temperatures, more volatile compounds are formed because of the low

reaction speed. Fan et al., (1987) reported the digestibility of 85% of cellulose by acid

hydrolysis after mild condition pyrolysis of bagasse. The product of pyrolysis is

directly used as fuel rather than its conversion into bioethanol or biogas. Garc a-

Pèreza, et al., (2002) carried out vacuumed pyrolysis of sugarcane bagasse and

http://www.sciencedirect.com/science/article/pii/S016523700100184X

http://www.sciencedirect.com/science/article/pii/S016523700100184X

http://www.sciencedirect.com/science/article/pii/S016523700100184X#AFF1

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reported 34.4% oil yield. The bio-oils obtained after vacuumed pyrolysis had low ash

contents and less viscosity, which makes it a potential valuable liquid fuel.

(d) Freeze Pretreatment

This pretreatment has achieved great attention because of its high effectiveness and

environment friendly features. The study of this pretreatment on rice straw has shown

the increase in enzymatic digestibility. A very few studies has been carried out on this

strategy. One of the major bottlenecks of this process is its high energy requirements

(Chang et al., 2011).

2.7.2. Chemical pretreatment

(a) Acidic Pretreatment

Acid has been widely used to convert complex lignocellulosic biomass into easily

degradable structure. Among all acids, most of the studies were carried on sulfuric

acid; however, other acids i.e. nitric acid, hydrochloric acid and phosphoric acid have

also shown positive effect on breakdown of complex structure (Panagiotopoulos et

al., 2012).

The pretreatment of biomass with sulfuric acid converts hemicelluloses into simple

monomeric sugars, thus enhances the accessibility and digestibility of cellulose.

Acidic pretreatment can be carried out at either low acidic condition with high

temperature, or high acidic condition with mild temperature (Taherzadeh and Karimi,

2008). The high concentration acidic pretreatment is an economic process; however,

corrosiveness of equipment, toxicity, acid recovery and degradation of glucose into

furan type inhibitors i.e. HMF (5-hydroxy methyl- furfural) and 2-furfuralaldehyde

makes this process inapplicable (Sun and Cheng, 2002; Almeida et al., 2007; Gírio et

al., 2010; Pedersen et al., 2010). In comparison, dilute acid pretreatment (DAP)

favors over concentrated pretreatment because of ease in its operation. DAP can be

carried out either for short retention time (1-5 min) at higher temperature, or long

retention time (30-90 min) at low temperature condition (Cruz et al., 2000). Moutta et

al., (2012) treated sugarcane bagasse with dilute H2SO4 and reported 90% removal of

hemicelluloses. Moreover, the glucose yield was also enhanced up to 65%. The major

drawback of this pretreatment is the formation of various types of inhibitors i.e. furan,

carboxylic acid and phenolic compounds. At higher temperature, glucose is degraded

into hydroxymethylfurfural (HMF); whereas xylose is degraded into furfural which is

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further degraded into levulinic acid and formic acid, respectively. All these inhibitors

may affect subsequent downstream phases that include enzymatic hydrolysis and

fermentation (Palmqvist and Hahn-Hägerdal, 2000). The pH of the process should be

monitored appropriately to reduce the formation of inhibitors. Another drawback

associated with the acidic pretreatment is its high cost which is mainly attributed to

neutralization of the process media. The neutralization of the media is important to

carryout subsequent enzymatic hydrolysis and fermentation (Taherzadeh and Karimi,

2008).

(b) Alkaline Pretreatment

For alkaline pretreatment of biomass, various studies on effect of sodium hydroxide

(NaOH), potassium hydroxide (KOH), calcium hydroxide (CaOH2), and ammonia

(NH3) pretreatment have been investigated (Wan et al., 2011). This method is

believed to increase the porosity of biomass by saponification and salvation reaction

involved in disrupting the linkages between hemicelluloses and other components of

biomass and induces the swelling of celluloses (Sun and Cheng, 2002). It has also

been studied that the ester bond linkages between lignin and xylan layers are

disrupted during alkaline pretreatment, thus helps in delignification of lignocellulosic

biomass. Zhao et al., (2010) pretreated sugarcane bagasse with 10% NaOH and

reported 96% of delignification and fiber swelling after pretreatment. The major

advantages associated with alkaline pretreatment are removal of acetyl group, lignin

and different uronic acid substitutions, which hinders the accessibility of cellulose to

hydrolytic enzymes (Li et al., 2010). However, the effect of this pretreatment on

solubilization of cellulose and hemicellulose is much weaker as compared to acid

pretreatment (Carvalheiro et al., 2008). Other advantages related to alkaline

pretreatment are requirement of low temperature to operate this process. Furthermore,

this pretreatment doesn’t require specialized corrosion resistant reaction equipment

(Digman et al., 2007). The major drawback associated with this pretreatment is

prolonged residence time from few hours to many days, and also the need of

neutralization for the subsequent enzymatic hydrolysis and fermentation process

(Wan et al., 2011).

(c) Organosolvent pretreatment

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In organosolvent pretreatment process; ethanol, methanol, acetone, ethylene glycol

and tetrahydrofuran alcohol is used either in presence or absence of catalyst (Mesa et

al., 2011). Different organic or inorganic acids (HCl or H2SO4) or bases (NaOH, NH3

or CaOH) are used as catalyst (Zhao et al., 2009). This pretreatment is specialized for

the biomass having higher lignin content and involved in disruption of bonds between

lignin and hemicelluloses. This pretreatment can lead to the recovery of pure lignin as

a byproduct which can be considered as an extra advantage of this pretreatment (Mesa

et al., 2011). Mesa et al., (2011) reported that pretreatment of bagasse with 30% (v/v)

ethanol at 195°C for 60 min yielded 29.1% glucose during subsequent enzymatic

hydrolysis. The removal of lignin also helps to greatly enhance the surface area for

enzyme accessibility and cellulose digestion (Koo et al., 2011). The major drawback

of this process is use of highly flammable and volatile solvents which can arise the

problem of pressure adjustment and also the recyclability of these solvent to make the

process economically feasible (Sun and Cheng, 2002) and also to make the process

feasible for subsequent enzymatic hydrolysis and fermentation process. Ethanol and

methanol are usually preferred because of their low boiling point and ease of

recyclability.

(d) Ozonolysis

In this process, ozone gas is used to disrupt lignin and hemicelluloses, which in turn

enhances the cellulose digestibility. Ozone acts as strong oxidant that degrades the

lignin by direct ring cleavage. The important features of ozone, like its solubility in

water and its behavior as a strong oxidant, make it an interesting option to carry out

lignocellulosic breakdown (Balat, 2011). It mainly degrades lignin and releases small

molecular weight components like acetic and formic acids (Williams, 2006). The

main advantages of this process is the activity of ozone in ambient temperature, and

little release of degradation byproducts that can act as inhibitors of downstream

phases (García-Cubero et al., 2009). Travaini et al., (2013) studied the effect of

ozonolysis pretreatment on sugarcane bagasse and found increase in glucose and

xylose yield up to 41.7 and 52.4%, respectively. Only xylitol, acetic, formic and lactic

acid degradation compounds were found, with no detection of HMF (5-

hydroxymethylfurfural) or furfural. The main disadvantages associated with ozone are

its high cost and its high quantity requirement to treat biomass (Sun and Cheng,

2002).

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(e) Ionic Liquid Pretreatment

Ionic liquid pretreatment (IL) is another method of reducing the recalcitrance of

biomass that has recently drawn a great deal of attention because of the unique

physical and chemical properties of ILs that are a very stable class of organic salts

with potential application as ―green solvents‖ (Qiu et al., 2012). The main advantages

of using ILs are related to their non-explosive, non-toxic, environment friendly, low

volatility, good recyclability and general stability under severe reaction conditions (Li

et al., 2010; da Silva et al., 2011; Qiu et al., 2012). IL pretreatment is considered as

an effective pretreatment method as it weakens van der Waals interactions between

cell wall components (Li et al., 2010; Bian et al., 2014). In grasses, the ester linkages

that are formed between lignin and arabinoxylan are disrupted during IL pretreatment

(Li et al., 2010). It is expected that IL pretreatment imparts compositional changes

and interact with the original biomass by hydrogen, ionic and Π-Π interaction in order

to dissolve its components (Karatzos et al., 2012). Anionic moieties of ILs act as

hydrogen ion acceptor and interact with hydroxyl groups present within hydrogen

bonding network of cellulose; however, cations interact with lignin though Π-Π

interaction (Qiu and Aita, 2013; Ninomiya et al., 2015). IL pretreatment causes

dissolution of biomass that can be rapidly precipitated with an anti-solvent and this

prevents reconstruction of crystalline phase of cellulose resulting in the formation of

porous and amorphous cellulose. These effects increase surface area availability for

cellulases adsorption and also increase cellulose digestion (Qiang Li et al., 2009; Qiu

et al., 2012; Bian et al., 2014). For biomass pretreatment, three of the most cited ILs

are imidazoliums i.e. [C4mim][Cl] (1-butyl-3-methylimidazolium chloride),

[C2mim][Cl] (1- ethyl-3-methylimidazolium chloride) and [C2mim][OAc] (1-ethyl-3-

methylimidazolium acetate). All these alkylimidazolium salts have been reported as

most effective agents for lignocellulosics dissolution (Karatzos et al., 2012).

2.7.3. Physicochemical pretreatment

(a) Autohydrolysis

Autohydrolysis is recommended as environmentally benign and clean process (Lei et

al., 2013) which doesn’t require any catalyst or corrosive compounds (Hongdan et al.,

2013). Biomass and water are heated from 130-230ºC for different time periods

(from few seconds to several hours) to carry out this pretreatment (Batalha et al.,

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2015). At high temperature (~200˚C) water has an acidic pH and acquires catalytic

properties which eliminates the requirement of catalyst to disrupt biomass (Mosier et

al., 2005). Auto-ionization of water and ionization of acidic species (uronic acid and

formic acid) at high temperature generate hydronium ions that catalyze the series of

reactions and cause reduction in degree of polymerization (DP) of hemicelluloses and

celluloses by hydrolysis of selective glycosidic bonds (Lee et al., 2009; Batalha et al.,

2015). During autohydrolysis, acetyl groups are released from substituted xylan

chains (along with other organic acids) which act as catalysts to assist in acid-

catalyzed hydrolysis of hemicellulose fraction of lignocellulosic biomass (Huang and

Ragauskas, 2012; Sun et al., 2014; Batalha et al., 2015; Pol et al., 2015). The main

compositional changes observed after autohydrolysis are lignin transformations and

depolymerization of hemicellulose and celluloses into oligomers and monomers due

to very high severity conditions (Batalha et al., 2015). These compositional changes

during autohydrolysis pretreatment create more number of structural changes

including increasing the reducing ends of plant polysaccharides for efficient

exoglucanase activity and hence increased cellulose digestibility (Huang and

Ragauskas, 2012; Hongdan et al., 2013; Batalha et al., 2015). The significant

increase in lignin content after autohydrolysis might be attributed to the removal of

significant amount of hemicellulose while retaining most of the lignin. Some

researchers suggested that the increased lignin content might be due to the

repolymerization of polysaccharides degradation product (such as furfural) and/or

polymerization with lignin, which forms a lignin like material termed as pseudo-lignin

(Li et al., 2007). The pseudo-lignin can also be generated from carbohydrate without

significant contribution from lignin, especially under high severity pretreatment

conditions (Sannigrahi et al., 2011).

(b) Steam explosion pretreatment

During steam explosion pretreatment, saturated steam (under high pressure) is

inserted into a reactor (filled with biomass) that raises the temperature to 160-270°C.

Following gas insertion, the pressure is abruptly reduced that explodes the biomass

with degradation of lignin and hemicellulose (Mabee et al., 2006). The extent of the

biomass disruption depends on particle size, temperature, residence time and moisture

content (Sun and Cheng, 2002). It is believed that, this pretreatment enhances

cellulose crystallinity by converting amorphous portion of cellulose in to crystalline.

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During steam hydrolysis, the acetyl group associated with hemicelluloses generates

acetic acids that help in degradation of hemicelluloses. The removal of hemicelluloses

from the surface enhances cellulose accessibility and enzymatic digestibility (Kabel et

al., 2007). In some cases like pretreatment of softwoods require addition of catalyst

like H2SO4 or SO2 because of low content of acetyl group present in their

hemicelluloses (Mackie et al., 1985). Martín et al., (2002) studied the effect of steam

explosion of sugarcane bagasse impregnated with different agents and found that

H2SO4 impregnated bagasse showed comparatively higher glucose yield i.e. 35%.

This pretreatment strategy can induce the formation of inhibitors; therefore, washing

of biomass is prerequisite before subsequent enzymatic hydrolysis and fermentation

(García-Aparicio et al., 2006). Steam explosion is considered as cost effective

pretreatment due to low energy requirement and high energy efficiency.

(c) Ammonia Fiber Explosion (AFEX)

This process is similar to steam explosions; however, during this process liquid

ammonia is used, and the process is carried out at moderate temperatures (60-120°C)

for less than 30 min with subsequent drop in pressure (Kumar et al., 2009; Bals et al.,

2011). Commonly, 1-2 kg of ammonia is loaded per kg of biomass. Similar to other

alkaline pretreatment, AFEX plays an important role to alter lignin structure. This

strategy doesn’t release the sugars due to low hemicelluloses solubilization; however,

this opens up the lignocellulosic structure (Chundawat et al., 2007). The cost of

ammonia and its recovery affect the price of process; however, ammonia can easily be

recovered (Holtzapple et al., 1992). Krishnan et al., (2010) studied on alkali based

AFEX pretreatment of bagasse and reported that it enhance the cellulose digestibility

up to 85%. He also reported that AFEX pretreatment plays important role in

breakdown of ester bond and other LCC linkages thus enhances cellulose and

hemicelluloses accessibility during pretreatment.

(d) CO2 Explosion

This method is similar to steam and AFEX explosions; however, this involves

insertion of high pressure CO2 into reactor followed by sudden pressure drop (Zheng

et al., 1995). In comparison to other explosive strategy, this technique requires low

temperature and less cost as compared to AFEX. Other advantages are related to its

non-inflammability and non-toxicity (Zheng et al., 1995). CO2 explosion also retains

the properties of acidic hydrolysis by formation of carbonic acid (due to reaction of

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CO2 with water); however, carbonic acid is less corrosive as compared to other acidic

pretreatment. Moreover, due to easy removal of CO2, it doesn’t create any waste or no

further processing is required before downstream enzymatic hydrolysis and

fermentation (Schacht et al., 2008).

2.7.4. Biological Pretreatment

The process of biological pretreatment involves variety of microorganisms to degrade

cellulose, hemicelluloses and lignin. Different studies have been done on brown rot,

white rot and soft rot fungi to degrade lignocellulosic material (Ghose, 1978). These

fungi produce lignin modifying enzyme, lignin peroxidases and manganese dependent

peroxidases which breakdowns the intact lignocellulosic structure by oxidative

reactions (Fan et al., 1987; Malherbe and Cloete, 2002). Brown rot fungi mainly

degrade cellulose; whereas white rot fungi and soft rot fungi degrades lignin and

cellulose. Specifically, White rot fungi is the most prominent fungi for lignin

degradation. Hatakka et al. (1983) studied that 35% of wheat straw was converted

into sugar when it was degraded with white rot fungi i.e. Pleurotus ostreatus;

however, the time required to complete that process was estimated as five weeks.

Keller et al., (2003) studied the effect of pretreatment with Cyathus stercoreus on

corn stover and determined three to five fold increase in digestion during subsequent

enzymatic hydrolysis. Kurakake et al. (2007) studied the effect of two bacteria

Sphingomonas paucimobilis and Bacillus circulans on paper mill waste and

determined 94% sugar recovery. Patel et al., (2007) pretreated sugarcane bagasse with

Phenerochaete chrysosporium and resulted in high amount of sugar obtained. He also

pretreated sugarcane bagasse with Aspergillus awamori and Pleurotus sajor-caju and

high amount of ethanol was obtained. The absence of chemicals and mild conditions

requirement creates the interest of scientists on this type of pretreatment; however,

major drawbacks associated with this pretreatment are slow rate of hydrolysis, due to

these reasons this process is not considered as an economically feasible process.

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Fig.2.3. Separation of cellulose, hemicelluloses and lignin after pretreatment of

lignocellulosic biomass (Mood et. al., 2013)

2.8. Enzymatic Hydrolysis

Enzymatic hydrolysis is an important step between pretreatment and fermentation.

Enzymatic hydrolysis is prerequisite to convert polymeric cellulose and

hemicelluloses into monomeric fermentable sugars. Lignin, acetyl and cellulose

crystallinity are the major factors that makes the process of enzymatic hydrolysis

difficult. Lignin acts as the solid block that hinders the activity of enzymes. Acetyl

group may involve in inhibition of enzymes and microbes. However, crystallinity of

cellulose makes it difficult for enzymes to find the site of adsorption for their function

(Chang and Holtzapple, 2000). Intact lignocellulosic structure is first degraded and

opened by different pretreatments, which is then hydrolyzed into fermentable sugars

during the process of enzymatic hydrolysis.

The hydrolysis mechanism involves three steps i.e. adsorption of enzyme, hydrolysis

of celluloses and hemicelluloses, and desorption. Cellulases are mainly divided into

three types on the basis of their mode of action i.e. endoglucanases, exoglucanases

and β-glucosidases (Philippidis, 1994). During first step of hydrolysis, endoglucanase

acts on amorphous part of cellulose and breaks down intermolecular bonds between

adjacent cellulose chains. Moreover, it acts on glycosidic bonds and creates new ends

at crystalline fractions for exoglucanase activity. Exoglucanase starts acting at the

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ends of cellulose chains and releases cellobiose and also glucose (Zhang and Lynd,

2004). In last step β-glucosidase converts cellobiose into fermentable monomeric

sugar i.e. glucose (Teeri, 1997). Cellulases have been produced by bacteria, plants and

fungi; however, fungi showed better results and some strains of Trichoderma reesei

produced 30g/l of cellulases (Chambel, 2008). The hemicelluloses of lignocellulosic

biomass are also degraded by similar enzymes like xylanases, mannases, arabinases

etc. (Saha, 2003).

Figure.2.4. Simplistic overview of some factors limiting efficient hydrolysis of

cellulose: (1) Product inhibition of cellobiohydrolases by cellobiose (majorly) and

glucose, respectively; (2) Unproductive binding of exoglucanases onto a cellulose

chain. (3) and (4) Hemicelluloses and lignin associated with or covering the

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microfibrils prevent the cellulases from accessing the cellulose surface; (5) Enzymes

(both cellulases and hemicellulases) can be non-specifically adsorbed onto soluble

lignin particles or surfaces; (6) Denaturation or loss of enzyme activity due to

mechanical shear, proteolytic activity or low thermostability (7) Product inhibition of

β-glucosidases by glucose (Jørgensen et al., 2007).

2.9. Fermentation

2.9.1. Separate Hydrolysis and Fermentation (SHF)

SHF is the simplest configuration during which each step is separated and operated at

its optimum conditions. During this process, a small portion of feedstock is used for

enzymatic production and then the whole batch of pretreated biomass is added to

reactor for enzymatic hydrolysis, which is subsequently fermented in a separate

fermenter. In fermenter, microbes convert pentoses and hexoses into bioethanol that is

later separated out by distillation. The major advantage of SHF is related to carrying

out all processes at their optimal physicochemical parameters; however, the main

disadvantage is that the sugar released inhibits enzymatic activity (Goldschmidt,

2008)

2.9.2. Simultaneous Saccharification and Fermentation (SSF)

During SSF, the enzymatic hydrolysis and fermentation process is carried out in same

reactor. The main advantage of this process is the reduction of cost that is related to

the use of separate reactors for hydrolysis and fermentation. Moreover, it also reduces

the possibility of enzyme’s feedback inhibition, which is caused by accumulation of

released sugar as a result to enzymatic hydrolysis (Kim, 2004). During this process,

sugar release is abruptly converted into ethanol; therefore, it also reduces the chances

of contamination. The bottleneck of this strategy is compromised physicochemical

conditions (Goldschmidt, 2008). Enzymatic hydrolysis is usually carried out at 40-

50°C (for common cellulases); whereas, the optimum temperature for fermentation is

around 30°C (Barta, 2011). Therefore, the process is carried out at 35-37°C, which

requires more amount of enzyme to complete hydrolytic process. Recent researches

are being done to develop thermo-tolerant enzymes, which can improve SSF

(Goldschmidt, 2008; Barta, 2011).

2.9.3. Simultaneous Saccharification Fermentation and Co-fermentation (SSFC)

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SSFC also helps to reduce the cost eliminating the need of separate reactor. During

this process, hexose and pentose fermenting microbes are added to convert it into

bioethanol; however, there is the competition between hexoses and pentoses for the

same organism that reduce the overall ethanol yield. Due to this reason this process

could not be applied to industrial scale (Goldschmidt, 2008).

2.10. Biofuel from sugarcane: A solution of Food vs. Fuel Debate

In 2008, due to the international community's concerns regarding rise in food price,

the food vs. fuel debate reached a global scale. A United Nations Special

Rapporteur on the Right to Food, Jean Ziegler named biofuels a "crime against

humanity" (Pedro, 2008), and similar concerns were shown by the World Bank's

President, Robert Zoellick (Elliott and Stewart, 2008). Later, Luiz Inácio Lula

(Brazilian President) gave a strong rejection to these claims by putting all the blame

instead on U.S. and European agricultural subsidies, and said that the problem is

restricted to U.S. ethanol produced from corn. The Brazilian President has also

claimed that his country's sugarcane based ethanol industry has not contributed to the

food price crises (Colitt, 2008). In June 2008, Oxfam released a report, which

criticized rich countries for their biofuel policies and claimed that it is not solving oil

or climate prices, but increasing the problem of food shortage. The report also include

that sugarcane based fuel is most favorable in term of greenhouse gas balance and

cost, whereas rich countries spent $15 million to support biofuels while blocking

sugarcane ethanol (Oxfam, 2008). A World Bank report released on July 2008,

reported that US and European fuels are major cause of increase in food price;

however, sugarcane biofuel did not affect the sugar price (Veja Magazine, 2008). In

July 2008, OECD published a report that agreed with World Bank report and critically

evaluated the limited effect on reduction of greenhouse gas (GHG) emission from the

biofuel generation by US and Europe. However, OECD claimed that biofuel

generated from sugarcane reduces GHG emission by 80% as compared to fossil fuels

(OECD, 2008).

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Chapter 03

Materials and Methods

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On the basis of types of the waste generated by sugar industry, this research was

divided in to two main parts. First part discussed the bioethanol production from

sugarcane molasses, and all these experiments were done in Department of

Microbiology, Quaid-i-Azam University, Islamabad, Pakistan. The second part of this

research focused the bioethanol production from sugarcane bagasse, and it was carried

out at The Department of Chemical and Biomolecular Engineering, The University of

Tennessee, Knoxville, USA.

PART A: ENHANCED PRODUCTION OF BIOETHANOL FROM

SUGARCANE MOLASSES

3.1. PHYSICOCHEMICAL PROPERTIES OF MOLASSES

3.1.1. Determination of Total Dissolved Solids (Brix)

The molasses was obtained from Murree Brewery, Rawalpindi, Pakistan. The total

dissolved solid of molasses (also called brix) was determined with the help of brix

refractometer. For this purpose, 1 g of molasses was added into 100 ml of water (in

500 ml beaker) and stirred with the help of magnetic stirrer bar to get a homogenized

mixture, and then filtered through Whatman filter paper. The temperature of the

solution was kept 20ºC by placing it inside water bath. A drop of solution was placed

on the dark rectangular surface of refractometer and its lid was closed to read the

scale. Final brix of molasses was calculated by multiplying scale reading with dilution

factor. This experiment was repeated in triplicates.

3.1.2. Determination of Specific gravity

To determine the specific gravity of molasses, 5 g of molasses was added into 500 ml

of water and dissolved with magnetic stirrer. After getting homogenized mixture, its

temperature was set at 20ºC in water bath and the solution was poured in to measuring

cylinder. The gravity hydrometer was placed inside the solution and allowed to buoy.

When all the air was removed and gravity hydrometer became centered and vertically

positioned then its reading was determined. The specific gravity was determined in

triplicates. Original specific gravity of molasses was calculated by multiplying the

specific gravity with dilution factor.

3.1.3. Determination of Reducing Sugar

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The reducing sugars in molasses were determined by dinitrosalicylic acid (DNS)

method proposed by Miller (1959). In this method, 1 g of molasses was dissolved in

100 ml of water with the help of magnetic stirrer bar. 1 ml of molasses solution was

taken in a separate test tube and 3 ml of DNS reagent was added. The tube was heated

at 90ºC in water bath for 5 min. The tube was dipped in ice cold water to stop the

reaction, and the optical density (OD) was noted at 540 nm with the help of

spectrophotometer. The OD values were compared with standard glucose calibration

curve formulated by same DNS method to determine the reducing sugar of molasses.

3.1.4. Determination of Total Sugars in Molasses

In sugarcane industry, Lane and Eynon method (1923) was employed for total sugar

determination in sugarcane molasses. To determine the total reducing sugar in

molasses, 5 g of molasses was added into 100 ml of distilled water along with the

addition of 5 ml of HCl in Erlenmeyer flask. The molasses was placed in water bath at

70ºC for 10 min to convert sucrose in to monomeric reducing sugars. After treatment,

NaOH was added to neutralize the solution and volume was increased up to 1000 ml,

after that a burette was filled with this solution. The Fehling’s A solution (12.5 ml)

was mixed with Fehling’s B solution (12.5 ml) in a conical flask and 25 ml of distilled

water was added in it and titrated against molasses dilution (filled in burette) at

boiling temperature until the color faded. A few drops of methylene blue indicator

was then added that turned solution to blue color, which was again titrated against

neutralized molasses dilution until disappearance of blue color and appearance of

brick red precipitates. Fehling’s factor was determined by repeating this titration with

known concentration of standard sugar solution filled in burette.

Sugar Concentration (%) = (Dilution factor × Fehling’s factor) × 100

Titrate value x Weight of sample (g) x1000

3.1.5. Determination of Non-Reducing Sugar

Sucrose is most abundant non reducing sugar in molasses, which doesn’t have

reducing ends to react with DNS reagent. The non-reducing sugar in molasses was

determined by following formula:

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Non-reducing Sugars (%) = [Total Sugar (%) - Reducing Sugar (%)] x 0.95

3.2. ISOLATION, SCREENING AND CHARACTERIZATION OF INDIGENOUS

YEAST STRAINS

3.2.1. Sample Collection

Different variety of fruits was selected for the isolation of yeast strains. Grapes,

Strawberry and Carrot were purchased from local market in Islamabad, Pakistan. Soil

sample was obtained from sugarcane field near Multan, Pakistan.

3.2.2. Sample Processing

The fruits collected from different sources were thoroughly washed with distilled

water to remove any chance of contamination. All the fruits were crushed with mortar

and pistil under sterilized conditions and placed separately in autoclaved reagent

bottles for two days at room temperature.

3.2.3. Isolation

Different varieties of yeasts were isolated by serial dilution of fruits and soil samples.

Crushed fruit’s juice (1 ml) was added in 9 ml of sterilized distilled water in test tube.

This solution was serially diluted in ten other test tubes (containing 9 ml of sterilized

distilled water in each) to make dilutions from 10-1

to 10-10

. All test tubes were

vortexed before next dilution to ensure the equal distribution of yeast cells. Similarly,

1 g of soil was added in 9 ml of sterilized distilled water and serially diluted in similar

fashion. All dilutions were plated on Wallerstein Laboratory Nutrient (WLN) agar

medium by spread plate method and petri plates were incubated at 30ºC for 48 to 72 h

(Appendix A). After incubation different colonies were selected on the basis of

different morphological characteristics and purified on separate WLN medium using

streak plate method.

3.2.4. Quantitative Screening

(a) Inoculum Preparation

All isolated yeast strains were grown on YPD broth. YPD medium was prepared by

mixing 1 g of yeast extract, 2 g of dextrose and 2 g of peptone in 100 ml of distilled

water and autoclaved. About 25 ml of YPD broth was taken in 100 ml Erlenmeyer

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flask and inoculated with a loop full of yeast strain. Same procedure was followed for

all isolated yeast strains and all flasks were incubated at 30ºC on shaker incubator

with 100 rpm. After 24 h, yeast cultures were used as inoculum for quantitative

screening (i.e. ethanol tolerance test) to select a comparatively better strain for

enhanced ethanol production.

(b) Ethanol Tolerance Test

Quantitative screening was done by determining tolerance of each isolated strain

against 10% and 15% ethanol concentration. For each yeast strain a set of two 250 ml

Erlenmeyer flasks with 50 ml YPD broths was prepared containing 10% and 15%

ethanol. All these sets of flasks were inoculated with 1 ml of 24 h old culture of their

respective strain and incubated at 30ºC for 72 h. Optical density was measured at 600

nm with spectrophotometer after every 24 h. The most ethanol tolerant strain was

selected for further study.

3.2.5. Identification of Selected Yeast

The selected strain was identified on the basis of morphological and molecular

characteristics.

3.2.5.1. Morphological Identification

The most ethanol tolerant strain was grown in WLN agar media and its morphology

was studied on plates after 48 h of growth. The texture, color and surface of colony

were examined. The microscopic examination of selected yeast was also carried out

by preparing slide in saline solution (0.9% NaCl) and yeast cells were examined by

using 100x magnification.

3.2.5.2. Molecular Characterization

(a) DNA Extraction

DNA isolation was carried out with a little modification in phenol chloroform

method. Selected yeast strain i.e. MZ-4 was cultured in YPD broth for 24 h, and then

1.5 ml of the yeast culture was taken in eppendorf and centrifuged for 5 min at 14000

rpm. Supernatant was removed and 400 µl distilled water was added into pellet along

with 200 mg of glass beads. In addition, 300 µL of 0.1M phosphate buffer, pH 8.0

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(Sigma-Aldrich); 300 µL of 50mM sodium acetate (Sigma-Aldrich); 50 µL Sodium

dodecyl sulfate solution alkaline solution (Sigma-Aldrich); 10mM EDTA (Sigma-

Aldrich), pH 5.5; 400 µL phenol–chloroform–isoamyl alcohol (25:24:1), pH 8.0

(Sigma-Aldrich) was added in the eppendorf and vortexed for 2 min and then it was

placed in ice bath for 3 min. The eppendorf was then centrifuged at 14000 rpm for 15

min and supernatant was collected which was washed once with phenol-chloroform-

isoamylacohal (25:24:1) and afterwards with chloroform-isoamyl alcohol (24:1).

Again supernatant was collected and double the amount of absolute alcohol was

added into it and placed it overnight at -20ºC to precipitate out DNA. Next day it was

again centrifuged at 14000 rpm for 10 min and pellet was washed with 80% alcohol

and dried afterwards. The DNA pellet collected at the end was dissolved in 100 µL of

1x Tris EDTA Buffer and stored at -20°C for further use.

(b) PCR Amplification

Using primers ITS1 50-TCCGTAGGTGAACCTGCGG-30 and ITS4 50-

TCCTCCGCTTATTGATATGC-30, the DNA fragments of the ITS1, 5.8S RNA, and

ITS2 regions were amplified by PCR. PCR reactions were carried out in an AB 2720

thermal cycler (USA) with controlled amplification conditions i.e. initial denaturation

at 94oC for 5 min, followed by 30 cycles of denaturation at 94

oC for 30 s, annealing at

55oC for 30 s, extension at 72

oC for 1 min, and the final extension at 72

oC for 10 min

(White et al., 1990).

(c) Sequence Analysis

For molecular determination of yeast strain, 18S rRNA sequence of isolated DNA

was determined and provided by Macrogen Seoul Korea which was then examined

through nucleotide blast programs of NCBI (National center for biotechnology

information) to determine the homology against partial sequencing of 18S rRNA

(Appendix B).

(d) Phylogenetic Analysis and Submission to Gene Bank

The nucleotide blast program ―Blastn‖ was used for searching the homology against

the partial 18S rRNA sequences. The sequences from NCBI gene bank, having

maximum score and percentage for sequence homology were retrieved, and

alignment of these sequence were done by Molecular Evolutionary Genetic Analysis

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(MEGA) program version 6.06. These sequences were later submitted to NCBI gene

bank where it was assigned by accession number.

3.3. SELECTION OF BEST COMMERCIAL YEAST STRAIN TO COMPARE

WITH BEST INDIGENOUS YEAST STRAIN

3.3.1. Source of Commercial Strains

Four commercial strains of S. cerevisiae were compared with newly isolated

indigenous strain (MZ-4) to determine the most efficient strain for enhanced

production of bioethanol from sugarcane molasses. Rossmoor strains has already been

reported to be used in baking industry (Sultana et. al., 2013) and purchased by local

market in Islamabad, Pakistan. Saf instant was purchased from local vendor in

Islamabad, Pakistan while Uvafem-43 was provided by Murree Brewery, Rawalpindi,

Pakistan and both these strains has been reported for alcohol production from various

sources (Bechem et. al., 2007; Schmidt et. al., 2011). Lalvin EC-1118 was Canadian

strain, and famous for Champaign production from grapes (Valero et al., 2005 and

Carreto et. al., 2008).

3.3.2. Maintenance of Cultures

All the yeast strains selected for study were maintained by culturing on YPD agar

medium. The commercial strains were first added in autoclaved water (1:2 yeast to

water ratio at 30ºC) and after 10 min it was streaked on YPD agar plate. Self-isolated

strain MZ-4 was also transferred from WLN to YPD. All plates were incubated at

30ºC for 48 h and then transferred to refrigerator for preservation.

3.3.3. Strain Activation and Inoculum Preparation

A standard method used in most of brewing industries for inoculum preparation

before ethanol fermentation was followed. Sugarcane molasses dilution having 1.030

sp.grv (5% sugar conc) was prepared by distilled water with the help of gravity

hydrometer. Low gravity dilutions were prepared to prevent the strains from adverse

effect of high osmotic pressure which could be created due to high gravity molasses.

Erlenmeyer flask (100 ml) was filled with 10 ml of the molasses dilution, which was

then autoclaved and incubated at 30ºC overnight for sterility check. After 24 h, a loop

full culture was collected from previously preserved petri plates and transferred into

these flasks which were then incubated in shaker incubator at 30ºC with 100 rpm for

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24 h. After completion of incubation time, 20 ml of the yeast culture was transferred

into Erlenmeyer flask (500 ml) containing 250 ml of sterilized molasses dilution

(1.050 sp. grv; 9% sugar conc.) and again incubated at 30ºC for 24 h in shaker

incubator with 100 rpm for inoculum development. This stepwise increase in

molasses gravity was adopted to acclimatize yeast cells to high osmotic environment.

Same procedure was followed for all the yeast strains and repeated for all the

experiments on molasses.

3.3.4. Osmotic Tolerance

Sugarcane molasses was diluted with distilled water to prepare various concentrations

between 18-58% (w/v) and specific gravity values in range of 1.050-1.150, with the

help of gravity hydrometer. The sugar concentration was estimated between 9-29%

(w/v) by dividing total sugar added with dilution factor. These set of dilutions were

prepared for all of the selected stains and inoculated by 24 h old inoculum of each

strain and incubated at 30ºC for 72 h. After completion of reaction, all the dilutions

were distilled to determine ethanol concentration with the help of high performance

liquid chromatography (HPLC) and fermentation efficiency of all the strains at

different sugar concentration was also calculated by the formula:

Fermentation efficiency % = Actual Ethanol yield × 100

Theoretical Yield

Where, Theoretical Yield (v/v) = Sugar concentration × 0.64

On the basis of actual ethanol yield and fermentation efficiency, most osmotic tolerant

commercial strains were selected and compared with best self-isolated indigenous

strain (MZ-4) under optimized physicochemical conditions for enhanced production

of bioethanol from molasses.

3.3.5. Effect of Physicochemical and Nutritional Parameters

Different physicochemical parameters i.e. pH, temperature, inoculum size and age

were optimized for best selected yeast strains to get maximum ethanol production

from sugarcane molasses. Effect of different nitrogen sources and chelating agents on

these strains for enhanced ethanol production was also determined.

(a) Effect of pH

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To study the optimized pH value of yeast strains, 500 ml molasses dilution having

optimized sugar concentration was taken in Erlenmeyer flasks (3 L capacity) and their

pH was adjusted in range of 3.0-6.0 with the help of 5N H2SO4 and 5N NaOH. All the

flasks were inoculated with 5% (v/v) of 24 h old yeast inoculum and incubated at

30ºC for 72 h. Same procedure was repeated for all the yeast strains selected from

previous experiments.

(b) Effect of Temperature

To determine the best temperature for all selected strains, 500 ml molasses dilution

which contained optimized sugar concentration in 3L Erlenmeyer flask was adjusted

with previously optimized pH with respect to the specific inoculating strain. All flasks

were inoculated with 5% (v/v) of 24 h old inoculum of that strain and incubated at

different temperatures ranges 27-39ºC for 72 h. Same experiment was repeated for all

selected strains.

(c) Effect of Inoculum size

Experiment was set on the basis of all previously optimized parameters and varying

quantity of inoculum was added in range of 2.5-12.5% (v/v) to determine the best

inoculum size of each selected strain. Yeast cells were stained with 0.01% (w/v)

methylene blue and live yeast cell count in 24 h old inoculum was determined by

haemocytometer. The inocula of all selected strains were centrifuged at 2500 rpm for

5 min and the supernatant was discarded. Sterilized saline solution [0.9% (w/v) NaCl]

was added to yeast cell pellet to dilute the cells concentration up to 300x106 per

milliliter counts. Ethanol content was determined in fermentation media after 72 h of

incubation at the optimized temperature of each strain.

(d) Effect of Inoculum Age

To determine the effect of inoculum age, selected strains were cultured for 12, 24, 36

and 48 h. Then the inocula were centrifuged and diluted with saline to adjust cell

count up to 300x106 per milliliter. Already optimized size of inocula was used to

carry out fermentation process. All conditions were set to optimized values and

incubation was carried out for 72 h.

(d) Effect of Nitrogen Sources

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Nitrogen plays a significant role to enhance ethanol yield, due to which various

nitrogen sources were studied to determine their effect in enhanced ethanol

production. Urea, ammonium sulfate, ammonium nitrate, di-ammonium phosphate,

and ammonium chloride in various concentration ranges from 0.05-0.15% (w/v) were

added in molasses dilution at optimized sugar concentration, with adjusted optimized

pH value and inoculated with 5% (v/v) of 24 h old inoculum. The fermentation media

was then incubated at their optimum temperature for 72 h. Same experiment was

repeated for all selected strains.

(f) Effect of Chelating Agents

Chelating agents such as EDTA (Ethylenediamine tetraacetic acid), potassium

ferrocyanide, and sodium potassium tartrate were studied by adding them in various

concentrations ranges from 0.0025-0.32% (w/v) in to molasses dilutions. Effect of

chelating agent on fermentation by selected yeast strains was studied at their

optimized molasses concentration and pH by inoculating 5% (v/v) of 24 h old yeast

culture. The fermentation process was carried out by providing incubation at already

determined optimized temperature for 72 h. Same procedure was followed for all

strains.

After the completion of all the experiments, ethanol content was determined in

fermentation media with HPLC after distillation. The fermentation efficiencies of all

the selected strains were again calculated under optimized physicochemical

parameter.

3.4. FED BATCH FERMENTATION

The most ethanol tolerant comercial strain i.e. Lalvin EC-1118 (18% (v/v) ethanol tolerance)

was selected and compared with newly isolated strain MZ-4 (15% (v/v) ethanol tolerance),

and their ethanol producing potential in molasses was determined by fed-batch fermentation.

During fed-batch fermentation molasses was fed in intervals to reduce the osmotic stress

faced by fermenting microbes due to high sugar concentration in batch fermenter. For this

reason different molasses dilutions were prepared for each strain, having specific gravity

ranges from 1.080-1.140, with the help of gravity hydrometer. Four sets of flask for each

molasses dilution were prepared and they were assigned label 12, 24, 36 and 48 h. Molasses

dilutions (500 ml) were added to 5 L flask along with optimized nutrient addition and

inoculum and pH was adjusted to optimized value. All the flasks were incubated at the

already optimized temperature of the inoculated strain. All the four sets were fed after every

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12, 24, 36 and 48 h, respectively. The drop in specific gravity was noted after their assigned

time and more pure molasses was added into it to raise the specific gravity equals to initial

condition. Same procedure was repeated until no reduction in specific gravity was noted after

the assigned time interval. Fermentation media was distilled when the process was completed

and ethanol was determined by HPLC. Same procedure was repeated for all the three strain.

Fermentation efficiency of each strain at various specific gravity with different feeding rate

was also calculated.

3.5. ANALYTICAL METHODS

3.5.1. Lane and Eynon Method

Lane and Eynon titration method was used to determine reducing sugars in any

substance. Fehling’s solution A was prepared by dissolving 3.463 g of copper

sulphate (CuSO4.5H2O) in 50 ml water and filtered through Whatman filter paper.

Fehling’s solution B was prepared by adding 17.3 g of sodium potassium tartrate

(KNaC4H4O6.H2O) in 30 ml of water along with the solution of NaOH (prepared by

adding 5 g of NaOH in 5 g of water). The solution was allowed to cool and its volume

was increased up to 50 ml by adding distilled water. Methylene blue indicator (1%

w/v) was prepared by dissolving 1 g of indicator in 100 ml of water.

Standardization of Fehling’s Solution

Fehling’s solution A (12.5 ml) was mixed with Fehling’s solution B (12.5 ml) in 250

ml conical flask and 25 ml of water was added into same flask. Burette was filled with

standard glucose solution that was prepared by adding 1.25 g of glucose in 250 ml of

water. The Fehling’s solution was titrated against glucose solution at boiling

temperature and 23.5 ml of glucose solution was added from burette. Methylene blue

solution was added into Fehling’s solution and again few drops of glucose solution

were added from burette until the blue color completely disappeared. Exactly 24.1 ml

of glucose solution was used to standardize titration which showed the Fehling’s

solution is quite appropriate for sugar determination. The Fehling’s solution was then

used to determine the amount of total sugar by using factor: 25 mL of Fehling’s

solution = 0.1205 g of RS (reducing sugar)

3.5.2. Dinitrosalicylic acid (DNS) Method

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Reducing sugar can be determined by DNS method proposed by Miller (1959). The

DNS reagent was prepared by mixing 10 g of dinitrosalicylic acid and 10 g of NaOH

in 500 ml of water. In another flask, 200 g of potassium sodium tartrate was dissolved

in 200 ml water and 2 g of melted phenol was added into it. Both solutions were

mixed and volume was increased up to 1 L. A stock containing 0.1 g of glucose in

100 ml of water was prepared and glucose solutions of different concentrations were

prepared. In 1 ml of each dilution, 3 ml of DNS reagent was added and heated at 90ºC

for 5 min. Then all tubes were cooled in ice cold water then 200µl from each solution

was diluted to 2.5 ml of water and optical density was measured at 540nm to

formulate calibration curve (Appendix C.1). Sugar concentration of molasses sample

was determined by using linear regression equation obtained by standard curve i.e.

y = bx + a

Where the b represents slope, and a represents intercept of the line

3.5.3. Estimation of Alcohol Contents

3.5.3.1. Distillation Method

After completion of fermentation process, a known volume of fermentation media

(i.e. 100 ml) was distilled at 78ºC. The ethanol was collected at receiving flask. The

ethanol content in collected fraction was determined with the help of HPLC and the

concentration of ethanol in fermentation medium was determined by the formula:

C1V1= C2V2

Where C1 and V1 were the concentration and volume of fermentation media heated in

distillation flask, whereas C2 is concentration of the ethanol in distillate that was

determined by HPLC, and V2 is volume of the distillated collected in receiving flask.

High Performance Liquid Chromatography (HPLC)

Ethanol content in collected distillate was determined by HPLC (Waters 1525) with

IC-PAKTM

Ion Exclusion column [50 Aº 7µM (300 × 7.8mm)] and RI detector

(Waters 2410) by using 0.5 mM H2SO4 as the mobile phase and injection volume of

20 μl at flow rate of 0.5ml/min. Ethanol calibration curve was developed by using

various concentration of ethanol i.e. 10, 20, 30, 40 and 50% (Appendix C.2). All

samples collected after distillations were compared with this HPLC calibration curve

to determine the exact concentration of ethanol. All the standards and samples were

filtered through 0.2 μm cellulose-acetate filter paper before HPLC analysis.

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3.5.4. Yeast Cell Counting: (Mills, 1941)

Live yeast cells were counted with the help of haemocytometer already cleaned with

70% ethanol. For this purpose, 1ml of 24 h old inoculum was diluted in 10 to 100 ml

(depending upon the concentration of cells) distilled water. After dilution 0.5 ml of

sample was mixed with 0.5ml of 0.01% (w/v) methylene blue. A micropipette was

then used to transfer 10 µl of sample into haemocytomter and allowed the cells to

settle down for 1 min. The yeast cells were then examined under microscope with

100x magnification. All the dead cells were stained blue while the live cells were

colorless. Cells in four large corners were counted while neglecting cells touching left

and lower margins. The live cell count was determined by following formula:

Live yeast cell counts (cell/ml) = (live yeast count / No of squares) x 104

PART-B: ENHANCED PRODUCTION OF BIOETHANOL FROM

SUGARCANE BAGASSE

3.6. BIOMASS PREPARATION

3.6.1. Raw Material

Sugarcane bagasse was supplied by Green Energy Inc. Vonore, TN, USA. Bagasse (5-8 cm

size) was air-dried for two-three days. It was placed inside plastic bags and stored at 4ºC for

further use.

3.6.2. Drying and Milling

Bagasse was dried for two-three days until its moisture content became less than 10%. The

dried bagasse was milled in Thomas Model 4 Wiley® Mill fitted with 40 mesh screen to get

final particle size of about 0.45 mm.

3.6.3. Soxhlet Extraction

Removal of extractives present in lignocellulosic biomass is one of the important

steps while performing structural analysis, as these extractives can create problems by

interfering with other chemicals and reduce the precision of particular analysis. After

size reduction, sugarcane bagasse was extracted by placing the biomass into thimble

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of soxhlet extractor. The upper part of thimble was connected with condenser while

its lower part was fixed with receiving flask containing dichloromethane (1:10 mass

to liquid ratio). Biomass was refluxed with dichloromethane for 24 hours at 4-5

extractions per hour according to Tappi method T204 cm-07. After extraction,

biomass was removed from extraction thimble, and solvent was evaporated near to

dryness in the chemical fume hood.

3.7. PRETREATMENT STRATEGIES

3.7.1. Autohydrolysis of Sugarcane Bagasse

Sugarcane bagasse was treated with hot water in 4560 mini-parr pressure reactor.

Pretreatment conditions were selected on the basis of previously reported data.

Batalha et al. (2015) reported 190ºC±5 for 10 min and Peterson et al. (2009)

suggested 205ºC±5 for 6 min were comparatively better conditions for maximum

sugar recovery from lignocellulosic biomass. To compare the effectiveness of these

two conditions suggested in previous studies, 5 g of sugarcane bagasse was mixed

with 100 ml of water (20:1 liquid to solid ratio) and heated under these conditions in

mini Parr reactor separately. Untreated bagasse was considered as control.

Temperature increase rate at both conditions was determined as 4-5ºC per minute.

Pressure was noted as 14 Barr and 18 Barr when reactor heated at 190ºC and 205ºC,

respectively. After completion of pretreatment, reactor was quenched in ice cool

water for rapid cooling.

Filtration and Solid Recovery after Autohydrolysis

After autohydrolysis each set of pretreated bagasse was filtered through VWR Grade

417 filter paper with the help of vacuum filtration technique. Solid content was

collected in filter paper and kept inside fume hood for complete drying. Biomass after

autohydrolysis was not oven dried to prevent reduction in pore size and damage of

cellulose structure (Zhang et al., 2012).

3.7.2. Ionic liquid (IL) Pretreatment

Ionic liquid (1-butyl-3-methylimidazolium acetate) was added in sugarcane bagasse to

set liquid to solid ratio of 20:1 (5% w/v bagasse solution) and heated up to 110 C 5

for 30 min. The impeller was adjusted at speed of 100 rpm and reactor was heated to

specific temperature (at ~ 4-5 C/minute). After completion of pretreatment time the

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reactor was dipped into ice bath (~5 minutes) for rapid cooling. Untreated bagasse and

water treated bagasse were used as control to compare the effectiveness of ionic liquid

pretreatment. In water-treated bagasse sample, water was added instead of ionic liquid

and heated at same conditions i.e. 110ºC for 30 min. All experiments were conducted

in duplicates.

Filtration and Solid Recovery after IL pretreatment

Post pretreatment, deionized water was added into the IL and biomass solution at a

ratio of 5:1 (water: IL) to recover the biomass. Deionized water was added to it at

room temperature and stirred vigorously to enable biomass regeneration. The solids

were washed repeatedly with deionized water to remove any remaining IL from the

samples until the washed solution appeared colorless and solids were collected. The

ionic liquid/water mixture and biomass were separated by vacuum filtration through

VWR Grade 417 filter paper. Then recovered biomass was dried for two days before

further experiments (Zhang et al., 2012).

3.7.3. Severity Factor

In order to compare the efficacy of various pretreatment techniques, the

severity factor of all the pretreatments were determined by using the

equation:

SF = log (t x exp((T − Tref)/14.75))

In the above equation, t is the treatment time in min, T is the treatment temperature,

Tref is the reference temperature (i.e., 100°C) and 14.75 is an empirically determined

constant (Soudham et al., 2015)

3.8. COMPOSITIONAL ANALYSIS OF SUGARCANE BAGASSE

In order to monitor changes in biomass composition as a result of pretreatments and to

calculate the sugar conversion from enzymatic hydrolysis, carbohydrate and lignin analysis

were carried out based on methods described in NREL/TP-510-42618.

3.8.1. Lignin Determination

(a) Acid Insoluble lignin (Klason lignin)

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Acid Insoluble lignin (Klason lignin) of the control and pretreated samples were

determined by Tappi T-249 method. An oven dried sugarcane bagasse sample (0.175

g) was taken in a digestion tube and 1.5 ml of 72% (w/v) H2SO4 was added into it.

The sample was stirred with glass rod and after covering the cap with Parafilm,

digestion tube was placed in water bath at 30ºC. Glass rod was left in digestion tube to

stir it occasionally during primary hydrolysis. After one hour the sample was removed

from the water bath and 42 ml of distilled water was added into it for secondary

hydrolysis step during which bagasse was heated in 3% (w/v) H2SO4. Disposable

pipette was used to clean the glass rod during addition of water. The digestion tube

was loosely caped and was autoclaved at 121ºC for one hour. After autoclave, when

digestion tubes were cooled down, the solution inside tube was filtered through G8

(glass fiber filter) and more distilled water was added to make the volume of filtrate

up to 50 ml. A part of filtrate was used to determine acid soluble lignin and the rest

was stored in refrigerator for carbohydrate analysis. Glass fiber filter was prepared by

placing it in crucibles and washed by filtering distilled water through it; and then

crucible was left in an oven at 105ºC for 2 h. After drying, the crucible was removed

from oven and placed in desiccator for 20 min and weighed. After filtration of sample

through G8 filters, the remaining residue on filters were placed in oven at 105ºC and

left overnight. Next day, the crucibles were again cooled inside desiccator for 20 min

and weighed. The residue collected on fiber divided by the dry weight of the bagasse

(i.e. 0.175g) showed the weight of klason lignin per gram of substrate. Same

procedure was repeated in duplicates for all treated and untreated samples.

(b) Acid soluble Lignin (ASL)

Acid soluble lignin was analyzed in aliquots of filtrates by UV-Vis

Spectrophotometer within six hours after secondary hydrolysis. The absorbance was

noted at 240 nm by using 1cm light path Quartz cells and deionized water was used as

blank. The sample was diluted 4 times. The total insoluble lignin was determined by

the formula:

ASL% = (UVabs x Total volume of filtrate x Dilution factor ) x 100

є x ODWsample x Path length

Where,

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UVabs = average UV-Vis absorbance for the sample at 240nm

Volume hydrolysis liquor = volume of filtrate, 0.050 L

Dilution = (Volume of sample + Volume of diluting solvent)/Volume sample

ε = Absorptivity of biomass at 240nm = 24 (L/g-cm)

ODWsample = weight of sample in grams

Path length= path length of UV-Vis cell in cm

3.8.2. Carbohydrate Analysis

Sugar analysis after wet chemistry was done to determine compositional analysis of

pretreated and untreated biomass. All treated and untreated samples were analyzed for

glucan, xylan, arabinan and mannan content with the help of HPLC.

3.9. STRUCTURAL ANALYSIS OF SUGARCANE BAGASSE

3.9.1. Fourier Transform Infrared Spectroscopy (FTIR)

To investigate and estimate chemical changes in the sugarcane bagasse samples after each

pretreatment, a Perkin Elmer Spectrum 100 FTIR spectrometer equipped with an Attenuated

Total Reflectance (ATR) sampling accessory (Perkin-Elmer Inc., Wellesley, MA, United

States) was used. Samples (20 mg) were pressed uniformly against the crystal surface via a

spring-anvil, and spectra were obtained after 32 scans accumulation from 4,000 to 600 cm−1

at

4 cm−1

resolution. The ATR correction, which was performed using the formula AATRcorr

=

AATR

(k/k0) and setting the correction factor k0 to 1000 cm-1

(M. Milosevic, Internal

Reflection and ATR Spectroscopy (John Wiley & Sons, 2012)), and the baseline correction

were carried out by the Perkin-Elmer Spectrum software 8 (Perkin-Elmer Inc., Norwalk, CT,

United States). All FTIR graphs represented wavenumber in cm-1

on the x-axis and

absorbance on the y-axis (Sun et al., 2015)

.

3.9.2. X-ray Powder Diffraction (XRD)

Crystallinity index (CrI) of untreated, autohydrolysed and IL treated sugarcane

bagasse material was analyzed by X-ray diffractometer (PANalytical 3040/60 X'pert

PRO, Netherlands) with CuKα radiation source (k = 0.1505nm). Patterns were

collected from 5 to 40 (2θ) with scan step of 0.01 , while the operating voltage and

current were 40kV and 30mA, respectively. The crystallinity index (CrI) was defined

using the equation:

CrI= (I002-Iam)/I002

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Where I002 is the maximum intensity of crystalline peak at 2θ (22 ), whereas Iam is the

scattered intensity due to the amorphous portion evaluated as the minimum intensity

between the main (I002) and secondary peaks (I101) (Qiu et al., 2012)

3.10. ENZYMATIC SACCHARIFICATION

A combination of two commercially available enzymes cellulase (Celluclast® 1.5L,

Sigma Aldrich) and β-glucosidase from almond (CAS No: 9001223; Sigma Aldrich)

was used for the hydrolysis of untreated, water-treated and ionic liquid-treated energy

cane bagasse. The activity of cellulase was determined as filter paper assay unit (FPU)

by LAP (Laboratory Analytical Procedures) TP-510-42628 documented by NREL.

3.10.1. Enzyme Loading Optimization

(a) For Autohydrolysed Samples

Cellulase and β-glucosidase was added in different ratios (5FPU:10IU; 10FPU:20IU,

20FPU:40IU) to determine the exact loading amount for maximum sugar release from

pretreated biomass. In structural and compositional analysis, there was no wide

difference observed in cellulose crystallinity and quantity, among samples

autohydrolysed at 190ºC and 205ºC. Thus the optimization was carried out only by

using bagasse pretreated at 190ºC for 10 min. Reducing sugar released was

determined by DNS method (Miller, 1959) after every 24 h until the process

completed.

(b) For Ionic liquid Treated Samples

In order to determine the maximum loading required for the saccharification of IL

pretreated bagasse, cellulases and β-glucosidases were added in different ratios i.e.

5FPU:10IU; 10FPU:20IU, 20FPU:40IU; 40FPU:80IU. The best loading amount

determined for IL pretreatment was added for further enzymatic hydrolysis of IL-

pretreated biomass.

3.10.2. Enzymatic Hydrolysis

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All pretreated and untreated bagasse samples were subjected to enzymatic hydrolysis

by adding 1g substrate in 100 ml (1% w/v) of 50mM sodium citrated buffer (pH 4.8).

The optimized dosage of cellulases and β-glucosidases was chosen for hydrolytic

process. The process was carried in rotary incubator with a shaking speed of 150 rpm

at 50ºC for 72 h. Aliquots (500 µl) were collected at 3, 6, 12, 24, 48 and 72 h. Each

aliquot was sealed and incubated for 5 min at boiling water to denature the cellulases

(Zhang et al., 2012). The aliquots were filtered through 0.20 µm Nylon syringe filter

(Millipore, Billerica, MA) before high performance liquid chromatography (HPLC)

analysis. All experiments were performed in duplicates.

(a) Cellulose Digestibility Percentage

Percentage digestibility of cellulose was calculated as:

Percentage digestibility= (gram cellulose digested/ gram cellulose added) x100

(b) Xylan Digestibility Percentage

Percentage digestibility of xylan was calculated as:

Percentage digestibility= (gram xylan digested/ gram xylan added) x100

3.11. FERMENTATION

Three strains of S. cerevisiae and one strain of P. stipites (ATCC 58785) were used to

determine an efficient strain for enhanced ethanol production in case of each

pretreatment study. A newly isolated strain of S. cerevisiae MZ-4 and two commercial

strains of S. cerevisiae i.e. Lalvin EC-1118 and Uvaferm-43 were used for

fermentation process. After enzymatic saccharification, liquid part of all pretreated

samples were subsequently autoclaved and allowed to cool before inoculation, and all

the four strains were inoculated separately with inoculum size of 5% v/v (containing

300 x 106

living cells/ml). Ethanol content was measured from fermentation media

after every 12 h to determine maximum ethanol concentration produced per gram of

treated sample (Boopathy and Dawson, 2008; Singhania et al., 2014).

3.12. ANALYTICAL METHODS

Carbohydrate Analysis of Biomass by HPLC

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HPLC analysis was done to quantify the sugars released during enzyme hydrolysis,

and to determine the compositional analysis of bagasse. An HPLC system (Perkin

Elmer Flexar HPLC, Perkin Elmer, Shelton, CT) with Bio-Rad Aminex HPX-87P

column and refractive index detector was used (Appendix C.3-C.6). The temperature

for the column was set at 85ºC and H2O was used as the mobile phase with a flow rate

of 0.25 ml/min. The Chromera® Chromatography Data Systems (CDS) software was

used for the analysis and interpretation.

Determination of Cellulase Activity (Filter Paper Assay)

In order to measure the cellulase activity, 50 mg Whatman No. 1 filter paper strip (1.0

x6.0 cm) was rolled and placed into enzyme assay tubes. Sodium citrate buffer

(50mM; pH 4.8) of the amount 1.0 ml was added into these tubes and were

equilibrated at 50ºC. Different dilutions of enzymes were prepared in citrate buffer

and 0.5ml of enzyme dilution was added into enzyme assay tube. All the tubes were

incubated at 50ºC for 60 min. When reaction was completed, all the tubes were

removed from water bath and reaction was stopped by adding 3.0 ml of DNS in each

tube. Sodium citrate buffer (1.5 ml) was used as blank, whereas, a substrate control

(1.5 mL citrate buffer + filter-paper strip) and an enzyme control (i.e. 1.0 mL citrate

buffer + 0.5 mL enzyme dilution) were used as process controls. For each enzyme

dilution separate enzyme control was prepared. All blank and controls were also

incubated at 50ºC for 60 min and reaction was stopped by adding 3.0 ml DNS. All the

tubes were covered properly to prevent evaporation and boiled for 5 min in water

bath. The tubes were then transferred to ice cold water to terminate the reaction.

When all the pulp settled down, 0.2 ml of sample from each tube was diluted to 2.5 ml

of water and absorbance was determined by UV-vis spectrophotometry at 540 nm.

The readings were compared with glucose standard curve which was prepared by

adding DNS in various concentrations of glucose-buffer solutions. The enzyme

dilution which released 2mg/ml of the glucose was determined by linear regression

formula obtained by standard curve of glucose. The filter paper assay unit was

determined by formula:

Filter paper activity (units /ml) = [0.37] / [enzyme] releasing 2mg/ml glucose

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Where [enzyme] represents the proportion of original enzyme solution present in the

directly tested enzyme dilution (that dilution of which 0.5 mL is added to the assay

mixture)

3.13. STATISTICAL ANALYSIS

The mean and standard deviation was determined for all experiments. One way

Anova was used to determine significance across groups, and post-hoc tukey tests

were used for pairwise comparison to determine significance between groups, carried

out by IBM SPSS software 23. P-values of less than 0.05 were considered as

significantly different. Graph preparation and standard deviation calculations were

performed in Microsoft Excel, 2010.

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Chapter 4

Results

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Enhanced production of biofuel from sugar industry waste 63

PART A: ENHANCED PRODUCTION OF BIOETHANOL FROM

SUGARCANE MOLASSES

For the enhanced production of bioethanol from sugarcane molasses, this part of

research was further divided into four main sections. In first section, the chemical

composition of sugarcane molasses, which was obtained from Murree Brewery,

Rawalpindi, Pakistan, was determined. In the next section, ethanol tolerant yeast

strain was isolated from fruits and identified at molecular level. In third section, most

tolerant self-isolated strain was compared with the already available commercial

strains during optimization process. In last section, an attempt was made to enhance

the production of ethanol by fed-batch fermentation process.

4.1. PHYSICOCHEMICAL PROPERTIES OF MOLASSES

The physicochemical properties of molasses were determined by conventional

methods. The results of various experiments to determine physicochemical properties

of molasses were illustrated in Table. 4.1. The dissolved solid particle of molasses

was determined with the help of brix refractometer. It was an important parameter to

be studied before start of the work because most of the ethanol industries diluted their

molasses according to its ºbrix values. The ºbrix of molasses during current research

was noted as 79.0 brix. Specific gravity is another important parameter to determine

before the start of new experiments. The specific gravity noted with the help of

gravity hydrometer was 1.4. The sugar concentration of molasses was determined by

various conventional methods. As molasses contained both reducing as well as non-

reducing sugars, therefore, a modified Lane and Eynon method was first used to

determine its total sugar concentration. During this experiment, molasses was heated

with the addition of HCl and all the non-reducing disaccharides present in molasses

were converted into monomeric reducing sugars, which were then detected by

Fehling’s solution and calculated as 49% (w/w). This value showed the amount of

both reducing as well as inverted sugars. The DNS method was used to determine the

reducing sugars in non-treated molasses, which was noted as 15% (w/w) that

excluded the values of non-reducing sugar. The value of non reducing sugar was

determined as 32.3% (w/w). The ability of yeast to convert all the sugars (reducing

and non-reducing) into ethanol made it important to determine the concentration of

total sugars present in molasses.

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Table 4.1: Physicochemical Properties of Molasses

No Components Amount

1. Brix 79.0 °Bx ±1.0

2. Specific Gravity 1.4 ± 0.05

3. Reducing Sugar (w/v) 15.0 % ±0.50

4. Non Reducing Sugar (w/v) 32.3% ± 1.0

5. Total Sugar (w/v) 49.0% ±1.0

4.2. ISOLATION, SCREENING AND CHARACTERIZATION OF

INDIGENOUS YEAST STRAINS

4.2.1. Isolation and Screening

Variety of sources were selected and processed for isolation of yeast. The number of

strains isolated from grapes and strawberry were 22 and 3, respectively, whereas no

yeast strain could be isolated from carrot and soil samples. All yeast strains were then

subjected to grow in YPD broth medium containing 10% and 15% ethanol.

The results showed that five of the isolated strains i.e. MZ-1, MZ-4, MZ-12, MZ-18

and MZ-22 were able to grow in 10% ethanol containing YPD broth that represented

the hight tolerance of these strains against the ethanol concentration present in that

medium as compared to the rest of the strains (Table. 4.2.a). Only one strain MZ-4

exhibited comparatively better growth and tolerance upto 15% ethanol concentration

in YPD broth (4.2.b). Thus, the strain MZ-4 was selected for further studies on the

basis of its high ethanol tolerance so that it could be compared with some very good

ethanol tolerant commercially available strains.

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Table 4.2(a): Screening of Isolated Yeast Strains against 10% Ethanol Tolerance

Yeast strain Optical Density at 600nm after every 24 hour

0 h 24 h 48 h

MZ-1 1.502 2.327 2.500

MZ-2 0.53 0.524 0.510

MZ-3 0.345 0.524 0.590

MZ-4 0.965 2.001 2.221

MZ-5 0.652 0.599 0.59

MZ-6 1.986 2.060 1.856

MZ-7 1.521 1.321 1.32

MZ-8 0.53 0.521 0.501

MZ-9 0.963 1.546 2.570

MZ-10 0.632 0.624 0.312

MZ-11 1.011 0.915 0.832

MZ-12 0.937 1.815 1.945

MZ-13 1.021 0.950 0.810

MZ-14 1.342 1.321 1.212

MZ-15 0.702 0.654 0.525

MZ-16 2.012 1.985 1.895

MZ-17 2.222 2.170 2.135

MZ-18 0.399 0.416 0.723

MZ-19 0.77 0.759 0.750

MZ-20 1.551 1.542 1.540

MZ-21 2.19 2.195 2.001

MZ-22 0.852 1.416 1.850

MZ-23 0.645 0.652 0.552

MZ-24 1..511 1.501 1.382

MZ-25 1.1 1.918 1.824

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Table 4.2(b): Screening of Isolated Yeast Strains against 15% Ethanol Tolerance

Yeast strain Optical Density at 600nm after every 24 hour

0 h 24 h 48 h

MZ-1 0.421 0.316 0.220

MZ-2 0.366 0.304 0.310

MZ-3 0.459 0.455 0.420

MZ-4 0.502 0.669 0.920

MZ-5 0.54 0.504 0.501

MZ-6 0.499 0.459 0.439

MZ-7 0.51 0.499 0.503

MZ-8 0.503 0.507 0.506

MZ-9 0.507 0.513 0.486

MZ-10 0.52 0.519 0.515

MZ-11 0.401 0.414 0.400

MZ-12 0.313 0.303 0.302

MZ-13 0.515 0.523 0.521

MZ-14 0.45 0.456 0.411

MZ-15 0.32 0.315 0.301

MZ-16 0.315 0.321 0.312

MZ-17 0.59 0.600 0.508

MZ-18 310 0.299 0.298

MZ-19 0.344 0.358 0.350

MZ-20 0.5 0.507 0.500

MZ-21 0.46 0.458 0.369

MZ-22 0.453 0456 0.432

MZ-23 0.415 0.421 0.422

MZ-24 0.65 0.621 0.525

MZ-25 0.666 0.655 0.620

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4.2.2. Identification of Selected Yeasts

The strain MZ-4 was selected and identified through microscopic examination and

morphology on WLN agar plate. Buddings were clearly observed on microscopic

examination that considered as one of the major characteristics of yeast strain. Strain

MZ-4 has dark green, large, oval and elevated colonies on WLN medium, that showed

its resemblance to Saccharomyces, which was further confirmed by molecular

identification.

18S rRNA nucleotide sequence obtained from Macrogen Incorporation, Seoul, Korea,

was further analyzed by using NCBI blast technique. Strain MZ-4 showed 100%

similarity with Saccharomyces cerevisiae type strain S288c. The sequence was

submitted to gene bank and an accession number i.e. KP970869 was assigned. The

evolutionary history of strain MZ-4 was represented by formulating the phylogenetic

tree using neighbor-joining method with the help of Mega-6 software (Fig.4.1)

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Fig.4.1: Phylogenetic tree of isolated Saccharomyces cerevisiae strain MZ-4

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4.3. SELECTION OF BEST COMMERCIAL YEAST STRAIN TO COMPARE

WITH BEST INDIGENOUS YEAST STRAIN MZ-4

4.3.1. Selection of Commercial Strains

Four commercial strains i.e. Rossmoor, Saf-Gold, Uvaferm-43 and Lalvin EC-1118

were selected and their ethanol tolerance from already published data was found as 6,

12, 18 and 18% (v/v) respectively (Bechem et al., 2007; Schmidt et al., 2011; Sultana

et al., 2013). The strains exhibited comparatively more ethanol and osmotic tolerance

along with better fermentation efficiency was selected to compare with newly isolated

indigenous strain MZ-4.

4.3.2. Osmotic Tolerance

Osmotic tolerance of all the strains was determined by carrying out fermentation of

different molasses dilutions and quantifying the ethanol yield with the help of HPLC

(Table 4.3). It was observed that the ethanol production for all the strains was

increased with the increase in molasses concentration due to more sugar availability;

however, after certain concentration of molasses, the ethanol production was reduced

because of their osmotic intolerance at high sugar concentrations. It was observed that

the commercial strains i.e. Rossmoor and Saf-instant, was producing 6.5 and 7.5%

(v/v) of ethanol when 15 and 17% (w/v) of sugar was present in the fermentation

medium, respectively. However, Uvaferm-43, Lalvin EC-118 and MZ-4 were tolerant

to 25% (w/v) sugar and they produced maximum of 9.3, 9.6 and 10.1% (v/v) of

ethanol, respectively.

The fermentation efficiency of the process at different sugar concentration was also

calculated. It was determined that the fermentation efficiency of the process decreased

with increase in sugar concentration. The Rossmoor strain showed fermentation

efficiency of 72.6% in presence of 9% (w/v) sugar concentration and yielded 4.1%

ethanol. The increase in sugar concentration up to 15% (w/v) enhanced ethanol yield

up to 6.5% (v/v), but the fermentation efficiency reduced to 67.3%. In case of Saf-

instant yeast, the maximum fermentation efficiency of 75.6% was noted with 4.2% of

ethanol yield when 9% (w/v) sugar was present in molasses. The maximum amount of

ethanol produced by this strain was 7.5% (v/v) with 17% (w/v) sugar concentration;

however, the fermentation efficiency was reduced to 57.5%. Uvaferm-43, Lalvin EC-

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1118 and MZ-4 strains showed 85.1, 85.1 and 83.3% fermentation efficiency with the

production of 4.8, 4.8 and 4.7 % (v/v) of ethanol respectively. The increase in sugar

concentration up to 25% (w/v) enhanced their ethanol yield up to 9.3, 9.6, 10.1%

(v/v), but the fermentation efficiency was reduced to 58.1, 60.0 and 63.1%

respectively. On the basis of these results, the commercial strain Lalvin EC-1118 was

selected for further studies to compare with self-isolated MZ-4 strain, as both of these

strains were tolerant to 25% (w/v) sugar concentration and showed comparatively

better ethanol yield and fermentation efficiency as compared to other strains.

4.3.3. Ethanol Quantification by HPLC

The HPLC chromatogram for ethanol showed its peak with retention time of 14.6 min

(Fig 4.2). The calibration curve for ethanol was formulated by plotting the area of

peak (at 14.6 min) against different concentration of ethanol (Appendix C2). The

ethanol yield (Table 4.3) was determined by distillation of fermentation media after

completion of process, and distillate was run through HPLC to compare with

calibration curve.

Table.4.3: Enhanced production of bioethanol by various yeast strains using different concentration of sugar.

Sugar

(%)

Actual Ethanol yield % (v/v) Fermentation Efficiency (%)

Rossmmor SAF-

instant

Uvaferm

-43

Lalvin

EC-

1118

MZ-4 Ross

moor

SAF-

Instant

Uvaferm-

43

Lalvin

EC-118 MZ-4

9 4.1±0.08 4.2±0.04 4.8±0.08 4.8±0.02 4.7±0.16 72.6 75.6 85.1 85.1 83.3

11 4.8±0.12 5.3±0.08 5.7±0.12 6±0.12 5.7±0.04 68.5 75.2 81.9 85.2 81.91

13 5.5±0.04 6.2±0.16 6.9±0.08 6.4±0.16 5.9±0.08 66.4 74.4 82.8 76.8 71.2

15 6.5±0.12 7.1±0.08 7.2±0.16 7.5±0.02 6.4±0.14 67.3 73.9 75.00 78.1 67.3

17 6.4±0.12 7.5±0.04 7.8±0.08 7.8±0.16 7.2±0.08 59.1 69.2 71.6 72.3 66.1

19 6.1±0.08 7.0±0.08 8.1±0.08 8.1±0.08 7.8±0.02 50.1 57.8 66.6 67.1 64.1

21 5.4±0.04 6.7±0.04 8.5±0.08 8.5±0.04 8.4±0.04 40.4 50.3 63.2 63.7 62.9

23 4.3±0.08 6.1±0.08 8.7±0.08 9.2±0.14 9.2±0.04 29.2 41.4 59.1 62.7 62.9

25 3.6±0.04 5.6±0.16 9.3±0.08 9.6±0.16 10.1±0.12 22.9 35.0 58.1 60 63.1

27 3.4±0.08 5.4±0.08 8.5±0.12 8.6±0.02 8.1±0.14 19.6 31.2 49.5 49.9 46.8

29 2.4±0.08 4.7±0.12 7.7±0.08 8±0..18 7.5±0.02 12.9 25.6 41.4 43.1 40.4

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Fig.4.2: HPLC chromatogram for ethanol detection

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4.3.4. Effect of Physicochemical and Nutritional Parameters

(a) Effect of pH

To determine the effect of pH on each of the selected strains, the production of

ethanol was studied by adjusting the pH in range of 3-6 (Fig.4.3, Appendix D1). It

was observed during the optimization study that the production of ethanol decreased

under either more acidic or more alkaline conditions. For the strain Lalvin EC-1118, it

was observed that the production of ethanol enhanced with the increase in pH of

medium; and it showed that ethanol production was significantly higher (P<0.05) i.e.

9.8% (v/v) when the pH was adjusted at 4.5 (Appendix F1). The further increase in

pH reduced the production of ethanol. Same trend was observed in MZ-4 strain which

showed that the increase in pH enhanced the ethanol content in fermentation medium

up to the pH 5.0. The maximum of 10.2% (v/v) of ethanol was produced at pH 5.0

(P<0.05); however, further increase in pH reduced the ethanol content obtained at the

end of fermentation (Appendix F1). Therefore, it could be concluded from the above

observations that the optimized pH value for both Lalvin EC-118 and MZ-4 was 4.5

and 5.0, respectively.

(b) Effect of Temperature

The effect of temperature on enhanced production of bioethanol by Lalvin EC-1118

and MZ-4 strains was studied in range of 27-39°C (Fig. 4.4, Appendix D2). During

this study, it was investigated that the production of ethanol increased with increase in

temperature; however, there was a certain limit for each microbial strain up to which

the temperature positively affected the activity of each strain. As the temperature

exceeded that specific point, a reduction in ethanol yield was clearly observed. The

study of temperature’s effect on Lalvin EC-1118 showed that the lower temperature

i.e. 27°C was not as effective for ethanol production as it was observed at 30°C;

however, ethanol production was decreased with further increase in temperature

beyond this limit. The production of ethanol was significantly higher (P<0.05) at

30°C for Lalvin EC-1118 strain (Appendix F2). Similar observation was recorded in

case of MZ-4 strain, which showed significantly higher ethanol production (P<0.05)

when the fermentation medium was incubated at 33°C, and the temperature

adjustment below or above 33°C adversely affected the fermentation process

(Appendix F2). Therefore, it was concluded that the optimum production of ethanol

by Lalvin EC-1118 [9.8% (v/v)] and MZ-4 [10.3% (v/v)] was achieved at 30°C and

33°C, respectively.

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Fig.4.3: Effect of pH on enhanced production of bioethanol by Lalvin EC-1118 and

MZ-4

Fig. 4.4: Effect of temperature on enhanced production of bioethanol by Lalvin EC-

1118 and MZ-4

0

2

4

6

8

10

12

3 3.5 4 4.5 5 5.5 6

Eth

ano

l % (

v/v)

pH

Lalvin EC-118 MZ4

0

2

4

6

8

10

12

27 30 33 36 39

Eth

ano

l % (

v/v)

Temperature (C)

Lalvin EC-118 MZ4

PhD Thesis


(c). Effect of Inoculum Size

After allowing the living cells to grow for certain time, different concentration of

exactly measured cell density was inoculated in order to get the highest concentration

of ethanol. The inoculum was added as a liquid culture; therefore, more amount of

inoculum added can dilute the reaction medium in terms of sugar concentration which

in turn reduces the final ethanol concentration. To determine the effect of inoculum

size, the inoculum having 3x108 cells/ml was prepared and the different volumes of

that inoculum in range 2.5-12.5% (v/v) was added in fermentation medium (Figure

4.5, Appendix D.3). It was observed that the production of ethanol was also enhanced

with the increase in inoculum; however, when the inoculum size increased beyond

certain limit, no further increase in ethanol content was observed in fermentation

medium. For the strain Lalvin EC-1118, significantly greater (P<0.05) amount of

ethanol was produced i.e. 10% (v/v), when 7.5% (v/v) of inoculum was added into the

medium. Similarly, 10.5% (v/v) of ethanol was produced when 10.0% (v/v) of MZ-4

inoculum was added into it, which was significantly higher (P<0.05) than other

inoculum sizes (Appendix F3). Further increase or decrease in inoculum reduced the

content of ethanol in fermentation medium.

(d). Effect of Inoculum Age

The effect of inoculum age was studied for both strains i.e. Lalvin EC-1118 and MZ-4

after developing the inoculum for different period of time i.e. 12-48 h (Fig. 4.6,

Appendix D4). Both the strains Lalvin EC-1118 and MZ-4 produced significantly

higher (P<0.05) ethanol of 10% and 10.5% (v/v) respectively when 24 h old inoculum

was used in fermentation medium (Appendix F4). The production was greatly reduced

with further increase in age of inoculum. In case of both strains i.e. MZ-4 and Lalvin

EC-1118, the best inoculum age was determined as 24 h. During inoculum

preparation, both strains after 24 h of incubation reached to actively growing log

phase. These actively growing cells experienced short span lag phase when inoculated

in a reaction flask. After 24 h the strains might be shifted to stationary or death phase

which adversely affected the density of living cells and reduced the ethanol

production.

PhD Thesis


Fig.4.5: Effect of inoculum size on enhanced production of bioethanol by Lalvin EC-

1118 and MZ-4

Fig. 4.6: Effect of inoculum age on enhanced production of bioethanol by Lalvin EC-

1118 and MZ-4

0

2

4

6

8

10

12

2.5 5 7.5 10 12.5

Eth

ano

l % (

v/v)

Innoculum % (v/v)

Lalvin EC-118 mz4

0

2

4

6

8

10

12

12hours 24hours 36hours 48hours

Eth

ano

l % (

v/v)

Inoculum age

Lalvin EC-118 MZ4

PhD Thesis


(e). Effect of Nitrogen Sources

To study the effect of different concentration of various nitrogen sources on Lalvin

EC-1118 and MZ-4, different concentrations of urea, ammonium chloride (NH4Cl),

ammonium nitrate (NH4NO3), ammonium sulfate ((NH4)2SO4) and di-ammonium

phosphate ((NH4)2HPO4) were added in fermentation media in range 0.05-0.15%

(w/v) to determine their effect on both of selected strains. The effect of nitrogen strain

on Lalvin EC-1118 is shown in Fig 4.7 and Appendix D5. Ethanol concentration was

increased from 10.1 to 10.2% (v/v) as the urea concentration was increased from 0.05

to 0.10% (w/v) in the medium inoculated with strain Lalvin EC-118. However, further

increase in urea concentration adversely affected the efficiency of this strain and it

reduced the ethanol production. In case of NH4Cl, the increase in its concentration

from 0.05 to 0.15% (w/v) resulted into enhancement in ethanol concentration from

10.1 to 10.4% (v/v). The addition of 0.05 % (w/v) NH4NO3 enhanced the ethanol

yield up to 10.3%; however, the further increase in its concentration reduced ethanol

content. The effect of (NH4)2SO4 on ethanol production was same as for NH4Cl. As

the amount of (NH4)2SO4 increased from 0.5 to 0.15%, the ethanol concentration

increased from 10.0 to 10.4%. The maximum concentration of ethanol was obtained

when (NH4)2HPO4 was added as nitrogen source, i.e., 10.3 to 10.5% ethanol was

produced after addition of 0.05 to 0.1% of (NH4)2HPO4; however, with increase in its

concentration up to 0.15%, the ethanol concentration reduced to 10.0%. After

observing all the data, it was concluded that (NH4)2HPO4 was the comparatively

better nitrogen source for the strain Lalvin EC-118 and it yields significantly higher

(P<0.05) ethanol concentration up to 10.5% (v/v) and required in less amount as

compared to (NH4)2SO4 and NH4Cl; therefore, it was added in fermentation media as

the best nitrogen source for further experiments (Appendix F5).

The effect of different nitrogen sources on enhanced production of bioethanol by MZ-

4 strain was also studied (Fig. 4.7, Appendix D5). It was found that increase in

amount of urea from 0.05 to 0.1% (w/v) in fermentation medium resulted

enhancement in ethanol yield from 10.5 to 10.7% (v/v). However, with further

increase in urea concentration, a decrease in amount of ethanol in fermentation

medium was observed. Similar trend was observed in the case of NH4Cl. An increase

in NH4Cl concentration from 0.05 to 0.1% (w/v) enhanced the ethanol yield from 10.6

to 10.8% (v/v); however, further increase resulted in reduction of ethanol

PhD Thesis


concentration. The effect of NH4NO3 and (NH4)2HPO4 on ethanol production was

found similar, 10.6% (v/v) of ethanol was produced in the presence of 0.1% (w/v) of

NH4NO3 and (NH4)2HPO4 in fermentation medium. Any increase or decrease in its

concentration reduced the final ethanol content in fermentation medium. In case of

(NH4)2SO4, ethanol concentration was increased from 10.5 to 10.7% (v/v) with an

increase in its concentration from 0.05 to 0.15%. From all the data, it was concluded

that, NH4Cl was the comparatively better nitrogen source for the strain MZ-4, that

could produce significantly higher (P<0.05) ethanol, i.e., 10.8% (v/v) at 0.1% (w/v) of

its concentration (Appendix F.6).

From the above study, it was concluded that the best nitrogen source for strain Lalvin

EC-1118 and MZ-4 was (NH4)2HPO4 and NH4Cl, 0.1% (w/v) of their concentration

resulted into 10.5% and 10.8% (v/v) of ethanol, respectively.

PhD Thesis


Fig.4.7: Effect of nitrogen source on enhanced production of bioethanol by Lalvin

EC-1118 and MZ-4

0

2

4

6

8

10

12

0.0

5%

0.1

0%

0.1

5%

0.0

5%

0.1

0%

0.1

5%

0.0

5%

0.1

0%

0.1

5%

0.0

5%

0.1

0%

0.1

5%

0.0

5%

0.1

0%

0.1

5%

Urea NH4Cl NH4NO3 (NH4)2SO4 (NH4)2HPO4

Eth

ano

l % (

v/v)

Nitrogen source (% w/v)

Lalvin EC-118

0

2

4

6

8

10

12

0.0

5%

0.1

0%

0.1

5%

0.0

5%

0.1

0%

0.1

5%

0.0

5%

0.1

0%

0.1

5%

0.0

5%

0.1

0%

0.1

5%

0.0

5%

0.1

0%

0.1

5%

Urea NH4Cl NH4NO3 (NH4)2SO4 (NH4)2HPO4

Eth

ano

l % (

v/v)

Nitogen source (% w/v)

MZ-4

PhD Thesis


(f). Effect of Chelating Agents

The effect of different chelating agents for the enhanced production of bioethanol

showed high significance difference (P<0.05) for the production of bioethanol.

Different types of chelating agents i.e. EDTA, K4Fe(CN)6, and NaK-Tartrate were

added in fermentation medium in concentration 0.0025-0.32 (w/v) (Fig 4.8, Appendix

D6) In case of Lalvin EC-118 strain, it was observed that the increased concentration

of EDTA in fermentation media also enhanced the ethanol yield. The addition of

0.04% (w/v) of EDTA and NaK-tartarate enhanced ethanol yield up to 10.7% (v/v);

however, further increase in its concentration adversely affected the ethanol

production. The maximum amount of ethanol obtained was 10.9% (v/v) when 0.04%

(w/v) of K4Fe(CN)6 was added into fermentation medium., which was significantly

greater (P<0.05) than the effect of other chelating agents (Appendix F7). However,

when the amount of chelating agent was increased more than 0.04% (w/v), it

adversely affected the fermentation process. Therefore; K4Fe(CN)6 was considered as

comparatively better option to be used in fermentation medium as compared to other

chelating agents for strain Lalvin EC-1118.

The effect of various chelating agents on ethanol production from strain MZ-4 was

also observed (Fig 4.8). An increase in ethanol concentration was observed with

increase in concentration of chelating agents. It was observed that the maximum of

10.9% (v/v) ethanol was produced when 0.04% (w/v) of EDTA was added into

fermentation medium, and the further increase or decrease in its concentration lowers

the ethanol production. Similarly, when 0.01% (w/v) of K4Fe(CN)6 was added into

fermentation medium, the production was increased up to the maximum of 11.1%

(v/v), which was significantly higher (P<0.05) than the effect of other chelating

compounds (Appendix F8). In case of NaK-tartarate, 10.9% (v/v) of ethanol was

produced in the presence of only 0.02% (w/v) of the chelating agent and its further

increase in concentration reduced the ethanol content. By comparing the effect of all

three chelating agents, it was concluded that K4Fe(CN)6 was the comparatively better

option to be used in fermentation medium, because it was required in low

concentration and yielded more ethanol as compared to other chelating agents.

PhD Thesis


0

2

4

6

8

10

120

.00

25

0.0

05

0.0

1

0.0

2

0.0

4

0.0

8

0.1

6

0.3

2

0.0

02

5

0.0

05

0.0

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0.0

2

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0.0

8

0.1

6

0.3

2

0.0

02

5

0.0

05

0.0

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0.0

2

0.0

4

0.0

8

0.1

6

0.3

2

EDTA K4FE(CN)6 NaK-Tartarate

Eth

ano

l % (

v/v)

Chelating agents ( % w/v)

Lalvin EC-118

0

2

4

6

8

10

12

0.0

02

5

0.0

05

0.0

1

0.0

2

0.0

4

0.0

8

0.1

6

0.3

2

0.0

02

5

0.0

05

0.0

1

0.0

2

0.0

4

0.0

8

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6

0.3

2

0.0

02

5

0.0

05

0.0

1

0.0

2

0.0

4

0.0

8

0.1

6

0.3

2

EDTA K4Fe(CN)6 NaK-Tartarate

Eth

anl (

% v

/v)

Chelating agents (%w/v)

MZ-4

PhD Thesis


Fig.4.8: Effect of chelating agents on enhanced production of bioethanol by Lalvin

EC-1118 and MZ-4

Fermentation Efficiency after Optimization

Fermentation efficiency of the process was determined again after optimization of

process and an increase in fermentation efficiency was noted (Table 4.4). It was

observed that the fermentation efficiency of strain Lalvin EC-118 was calculated as

60.0% with 9.6% (v/v) of bioethanol at 25% (w/v) of sugar concentration; however,

the efficiency was increased to 68.1% with 10.9% (v/v) of ethanol production after

optimization of physicochemical parameters utilizing same sugar concentration. In

case of MZ-4 strain, fermentation efficiency before optimization was calculated as

63.1% with 10.2% (v/v) ethanol yield with 25% (w/v) of sugar; however after

optimization it was increased up to 69.3% with 11.1% (v/v) ethanol yield with same

sugar concentration.

Table.4.4: Enhancement of fermentation efficiency due to optimization

Strains

Before optimization After optimization

Ethanol

Yield % (v/v)

Fermentation

Efficiency (%)

Ethanol

Yield % (v/v)

Fermentation

Efficiency (%)

Lalvin EC-

1118

9.6±0.16 60.0 10.2±0.16 68.1

MZ-4 10.1±0.12 63.1 11.1±0.16 69.3

4.3. FED BATCH FERMENTATION

During fed batch fermentation, substrate was added after specific intervals in order to

reduce the osmotic pressure of the fermentation medium faced by fermenting

microbes. The effect of specific gravity and feeding interval on enhanced production

of bioethanol was also studied. Those conditions were preferred for fed batch

fermentation which showed enhanced ethanol production and better fermentation

efficiency. The process completion time for each step was also determined because of

the time-constrained issues faced by industry.

PhD Thesis


4.3.1. Effect of Specific Gravity

The specific gravity was adjusted within range of 1.080-1.140 in order to determine

the best specific gravity for the fed-batch fermentation using strains Lalvin EC-118

and MZ-4 (Fig. 4.9). For the Lalvin EC-118, it was observed that the increase in

specific gravity positively affected on production of ethanol. This strain produced

significantly greater (P<0.05) ethanol, when specific gravity was adjusted to 1.090

(Appendix F9). The production of ethanol at specific gravity of 1.080 was determined

as 11.7% (v/v) that was increased to 13.9% with increase in gravity up to 1.090. No

increase in ethanol yield was observed with further increase in specific gravity beyond

this point. The higher initial specific gravity of fermentation media increased the

osmotic pressure, and hence decreased the ethanol yield. The use of very high gravity

molasses i.e. 1.140 reduced the ethanol yield up to 9.7% (v/v).

In case of strain MZ-4 for fed batch fermentation, ethanol production was increased

from 12.5 to 13.5% (v/v) with the increase in specific gravity from 1.080 to 1.090.

Thus again it was observed that MZ-4 strain produced significantly greater (P<0.05)

ethanol, when specific gravity was adjusted to 1.090 (Appendix F10). The further

increase in specific gravity reduced the final ethanol concentration. Similar to Lalvin

EC-118 strain, the production of ethanol reduced to 9.6% (v/v) by using very high

gravity molasses i.e. 1.140. All these observation showed that 1.090 was the best

specific gravity in case of fed batch fermentation which was easily tolerated by

fermenting microbes and it showed enhanced ethanol production.

4.3.2. Effect of Viscosity

The viscosity of each molasses dilution was determined and it was observed that the

viscosity increases with the increase in amount of molasses, which can create an

unfavorable environment for the growth and productivity of microbial strains. The

viscosity of all the molasses dilutions were determined as mentioned in Table 4.5 and

4.6. The lowest viscosity was determined at the specific gravity of 1.080 as 1.86mP-s,

whereas with the increase in specific gravity an increase in viscosity was also noted.

The maximum viscosity of 4.06mP-s was determined with the highest specific gravity

1.040 used in this study. The result revealed that, high osmotic pressure is not the only

bottleneck of the high gravity fermentation process, but the increase in viscosity was

PhD Thesis


also related to high molasses content that can create an unfavorable environment for

the growth of fermenting microorganisms.

4.3.3. Effect of Feeding Rate

The molasses was fed after various intervals to determine the exact feeding rate for

the maximum production of ethanol at any specific gravity (Fig. 4.9). In case of

Lalvin EC-1118, no regular trend in ethanol production was observed with the

difference in feeding interval; however, in most cases it was observed that more delay

in feeding time decreased the production of ethanol. From the Fig 4.8, it was inferred

that the feeding interval of 12 h at optimized specific gravity i.e. 1.090, the ethanol

content was enhanced up to 13.9% (v/v); however, at the same specific gravity, the

ethanol content reduced to 10.2% (v/v) when feeding was done after every 48 h. The

lowest content of ethanol was determined as 8.3% (v/v) at specific gravity of 1.140 at

feeding rate of 48 h, which showed that the higher specific gravity and delayed

feeding rate adversely affected the production of bioethanol.

In case of strain MZ-4, it was observed that there was no regular trend in increase in

ethanol yield; however, in most cases 24 h was considered as the best feeding interval

to get the enhanced production of ethanol. It was observed that the maximum of

13.5% (v/v) of ethanol was produced when specific gravity was adjusted at optimized

value i.e. 1.090 and fed after every 24 h. It was also observed in most cases that the

delay in feeding time reduced the concentration of ethanol in fermentation medium.

The comparison of both strains showed that the commercial strain Lalvin EC-1118

was more active, and rapidly converted sugars into ethanol; however, strain MZ-4

required more time for the conversion of sugars into ethanol.

Fermentation Efficiency

The fermentation efficiency of the fed batch process at different specific gravity of

molasses with different feeding rate was determined for both strains i.e. Lalvin EC-

1118 and MZ-4. It was observed that higher specific gravity and long feeding

intervals had negative impact on actual ethanol yield, thus reduced the fermentation

PhD Thesis


efficiency of the process. For Lalvin EC-1118, it was observed that maximum ethanol

production was obtained at 1.090 specific gravity with 12 h feeding rate and

maximum fermentation efficiency i.e. 81.1% was noted (Table 4.5). In case of strain

MZ-4, maximum ethanol yield was obtained at specific gravity 1.090 with 24 h

feeding interval, and maximum fermentation efficiency of 83.2% was also observed

under same conditions (Table 4.6).

Process completion time

Process completion time is one of the important factors to determine in case of fed-

batch fermentation as it directly depends on feeding rate and most of the ethanol

industries face the problem of time constraint. The completion time of the process at

different conditions by using strain Lalvin EC-1118 is shown in Table. 4.5. It was

observed that the maximum quantity of ethanol from strain Lalvin EC-118 was 13.9%

with 81.1% fermentation efficiency within 84 h. In case of MZ-4 strain, the process

completion time under different conditions is shown in Table. 4.6. The maximum

amount of ethanol obtained from strain MZ-4 was 13.5% (v/v) exhibiting 83.2%

fermentation efficiency within 120 h.

PhD Thesis


Fig.4.9: The effect of specific gravity and feeding rate for enhanced production of

bioethanol during fed-batch fermentation using strains Lalvin EC-1118 and MZ-4

0

2

4

6

8

10

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14

16

12

hrs

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hrs

.

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hrs

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hrs

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hrs

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hrs

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hrs

.

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hrs

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hrs

.

36

hrs

.

48

hrs

.

1.080 1.090 1.100 1.110 1.120 1.130 1.140

Eth

anl(

v/v)

%

Feeding intervals at different specific gravity

Lalvin EC 118

0

2

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8

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12

hrs

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24

hrs

.

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hrs

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hrs

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hrs

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hrs

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hrs

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hrs

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hrs

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hrs

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hrs

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hrs

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hrs

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hrs

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hrs

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hrs

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hrs

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36

hrs

.

48

hrs

.

12

hrs

.

24

hrs

.

36

hrs

.

48

hrs

.

1.080 1.090 1.100 1.110 1.120 1.130 1.140

Eth

ano

l (v/

v) %

Feeding intervals at different specific gravity

MZ-4

PhD Thesis


Table.4.5: The effect of specific gravity and feeding rate of molasses on fermentation

efficiency and process completion time of fed batch fermentation using Lalvin EC-

1118.

Specific

Gravity

Viscosity

(milliPascal-

second)

Sugar

Conc.

% (w/v)

Feeding

Time

(h)

Actual

Ethanol

%(v/v)

Fermentation

Efficiency

(%)

Process

Completion

time (h)

1.080 1.86 15 12 11.5 75.9 60

1.080 1.86 15 24 11.7 80.4 96

1.080 1.86 15 36 10.3 74.5 120

1.080 1.86 15 48 9.6 64.4 144

1.090 2.13 17 12 13.9 81.1 84

1.090 2.13 17 24 12.5 78.4 120

1.090 2.13 17 36 11.1 71.8 108

1.090 2.13 17 48 10.2 64 144

1.100 2.34 19 12 13 71.1 96

1.100 2.34 19 24 10.9 60 72

1.100 2.34 19 36 11 60.6 108

1.100 2.34 19 48 9.9 53.9 144

1.110 2.86 21 12 10.7 59.7 72

1.110 2.86 21 24 12.2 64 96

1.110 2.86 21 36 11 56.4 108

1.110 2.86 21 48 10.1 50.7 96

1.120 3.20 23 12 11.9 50.5 84

1.120 3.20 23 24 9.4 37.1 96

1.120 3.20 23 36 10.3 44.3 108

1.120 3.20 23 48 10.2 69.0 96

1.130 3.60 25 12 8.6 41.4 96

1.130 3.60 25 24 10.4 51.3 96

1.130 3.60 25 36 10.2 52.1 108

1.130 3.60 25 48 9.9 47.5 96

1.140 4.06 27 12 9.1 42.3 96

1.140 4.06 27 24 9.1 41.0 96

1.140 4.06 27 36 9.7 45.1 108

1.140 4.06 27 48 8.2 36.4 96

PhD Thesis


Table.4.6: The effect of specific gravity and feeding rate of molasses on fermentation

efficiency and process completion time of fed batch fermentation using MZ-4

Specific

Gravity

Viscosity

(milliPascal-

second)

Sugar

Conc. %

(w/v)

Feeding

Time

(h)

Actual

Ethanol

%

(v/v)

Fermentati

on

Efficiency

(%)

Process

Completion

time (h)

1.080 1.86 15 12 12.1 81.4 84

1.080 1.86 15 24 12.5 81.7 96

1.080 1.86 15 36 10.8 67.7 120

1.080 1.86 15 48 9.6 65.4 144

1.090 2.13 17 12 12.5 78.8 84

1.090 2.13 17 24 13.5 83.2 120

1.090 2.13 17 36 10.3 68.6 108

1.090 2.13 17 48 10 67.7 144

1.100 2.34 19 12 12 75.4 60

1.100 2.34 19 24 12.4 72.1 96

1.100 2.34 19 36 11.6 64.8 108

1.100 2.34 19 48 10.6 62.6 144

1.110 2.86 21 12 12 64.2 72

1.110 2.86 21 24 12.6 68.9 96

1.110 2.86 21 36 11.4 61 72

1.110 2.86 21 48 10 52 144

1.120 3.20 23 12 9.2 46.3 84

1.120 3.20 23 24 11.4 59.8 96

1.120 3.20 23 36 10.6 56 108

1.120 3.20 23 48 9.1 45.5 96

1.130 3.60 25 12 10.4 50 72

1.130 3.60 25 24 10.8 53.4 96

1.130 3.60 25 36 9.1 44.1 108

1.130 3.60 25 48 8.3 40.9 96

1.140 4.06 27 12 9.2 41.8 84

1.140 4.06 27 24 9 39.9 96

1.140 4.06 27 36 9.5 44.2 108

1.140 4.06 27 48 8.2 37.3 96

PhD Thesis


PART-B: ENHANCED PRODUCTION OF BIOETHANOL FROM

SUGARCANE BAGASSE

Sugarcane bagasse is the second major type of waste generated by sugar industry.

During this part of study, sugarcane was pretreated by different methods i.e.

autohydrolysis and ionic liquid pretreatment. In next section, the chemical and

structural composition of both types of pretreated bagasse was determined in

comparison to their respective controls. After that, the pretreated bagasse was

subjected to enzymatic hydrolysis with their optimized enzyme loading amount. In

last step, both types of pretreated bagasse was subjected to fermentation by four

different types of yeast strains, to determine the bioethanol produced from each type

of pretreated bagasse.

4.4. BIOMASS PREPARATION

The sugarcane bagasse was dried and milled up to the size of 0.45mm as shown in

Fig.4.10.

4.5. PRETREATMENT OF SUGARCANE BAGASSE

(i) Autohydrolysis

The milled sugarcane bagasse was autohydrolyzed at 190°C for 10 min and 205° C

for 6 min. The bagasse obtained after pretreatment was shown in figure 4.11(a). In the

figure, it could be observed that there was color variation in bagasse pretreated at

various conditions. The increased intensity in color and less porous texture of bagasse

was observed with increase in pretreatment temperature of autohydrolysis.

(ii) Ionic liquid (IL) Pretreatment

The milled sugarcane bagasse was pretreated with IL at 110°C for 30 min. Untreated

bagasse and the bagasse treated with water under same conditions were considered as

control. The bagasse obtained after pretreatment was shown in figure 4.11(b). The

bagasse pretreated with water at 110ºC for 10 minutes showed similarity in color and

texture with the untreated bagasse. However, the IL-pretreated bagasse showed more

dark color that revealed the removal of lignin during pretreatment.

PhD Thesis


Fig.4.10: Drying and milling of sugarcane bagasse through 40 Mesh sieve size: before

drying and milling (left); after drying and milling (right)

Fig.4.11 (a): Sugarcane bagasse autohydrolyzed at 190°C for 10 min; 205°C for 6

min and untreated bagasse control

PhD Thesis


Fig.4.11(b): Sugarcane bagasse, IL pretreated at 110°C for 30 min; water treated at

110°C for 30 min (control) and untreated bagasse (control)

4.5.1. Severity Factor

(i). Autohydrolysis

Sugarcane bagasse was pretreated with autohydrolysis, and the severity of each

pretreatment reaction was determined as shown in Table 4.7(a). The severity factor

determined for this autohydrolysis pretreatment at 190ºC for 10 min and 205ºC for 6

min was 3.64 and 3.86, respectively.

Table 4.7(a): Severity factors for different conditions of autohydrolysis

Pretreatment Temperature Time Severity Factor

Untreated Bagasse - - -

Autohydrolysis 190 °C 10 min. 3.64

Autohydrolysis 205 °C 6 min. 3.86

(ii). Ionic liquid Pretreatment

The severity factor for IL pretreatment was determined and shown in Table 4.7(b).

The severity factor for ionic liquid pretreatment and its water treated control at 110ºC

for 30 min was same i.e. 1.77 because same pretreatment conditions were used to

determine the difference in different pretreatments.

Table 4.7(b): Severity factors for IL pretreatment conditions

Pretreatment Temperature Time Severity Factor

Untreated Bagasse - - -

Water-treated

control

110 °C 30 min. 1.77

Ionic liquid

[C4mim][OAc]

pretreatment

110 °C 30 min. 1.77

PhD Thesis


4.6. COMPOSITIONAL ANALYSIS OF SUGARCANE BAGASSE

4.6.1. Lignin Determination

(i). Autohydrolyzed Bagasse

The effect of autohydrolysis on lignin content of bagasse was shown in Table 4.8(a).

The lignin determination of sugarcane bagasse showed that the total amount of lignin

present in untreated bagasse was 31.9%, with 27.5% of acid insoluble (klason lignin)

and 4.4% of acid soluble lignin. After autohydrolysis of sugarcane bagasse at 190°C

for 10 min, the acid soluble lignin (Klason lignin) was increased to 37.7% which

increased the total lignin content to 41.8%; however, the acid soluble lignin was

reduced to 3.1%. Same trend was observed after autohydrolysis at 205°C for 6 min

that the klason lignin was increased from 27.5 to 36.0% due to which the total lignin

was increased from 31.9 to 39.4%, whereas acid soluble lignin was reduced to 3.4 %.

Table 4.8(a): Lignin Determination of Autohydrolyzed Sugarcane bagasse

Pretreatment Temperature Time Klason

lignin ASL

Total

Lignin

Untreated control - - 27.5±0.2 4.4±0.1 31.9±0.4

Autohydrolysis 190 °C 10 min 37.7±0.3 3.1±0.2 41.8±0.7

Autohydrolysis 205 °C 6 min 36.0±0.4 3.4±0.1 39.4±0.7

ASL= acid soluble lignin

(ii). Ionic liquid Pretreated Bagasse

During current study, the content of lignin was reduced from 31.9% to 28.3% after IL

pretreatment (Table 4.8b). The amount of klason lignin was actually reduced from

27.5 to 21.3% in IL-pretreated sample; however, the overall increase in lignin content

was observed due to increase in acid soluble lignin from 4.4 to 8.0%. The lignin

content in water treated control remained almost same as untreated control. The total

lignin content in water treated bagasse was reduced from 31.9% to 30.9%; and the

PhD Thesis


amount of klason lignin and acid soluble lignin was reduced from 27.5 to 26.8% and

4.4 to 4.1%, respectively, which was not a noticeable difference as compared to IL

pretreated sample.

Table 4.8(b): Lignin determination of IL pretreated sugarcane bagasse

Pretreatment Temp. Time Klason

lignin

ASL Total

Lignin

Untreated Control - - 27.5±0.2 4.4±0.1 31.9±0.4

Water-Treated

Control 110 °C 30 min 26.8±0.2 4.1±0.1 30.9±0.4

IL-pretreated

Bagasse 110 °C 30 min 21.3±0.1 8.0±0.2 28.3±0.4

ASL= acid soluble lignin

4.6.2. Carbohydrate Determination

The carbohydrate content was determined by HPLC and the effect of pretreatments on

the compositional changes was analyzed. The Figure 4.12 is the chromatogram which

showed the peak of glucose, xylose, arabinose and mannose with retention time of 29,

32, 37 and 39 min, respectively. The area of peaks was compared with standard curve

formulated by plotting the graph of different concentration of sugars against area of

defined peak (Appendix C).

PhD Thesis


Fig. 4.12: Chromatogram showing peak of glucose, xylose, arabinose and mannose

standard (with retention time of 29, 32, 37 and 39 min respectively)

PhD Thesis



The effect of different pretreatment strategies on composition of sugarcane bagasse

was shown in Table 4.9(a). It was observed that the glucan content in untreated

bagasse was noted as 37.7%; however, this amount increased to 47.7 and 51.2% in

samples autohydrolyzed at 190°C for 10 min and 205°C for 6 min. The increase in

cellulose content in bagasse might be due to removal of other components. The xylan

content in untreated bagasse was determined as 18.5% which was reduced to 8.2 and

3.8% in autohydrolyzed at 190 and 205 °C respectively. Similar removal of arabinan

was detected during autohydrolysis, and its amount was decreased from 0.1 to 0.04%

in sample autohydrolyzed at 190°C. In the sample autohydrolyzed at 205°C, arabinan

was removed to the extent that it couldn’t be quantified by HPLC. The quantity of

mannan was quantified as 0.004 % only in the sample pretreated at 205°C. In other

samples mannan could not be quantified; therefore, the effect of pretreatment on

mannan changes could not be determined. In addition, no peak for galactan could be

observed in bagasse sample.

Table 4.9(a): Carbohydrate determination of autohydrolyzed sugarcane bagasse

Pretreatme

nt

Temp Time Glucan

%

Xylan

%

Arabinan

%

Mannan

%

Untreated

control

- - 37.7±0.7 18.5±0.3 0.1±0.0 N.Q

Autohydrolysis 190°C 10 min 47.7±0.6 8.2±0.4 0.04±0.1 N.Q

Autohydrolysis 205°C 6 min 51.2±0.3 3.8±0.1 N.Q 0.004±0.0

N.Q=not quantified

(ii). Ionic Liquid Pretreated Bagasse

The carbohydrate content of IL pretreated bagasse in comparison to untreated and

water treated control is shown in Table.4.9 (b). It was observed that the glucan

PhD Thesis


content in untreated bagasses was 37.7% and it was reduced to 37.2% in IL-pretreated

sample. Similarly, in water treated control the difference was quite negligible i.e.

38.5% glucan. The xylan content in IL –pretreated bagasse also reduced from 18.5 to

12.3%; however, the content of xylan was increased in water treated control sample.

The content of arabinan was also increased after pretreatment from 0.1 to 6.07% and

7.0% in case of water treated control and IL pretreated bagasse samples, respectively.

Mannan was quantified as 0.1% in both pretreated samples, but it couldn’t be

quantified in untreated bagasse sample. The galactan was not determined in any of

these samples.

Table.4.9.b: Carbohydrate determination of IL pretreated sugarcane bagasse

Pretreatment Temp Time Glucan

%

Xylan

%

Arabinan

%

Mannan

%

Untreated

control - - 37.7±0.7 18.5±0.3 0.1±0.0 N.Q

Water Treated

Control 110°C 30 min 38.5±0.0 18.9±0.4 6.07±0.1 0.1±0.0

IL-pretreated

bagasse 110°C 30 min 37.2±1.0 12.3±1.0 7.0±0.0 0.1±0.0

4.7. EFFECT OF PRETREATMENT ON BIOMASS STRUCTURE

4.7.1. Fourier Transform Infrared Spectroscopy (FTIR)

Attenuated total reflection-Fourier transform infrared (ATR–FTIR) spectroscopy was

conducted and different absorption bands were used to monitor the chemical changes

of lignin and carbohydrates.

Table 4.10: Assignments of FTIR-ATR absorption bands for bagasse

Wavelength

(cm-1

)

Assignment of Peaks

3336 O-H stretching, related to cellulose-hydrogen bonds

2916 C-H stretching, related to biomass methyl/methylene group

1604 Lignin aromatic ring stretch mode

1635 Ring-conjugated C=C bond in coniferaldehyde

1514 Aromatic skeletal from lignin

1424 CH2 scissor motion in cellulose

1370 C–H stretch in cellulose and hemicellulose

PhD Thesis


1243 Acetylated hemicellulose

1103 Crystalline cellulose

898 Amorphous components

(Adapa et al., 2011; Sun et al., 2015)


The FTIR spectrum of autohydrolyzed bagasse sample is shown in Fig. 4.13(a). The

decrease in spectral intensity at 3300cm-1

showed reduction in cellulose hydrogen

bonding. Two main lignin features were monitored that correspond to lignin aromatic

ring stretch mode (1604 cm-1

) and ring-conjugated C=C bond in conifer aldehyde

(1635 cm-1

), respectively. High temperature autohydrolysis clearly showed

diminished C=C bond in conifer aldehyde, as indicated by peak at 1631 cm-1

, and also

significant modification of the aromatic ring which exhibited the removal of lignin

aromatic ring stretch. Similarly, the significant peaks at 1514 cm-1

(aromatic skeletal

from lignin) were observed for all the autohydrolyzed samples, which might be due to

removal of large content of hemicellulose during autohydrolysis from sugarcane

bagasse sample. A significant peak at 1370 cm-1

(C–H stretch in cellulose and

hemicellulose) in samples obtained after autohydrolysis was likely due to the removal

of major hemicelluloses which comparatively increased the cellulose content in

bagasse, as also determined during compositional analysis. The reduction in peak at

1243 in autohydrolyzed samples showed the reduction in acetylated hemicelluloses.

The peak intensity at 1103 (crystalline cellulose) and peak reduction at 898

(amorphous cellulose) clearly showed the increase in crystallinity of autohydrolyzed

sugarcane bagasse.

(ii) Ionic liquid Pretreated Bagasse

The FTIR spectrum of autohydrolyzed bagasse sample is shown in Fig. 4.13(b). The

increase in spectral intensity at 3300cm-1

in ionic liquid treated bagasse showed

reduction in cellulose hydrogen bonding. The reduction in lignin aromatic ring stretch

mode (1604 cm-1

) and ring-conjugated C=C bond in conifer aldehyde (1635 cm-1

)

showed the removal of lignin component from IL treated sample; however, these two

peaks were increased in water treated control. A significant increase in peak at 1370

cm-1

(C–H stretch in cellulose and hemicellulose) was likely due to the removal of

major hemicelluloses which comparatively increased the cellulose content in bagasse.

The reduction in peak intensity at 1243 cm-1

in IL pretreated bagasse demonstrated the

PhD Thesis


good effect of this pretreatment strategy on removal of acetylated hemicellulose. The

peak at 1103 cm-1

(referring to crystalline cellulose) was diminished after ionic liquid

pretreatment indicated that IL played major role to decrease the cellulose crystallinity.

These results clearly demonstrated that both pretreatments effectively weakened the

van der Waals interaction between cell wall polymers.

4.13 (a): FTIR spectra of untreated, autohydrolysis and IL ([C4mim][OAc]) pretreated

bagasse from 1800-600 cm-1

region

PhD Thesis


4.13 (b): FTIR spectra of untreated, autohydrolysis and IL ([C4mim][OAc])

pretreated bagasse from 4000 to 1800 cm-1

region

4.7.2. XRD Analysis

XRD analysis was performed to examine the cellulose crystallinity index of biomass.

The X-Ray diffraction patterns of pretreated and untreated bagasse were determined

and two diffraction regions were noticed at 22 (2θ) and 19 (2θ) which corresponds to

I002 (crystalline region) and Iam (amorphous region), respectively to calculate the

crystallinity index.


The XRD analysis of autohydrolyzed samples didn’t show any major difference in

peaks at 22° and 15° (Fig 4.14.a). The valley between these two peaks also didn’t show

any remarkable difference among these peaks.

(ii). Ionic Liquid Treated Bagasse

From the Fig. 4.14(b), a remarkable difference in diffraction pattern was observed in

sugarcane bagasse after IL pretreatment. Diffraction pattern of IL pretreated sample

showed the disappearance of secondary peak (I101) at 2θ value of 15 , and the

crystallinity peak (I002) at 22° was also found to be weaken that represented the profile

PhD Thesis


of nearly amorphous cellulose. In addition, the broadening of amorphous valley

between the crystalline and secondary peak was also observed.

PhD Thesis


4.14. XRD analysis of untreated, autohydrolyzed and IL pretreated bagasse

PhD Thesis


Crystallinity Measurement of Autohydrolyzed and IL-Pretreated Bagasse

In order to determine the crystallinity aspect from FTIR analysis, the peak heights

were used as quantitative parameter as stated by Adapa et al. (2011). In order to

determine the crystallinity of cellulose structure, the ratio at the peak intensities at

1424/898 cm-1

was determined, which was referred as an empirical crystallinity index

proposed by Nelson and O’Connor or Lateral order index (LOI) which has been used

to show the presence of cellulose I structure in cellulose material. The ratio of

1370/2916 cm-1,

also proposed by Nelson and O’Connor (1964), known as total

crystallinity index (TCI), was used to evaluate the infrared crystallinity (IR) ratio.

Thus, the higher values of both LOI and TCI are indicative of the more ordered

structure of cellulose and higher crystallinity of biomass. A great reduction in LOI

and TCI of IL-pretreated sample indicated the reduction in crystallinity of cellulose

(Table 4.11). The order of crystallinity determined by LOI was 205ºC > 190ºC >

110ºC > untreated bagasse >IL pretreated bagasse which also corroborated with the

CrI (crystallinity index) determined by XRD analysis (Table 4.11). Crystallinity index

(CrI) of untreated bagasse samples was determined as 0.61, whereas CrI of the

samples autohydrolyzed at 110˚C, 190˚C and 205˚C was 0.62, 0.65 and 0.68,

respectively (Table 4.11). The CrI of IL treated sample was observed as 0.25 which

exhibited reduction in cellulose crystallinity in bagasse.

Table 4.11: Crystallinity measurements of autohydrolyzed and IL pretreated bagasse

Samples FTIR Crystallinity Ratio Crystallinity

Index (CrI)

based on

XRD

Lateral order

index (LOI)

(1424 cm-1

/898

cm-1

)

Total crystallinity

index (TCI)

(1370 cm-1

/2916 cm-1

)

Untreated bagasse 0.47 ±0.0 0.65 ±0.00 0.61

utoh drol sis

for 10 min) 0.78 ±0.03 0.61 ± 0.04 0.65

utoh drol sis

for 6 min) 1.11 ±0.02 0.55 ±0.02 0.68

Ionic liquid (IL) treated

for min 0.24 ±0.01 0.47 ±0.02 0.25

ater treated-control

for min 0.56 ±0.01 0.76 ±0.01 0.62

PhD Thesis


4.8. ENZYMATIC SACCHARIFICATION

4.8.1. Enzyme Loading Optimization


The optimized amount of enzyme that is required by a specific pretreated sample to

convert maximum of carbohydrates in to reducing sugars was determined. This is an

important parameter to reduce the cost of process because this makes it possible to

add only required amount of enzyme for the saccharification process and it would

make the process more economically feasible. For the autohydrolyzed samples the

bagasse treated at 190°C was taken as standard for enzymatic loading optimization

and it was observed that with the increase in enzymatic loading (i.e. cellulases and

beta-glucosidases ratio) the amount of released reducing sugar was also increased.

The addition of 5FPU cellulases with 10IU β-glucosidases released 2.7 mg/ml of the

reducing sugar. When increase concentration of enzyme i.e. 10FPU cellulases with

20IU β-glucosidases was added, the amount of released reducing sugars was increased

up to 3.1 mg/mL; however, no further increase in sugar was observed on further

increase in enzyme loading (Fig.4.15.a). Therefore, 10FPU cellulases with 20IU β-

glucosidases was selected to carry out further experimentations. These results showed

that, 10FPU cellulases with 20IU β-glucosidases were the maximum amount of

enzyme which was required for the maximum conversion of cellulose and

hemicellulose into monomeric reducing sugars.

(ii). Ionic liquid Pretreated Bagasse

It was observed during enzyme loading optimization of IL-pretreated sample that the

addition of 5FPU cellulases with 10IU β-glucosidases released 5.0 mg/ml of the

reducing sugar. When increase concentration of enzyme i.e. 10FPU cellulases with

20IU β-glucosidases was added, the amount of released reducing sugars was increased

up to 5.3 mg/mL. However, enzyamatic hydrolysis of IL-pretreated sample with

20FPU of cellulases with 40IU of β-glucosidases released maximum of 6mg/mL of

reducing sugars, which was detected by DNS method. No further increase in reducing

sugar was observed with further increase in enzyme loading (Fig.4.15.b). An

interesting observation was that, IL pretreated sample required more enzyme loading

and released more amount of sugar as compared to autohydrolyzed sample.

PhD Thesis


Fig.4.15(a): Enzyme loading optimization for autohydrolyzed samples

Fig.4.15 (b): Enzyme loading optimization for IL pretreated samples

0

0.5

1

1.5

2

2.5

3

3.5

0 24 hrs 48 hrs 72 hrs

Re

du

cin

g su

gar

(mg/

ml)

Time (h)

5FPU Cellulases : 10IU B-glucosidases

10FPU Cellulases:20IU B-glucosidases

20FPU Cellulases.:40IUB-glucosidase

0

1

2

3

4

5

6

7

0 24 48 72

red

uci

ng

suga

r (m

g/m

l)

Time (hrs)

5FFPU Cellulase:10IUB-glucosidases 10FPU Cellulases:20IU B-glucosidases

20FPU Cellulase.:40IU B-glucosidase 40FPU Cellulase: 80IU B-glucosidase

PhD Thesis


4.8.2. Enzymatic Hydrolysis

Fig.4.16: Chromatogram of sugarcane bagasse hydrolysate obtained after enzymatic

hydrolysis

PhD Thesis


During enzymatic hydrolysis, the release of glucose and xylose were determined by

HPLC. The chromatogram showed the glucose and xylose peaks at retention time of

29 and 32min respectively (Fig 4.16). The area of peaks was compared with standard

curve formulated by plotting the graph of different concentration of sugars against

areas of defined peak (Appendices C).

(i) Autohydrolyzed Bagasse

The variation in glucose concentration released by enzymatic hydrolysis of sugarcane

bagasse pretreated with different strategies was shown in Fig. 4.17.a and Appendix

E.1.1. It was observed that, from an untreated bagasse 0.47mg/ml of glucose was

released after 72 h of enzymatic hydrolysis; however, when the bagasse was

pretreated at 190ºC for 10 min and 205ºC for 6 min, the glucose concentration was

noted as 2.3 mg/ml and 3.5 mg/ml, respectively. These findings showed that the

release of glucose from bagasse pretreated at 205°C was significantly higher (P<0.05)

than other autohydrolyzed bagasse samples (Appendix G1). The amount of xylose

released after enzymatic hydrolysis of various autohydrolyzed was shown in Fig.

4.17.b and Appendix E.1.3. The released xylose concentration from bagasse

pretreated at 190°C was significantly greater (P<0.05) than other autohydrolyzed

samples (Appendix G2). It was determined that the release of xylose from untreated

bagasse was 0.03mg/ml; however, the autohydrolyzed bagasse at 190°C for 10 min

and 205°C for 6 min released 0.4mg/ml and 0.24mg/ml of xylose, respectively.

(a).Cellulose Digestibility (%)

The cellulose digestibility of autohydrolyzed and untreated bagasse was determined

by dividing the amount of cellulose digested by total amount of cellulose present in

pretreated sample. It was observed that the autohydrolyzed samples at 190ºC for 10

min and 205ºC for 6 min showed cellulose digestibility of 46.9% and 62.1% after 72 h

of enzymatic hydrolysis; however, untreated bagasse showed cellulose digestibility of

only 11.4% (Fig. 4.18.a, Appendix E.1.2).

(b). Xylan Digestibility (%)

The hydrolytic process of autohydrolyzed samples was completed at 72 h and xylan

digestion was determined by dividing the amount of xylan digested over the total

amount of xylan in pretreated sample. The figure 4.8.b showed that 4.6% and 5.7%

xylan digestibility was noted in samples autohydrolyzed at 190ºC and 205ºC,

respectively; however, only 1.4% xylan digestibility was determined in untreated

bagasse sample (Appendix E.1.4).

PhD Thesis


Fig.4.17.a. Glucose concentration released from autohydrolyzed samples during


Fig.4.17(b): Xylose concentration released from autohydrolyzed samples during


0

0.5

1

1.5

2

2.5

3

3.5

4

0 24 48 72

glu

cose

(m

g/m

l)

Time (hrs) Untreated ControlAutohydrolyzed 190 C-10 minAutohydrolyzed 205 C-6 min

0

0.1

0.2

0.3

0.4

0.5

0.6

0 24 48 72

Xyl

ose

(m

g/m

l)

Time (hrs)

Autohydrolyzed 190 C-10 min


Untreated Control

PhD Thesis


Fig.4.18 (a): Cellulose digestibility from autohydrolyzed samples during enzymatic

hydrolysis

Fig 4.18 (b): Xylan digestibility from autohydrolyzed samples during enzymatic

hydrolysis

0

10

20

30

40

50

60

70

0 24 48 72

Ce

llulo

se d

ige

stib

ility

pe

rce

nta

ge

Time (hrs) Untreated Control



0

1

2

3

4

5

6

7

0 24 48 72

Xyl

an D

ige

stib

lity

(%)

Time (hrs)

Untreated Control



PhD Thesis


(ii) Ionic liquid Pretreated Bagasse

The variation in glucose and xylose concentration released by enzymatic hydrolysis of

sugarcane bagasse pretreated with IL was shown in Fig 4.19.a (Appendix E.2.1).

Enzymatic saccharification of the IL pretreated bagasse (at 110ºC for 30 min) released

4.0 mg/ml glucose, untreated and water treated control showed the release of 0.47 and

0.8mg/ml of glucose respectively after 72 h of enzymatic hydrolysis. The release of

xylose from IL-pretreated sample was determined as 0.4 mg/ml; however the release

of xylose from water treated control and untreated control was determined as 0.1 and

0.03 mg/ml after 72 h of enzymatic hydrolysis (Fig 4.19.b, Appendix E.2.3). IL-

pretreated samples showed significantly greater (P<.005) released glucose and xylose

concentration than all other pretreated samples (Appendix G1 and G2). It was also

observed that the enzymatic hydrolysis of IL-pretreated sample was quite fast and it

released 3.3 mg/ml of glucose within 3 h after the start of hydrolysis.

(a). Cellulose Digestibility (%)

It was determined that the glucose release from IL-pretreated sample was more rapid;

therefore, the cellulose digestibility after 3 h of enzymatic hydrolysis was determined.

The cellulose digestibility of 79.8% was determined after 3 h; however, for the

untreated control and water treated control the digestibility (%) after 3 h was noted as

15.6% and 8.09%, respectively. The hydrolytic process was completed after 72 h and

cellulose digestibility of 11.3, 19.3 and 97.4% was determined from hydrolysis of the

untreated control, water treated control and IL-pretreated bagasse sample (Fig 4.20.a,

Appendix E2.2).

(b). Xylan Digestibility (%)

Xylan digestibility of IL pretreated sample was reported as 65.1% after 3 h, whereas

for water treated sample at 110ºC and untreated bagasse sample, the xylan

digestibility percentages were noted as only 4.2 and 0.1 % respectively (Fig 4.20.b,

Appendix E.2.4). The hydrolytic process completed at 72 h and hemicellulose

digestion was determined as 98.6% from IL pretreated bagasse while 8.3% and 1.4%

respectively hemicellulose digestibility was noted from water treated control and

untreated control samples respectively. Enzymatic hydrolysis results clearly showed

that soluble sugars were released faster to a greater extent when an IL pretreatment of

sugarcane bagasse was used rather than the autohydrolysis pretreatment. It was also

observed that total processing time to reach 60% cellulose digestibility was about 48 h

with autohydrolysis but it was reached up to 79.8% within 3 h with IL pretreatment.

PhD Thesis


Fig. 4.19.a. Glucose concentration released from IL pretreated samples during


Fig.4.19.b. Xylose concentration released from IL pretreated samples during


0

0.5

1

1.5

2

2.5

3

3.5

4

4.5

0 24 48 72Glu

cose

co

nce

ntr

atio

n (

mg/

ml)


Water-treated control (110 C-30 min)

IL-treated control (110 C-30 min)

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

0 10 20 30 40 50 60 70 80

Xyl

ose

co

nce

ntr

atio

n (

mg/

ml)



IL-treated control (110C-30 min)

PhD Thesis


Fig. 4.20 (a): Cellulose digestibility from IL pretreated samples during enzymatic

hydrolysis

Fig. 4.20 (b): Xylan digestibility from IL pretreated samples during enzymatic

hydrolysis

0

10

20

30

40

50

60

70

80

90

100

0 12 24 36 48 60 72

Ce

llulo

se D

ige

stib

ility

(%

)

Time (hrs)

Untreated Control


IL-treated control (110 C-30 min)

0

20

40

60

80

100

120

0 10 20 30 40 50 60 70 80

Xyl

an d

ige

stib

ilty

(%)


Water treated control

IL treated

PhD Thesis


FERMENTATION

Ethanol production from fermentation of pretreated/enzymatically deconstructed

samples of sugarcane bagasse showed that different microbial strains have different

abilities to produce ethanol from the available sugars.


Ethanol production obtained from bagasse autohydrolyzed at 190ºC for 10 min was

66.02 mg/g-substrate when it was fermented with S. cerevisiae (Lalvin EC-118);

whereas, bagasse pretreated at 205ºC showed maximum ethanol production (70.9

mg/g-substrate) when S. cerevisiae (MZ-4) was used as fermenting organism. Only

15.5 mg/g-ethanol was released when enzymatic hydrolysis were carried out with an

untreated bagasse and then subjected to fermentation process by using MZ-4 strain

(Fig 4.21.a, Appendix E.3).

(ii) Ionic Liquid Pretreated Bagasse

Best ethanol production (78.8 mg/g-substrate) was obtained when sugarcane bagasse

was pretreated with IL ([C4mim][OAc]) at 110ºC for 30 min and fermented with S.

cerevisiae (MZ-4) strain (Fig 4.21.b, Appendix E.3). This strain can be considered as

the most tolerant strain in [C4mim][OAc] pretreatment conditions and played

important role in production of maximum ethanol. The maximum ethanol production

obtained from water treated bagasse at 110ºC for 30 min was 28.42mg/g-substrate

when it was fermented with S. cerevisiae (Lalvin EC-118). Pitchia stipites was

considered as more efficient to ferment xylose into ethanol but it showed less

production with all pretreated samples. During fermentation process, IL-pretreated

samples showed significantly greater (P<0.05) production of ethanol than all

pretreatment strategies; however, significance was not observed between IL-

pretreated and samples autohydrolyzed at 205°C (Appendix G3).

PhD Thesis


Fig.4.21: Production of Bioethanol from untreated, autohydrolyzed and IL pretreated

sugarcane bagasse

0

10

20

30

40

50

60

70

80

90

Untreatedcontrol

Water treatedcontrol 110 C

Ionic liquid110C

Autohydrolysis190 C

Autohydrolysis205 C

Eth

ano

l (m

g/g-

sub

stra

te)

Pretreatment Strategies

Uvaferm-43 Lalvin EC-118 MZ-4 Pitchia stipilis

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Chapter 05

Discussion

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Bioethanol is a type of liquid fuel which can easily be produced from sugar rich

materials and mainly used in substituent to gasoline. The main focus of this study was

to enhance the production of bioethanol from sugar industry waste. There were two

main types of sugar industry wastes that were utilized in this study for the bioethanol

production i.e. sugarcane molasses and sugarcane bagasse. Various fermenting yeasts

were tested under optimized physicochemical parameters and various pretreatment

strategies were employed (specifically for sugarcane bagasse) to determine more

effective conditions and microbial strain which are required to enhance the bioethanol

yield from sugar industry waste.

PART A: BIOFUEL PRODUCTION FROM SUGARCANE MOLASSES

5.1. Physicochemical Analysis of Sugarcane Molasses

The physicochemical properties of molasses vary along with the source and type of

molasses. During current study, various physicochemical properties i.e. sugar content,

brix, specific gravity was determined. Curtin et al., (1983) suggested that the large

variation in total sugar or carbohydrate content was observed in molasses from

various sources and types. The sugar content of molasses in current study was

determined as 49% with 15% reducing and 32.3% non-reducing sugar. Previously it

was believed that the sugar mill could control the sucrose content depending up on the

production technology employed by the sugar industry. It was also observed that the

centrifugation process which helped in separation of sugar and water played an

important role in determining the content of sugar present in molasses (Curtin et al.,

1983). The term brix was originated for a pure sugar (specifically sucrose) solution;

however, for the molasses this term is used for total solid content due to the presence

of various types of sugars i.e. fructose, glucose, raffinose and many non- sugar

organic contents (Curtin, 1983). The brix of molasses during current research was

noted as 79 °brix. Curtin et al., (1983) suggested that the term brix only represent a

number related to specific gravity rather than dry matter content or sucrose. Specific

gravity can be defined as the ratio of the density of a substance to the density of

reference material i.e. water. In current study, the specific gravity was noted with the

help of gravity hydrometer and was found 1.40; however, in some previous studies,

the specific gravity was determined as 1.43 for a test molasses (Olbrich, 1963).

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5.2. Isolation, Screening and Characterization of Indigenous Yeast

During pioneer studies on fruit ecology, the majority of yeasts were isolated from

fresh grapes (Fleet et al., 2002). In our current study, the isolation of yeast strains was

carried out from various fruits and soil samples. Twenty five yeast strains were

isolated and screened for the ethanol tolerance, and one of the strains, which was

isolated from grapes showed maximum of 15% (v/v) ethanol tolerance and labeled as

MZ-4. Strain MZ-4 was identified through morphological and molecular methods

which showed 100% resemblance with S. cerevisiae strain S288C. Previously Li et

al., (2010) worked on varieties of the yeasts associated with grapes. The yeast strains

identified in grapes were Sporidiobolus pararoseus, Cryptococcus carnescens,

Cryptococcus flavescens, Candida inconpicua, Candida quercitrusa, Cryptococcus

magnus, Hanseniaspora guilliermondii and Zygosaccharomyces fermentat. Various

studies on grapes microbial ecology showed that the most common yeast found in

healthy grapes were Saccharomyces cerevisiae (Fleet et al., 2002; Barata et al., 2012).

Cabrera et al., (1988) worked on five different yeast strains and the best yeast for the

production of ethanol was determined as S. cerevisiae (Cabrera et al., 1988).

Different types of yeast strains showed variation in their ethanol tolerance ability, and

their intolerance against high concentration of ethanol affect the productivity of the

bioethanol (Lam et al., 2014). In present study, MZ-4 strain was tolerant up to 15%

ethanol; therefore, this strain was selected for further experimentations to enhance the

production of bioethanol. The increase in ethanol tolerance among various strains can

be attributed to the increase in proportion of unsaturated fatty acids, ergosterol and the

biosynthesis of phospholipids with in cytosol and membrane structures. It was

suggested that the decrease in sterol, protein ratio and phospholipids enhanced the

plasma membrane fluidity, thus adversely affected the ethanol tolerance of microbial

strain (Alexandre et al., 1994; Dinh et al., 2008). Basso et al., (2011) reported that the

increase in ethanol content in fermentation medium might decrease the number of

living cells by affecting their enzyme activity and growth of fermenting microbe

which in turn reduced the final ethanol yield.

5.3. Comparison of Commercial Strains with Indigenous Strain MZ-4

In order to compare the newly isolated strain MZ-4 with the best available

commercial strains, four commercial strains were selected i.e. Rossmoor, Saf-gold,

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Uvaferm-43 and Lalvin EC-1118. Two main problems faced by fermenting microbes

were osmotic and ethanol intolerance (Basso et al., 2011); therefore, the best ethanol

tolerant and osmotic tolerant strain was selected for comparison. Ethanol tolerance for

the commercial strains from already published data was found as 6, 12, 18 and 18%

(v/v) respectively (Bechem et al., 2007; Schmidt et al., 2011; Sultana et al., 2013).

For the selection of most efficient strain, osmotic tolerance of all the commercial

strains and indigenous MZ-4 strain was also determined. MZ-4 showed the best

osmotic tolerance among all the five strains and Lalvin EC-1118 was the most

efficient among industrial strains only. Lalvin EC-1118 and MZ-4 showed best

ethanol production of 9.6 and 10.1% (v/v) respectively in the presence of 25% (w/v)

sugar in molasses dilutions. It was observed that the increase of sugar concentration in

fermentation medium had positive effect on increase of ethanol production. However,

there was certain limit up to which any strain could tolerate the sugar concentrations,

beyond which the ethanol yield started decreasing. These findings justified a previous

study which revealed that the increase in sugar concentration enhanced the production

of ethanol up to 5.3% (w/v) from 30% (w/v) of the sugar concentration in

fermentation medium (Periyasamy et al., 2009). Arshad et al., (2008) during his study

determined the production of 7.7% (v/v) of ethanol in presence of 16% (w/v) sugar in

the fermentation medium. Many previous studies reported that 15% (w/v) was the

optimum sugar concentration for the maximum production of bioethanol (Amutha and

Gunasekaran, 2001; Alegre et al., 2003; Mariam et al., 2009).

The determination of fermentation efficiency at different sugar concentration revealed

that, yeast strains were not able to utilize sugar completely (i.e. 100% fermentation

efficiency) at increased sugar concentration. It was observed, at 25% (w/v) of sugar

concentration, Lalvin EC-1118 and MZ-4 strain converted sugar in to bioethanol up to

60 and 63.1% (w/v), respectively. This study confirms the previous studies that

showed the decrease in fermentation efficiency with an increase in sugar

concentration. Fadel et al., (2013) and Nofemele et al., (2012) reported 75% and 76%

of fermentation efficiency with actual ethanol yields of 8.4 and 7.6% (v/v)

respectively.

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Two strains i.e. Lalvin EC-1118 (commercial strain) and MZ-4 (indigenous strain)

were selected on the basis of their best ethanol and osmotic tolerance and the

physicochemical conditions of both strains were optimized to determine the maximum

ethanol could be produced by these strains. The effect of pH on fermentation process

was discussed by Methewson et al., (1980) during his study that slightly low pH not

only promote the growth of fermenting yeast, but also retard the growth of

contaminating lactic acid bacteria. During current study, the effect of pH on enhanced

production of bioethanol was studied which showed that pH 4.5 and 5.0 were best for

the strains Lalvin EC-1118 and MZ-4, respectively. The optimum pH of both strain

lies between the range 4.0 to 5.0 as already suggested in previous studies (Patrascu et

al., 2009; Periyasamy et al., 2009; Mukhtar et al., 2010). In various studies, it was

observed that the increase in pH of fermentation medium degraded the ethanol into

various organic acids and glycerol, thus reduced the ethanol production; however, the

increase of pH up to 7.0 enhanced the production of glycerol and increased the

activity of aldehyde dehydrogenase enzyme which favored the conversion of

acetaldehyde in to acetic acid, thus reduced the production of bioethanol (Wang et al.,

2001).

It has been widely studied that the temperature has crucial effect on yeast growth and

fermentation process (Laluce et al., 1993; Phisalaphong et al., 2006; Periyasamy et

al., 2009). In current study, the effect of temperature on both strains was studied to

determine their optimum temperature and it was observed that Lalvin EC-1118 and

MZ-4 showed maximum ethanol production at 30 and 33°C, respectively. The

previous studies on ethanol production from S. cerevisiae also indicated that the best

temperature for the yeast growth and production of bioethanol ranges 30 to 35°C

(Periyasamy et al., 2009; Mukhtar et al., 2010). Different studies revealed that the

increase in temperature exhibited adverse effect on enzyme’s catalytic activity, and at

very high temperature cellular proteins were disrupted and denatured which affected

the growth of yeast cells structures (Mariam et al., 2009; Dhaliwal et al., 2011;

Nofemele et al., 2012).

The effect of inoculum size and age on production of bioethanol was also determined

in current study, and it was observed that the strain Lalvin EC-1118 and MZ-4

produced maximum of ethanol when 7.5 and 10% (v/v) of 24 h old culture

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(containing 3x108

live cells per ml) were inoculated, respectively. These results

confirmed the findings of previous researches demonstrating that the inoculum size

and age directly affect final ethanol yield. It was determined by Munene et al., (2002)

that the inoculum size of 7×106 viable count/ml yielded maximum ethanol with

minimum byproducts i.e. glycerol. Similar observations were reported by Laopaiboon

et al., (2007) that 1x108

cells/ml was the optimized inoculum size for maximum

ethanol production. Later, Perisyasmi et al., (2009) during his study on S. cerevisiae

revealed that 2 g of yeast inoculum exhibited maximum production of bioethanol.

Further studies carried by Benerji et al., (2010) determined that 1.5% (v/v) of 48 h old

inoculum was best for the maximum production of bioethanol.

During early times, it was considered that the fermentation process is not affected by

the addition of nitrogen source; however, with the advent of researches it was found

that the efficiency of fermenting microbes are greatly affected by various nutrient

supplements (Casey et al., 1983). The majority of researchers, who studied the effect

of nitrogen sources on the production of bioethanol, determined that the change in

amount of nitrogen in fermentation medium enhanced ethanol yield by altering the

activity of enzymes involved in fermentation (Thomas et al., 1996; Yalçin and Özbas,

2004). During current study, various nitrogenous sources i.e. urea, ammonium

chloride (NH4Cl), ammonium nitrate (NH4NO3), ammonium sulfate ((NH4)2SO4) and

di-ammonium phosphate ((NH4)2HPO4) were added in fermentation medium in range

0.0-0.15% (w/v) to compare the effectiveness of these compounds as compared to

urea. The best nitrogen source for Lalvin EC-1118 and MZ-4 was determined as 0.1%

(w/v) of (NH4)2HPO4 and NH4Cl, respectively. These results verified the effect of

various nitrogen sources reported in previous studies; however, the type and quantity

of the nitrogen source depends on fermenting strain and source of substrate.

Previously, Bafrncová et al., (1999) studied that the addition of urea as nitrogen

source played an important role to enhance the tolerance of fermenting strain against

high concentration of ethanol. Similarly, Mukhtar et al., (2008) has been studied the

effect of urea and (NH4)2HPO4 and showed enhancement of bioethanol production by

the addition of both of these nitrogenous sources. Nofemele et al., (2012) inferred

from his research that 2 g/L of urea was best to enhance the ethanol yield up to

maximum limit. Another study conducted by Maharjan et al., (2012) revealed that

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ethanol yield increased up to 7.3 mg/ml by the addition of di-ammonium phosphate at

concentration of 1mg/ml.

The presence of various heavy metals in molasses e.g. Zn, Cu, Fe and Mn plays an

important role in growth of yeast cells but their higher concentrations can have

inhibitory effects. Due to this reason, the use of various chelating agents improves the

fermentation product by binding to toxic metals present in crude fermentation

medium (Pandey and Agarwal, 1993). During current study, various chelating agents

i.e. EDTA, K4Fe(CN)6, and NaK-Tartrate were added in to fermentation medium in

concentration range 0.0025-0.325% (w/v) to determine their effect on production of

bioethanol. It was observed that Lalvin EC-1118 and MZ-4 produced maximum

amount of ethanol in the presence of 0.04% and 0.01% (w/v) of K4Fe(CN)6,

respectively. The current study confirmed the findings reported in previous studies

that addition of chelating agents had positive effect on enhanced production of

bioethanol. Panday and Agarwal (1993) compared the effect of EDTA, K4Fe(CN)6,

and NaK-Tartrate and determined that the addition of EDTA as chelating agent at its

optimal dose i.e. 0.05 g/L showed maximum production of bioethanol. Similar results

were reported by Benerji et al., (2010), who found that addition of 1.2 g/L NaK-

Tartrate enhanced the production of bioethanol up to 12.0% (v/v). In a different study

conducted by Yadav et al., (1997) maximum production of ethanol was also observed

by the addition of 250 ppm of K4Fe(CN)6, and it reduced content of Fe from 85 to 47

ppm, and Cu from 7.3 to 5.4 ppm.

5.4. Fed Batch Fermentation

Different methods have been proposed for the production of bioethanol from

sugarcane molasses i.e. batch, fed batch and continuous fermentation (Yoshida et al.,

1973). The batch fermentation was the most common method in brewing industry for

the production of bioethanol. One of the major bottlenecks of batch fermentation is

the intolerance of fermenting microbes against high sugar concentration present in

molasses. Increase in ethanol production can be observed by using higher sugar

concentration up to certain limit, beyond which it hinders the growth of fermenting

microbes thus stopped fermentation process (Cheng et al., 2009). In present study, the

fed-batch fermentation was also employed to compare the difference in ethanol yield

by using batch or fed-batch fermentation. Fed-batch fermentation processes are

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widely applied to reduce substrate inhibition to achieve high productivity. During fed

batch fermentation, substrate is added after regular interval to reduce its inhibitory

effect and enhance the productivity without the removal of fermentation broth (Hunag

et al., 2012). The process is more successful in terms of high ethanol yield with fast

saccharification rate, decreased substrate inhibition and process completion time

(Cheng et al., 2009). Optimization of fed batch process is quite challenging because

of some physical constrained and non-linear dynamic equations which governed the

process (Hunag et al., 2012). In present study, enhanced production of bioethanol was

studied employing fed batch fermentation while using Lalvin EC-1118 and MZ-4 as

fermenting strains. The results of fed-batch fermentation were quite interesting as

compared to batch fermentation. It was observed that Lalvin EC-1118 strain produced

13.9% (v/v) of ethanol at 1.090 sp. grv. (17% (w/v) sugar) when molasses was fed

after every 12 h, whereas MZ-4 strain produced maximum ethanol of 13.5% (v/v) at

same sugar concentration but the optimum feeding time for this strain was 24 h. It was

observed during the study that more delay in feeding time and higher initial specific

gravity also reduced the final ethanol production. The reduction in ethanol yield with

increased specific gravity of molasses can be attributed to high osmotic pressure

created in fermentation medium with increase in sugar concentration which affected

the growth of yeast cells and also fermentation process (Cheng et al., 2009). Roukas

et al. (1991) illustrated that the increase in sugar concentration can cause plasmolysis

by decreasing water activity, thus reduced the rate of fermentation. In previous

studies, Laopaiboon et al., (2007) has been reported 120 g/L of ethanol obtained after

fed batch fermentations. Hunag et al., (2012) worked to optimize feeding rate, feeding

time, feeding glucose of a fed batch fermentation process which depicted an enhanced

productivity by 4.4%. Similarly, Bideaux et al. (2006) has been shown that high

performance fed batch fermentation enhanced the ethanol yield up to 0.34 g/g when

700 g/L of glucose solution was fed to fermenter and also helped in minimizing the

glycerol production during the process.

The comparison of fermentation efficiency showed that the fermentation efficiency of

the process increased to 81.1% when Lalvin EC-1118 strain was used for

fermentation; however, in case of MZ-4 the fermentation efficiency was increased to

83.2%. An increase in fermentation efficiency during fed batch fermentation showed

that the step wise addition of molasses helped microbes to avoid the problems of

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osmotic intolerance, and it efficiently converted majority of available sugar into

bioethanol. In comparison, MZ-4 showed better fermentation than Lalvin EC-1118

strain which can be beneficial in terms of utilizing maximum of sugar to enhance

production of bioethanol. Li et al., (2012) showed an increase in fermentation

efficiency up to 90% by employing fed batch fermentation using E. coli as fermenting

microbe.

During evaluation of process completion time, it was noticed that the time required to

complete fed batch fermentation by using Lalvin EC-1118 and MZ-4 strain under

optimized condition was 86 h and 120 h, respectively. The industries which are more

concerned with time constraints and distillation cost should prefer Lalvin EC-1118

strain over MZ-4 because it showed rapid conversion of sugar in to bioethanol and

also ethanol concentration was much higher in case of utilizing former strain. In

contrast, those industries, which are much concerned about their fermentation

efficiency and prefer to convert maximum of the sugar into bioethanol should prefer

MZ-4 strain because of its better fermentation efficiency.

In Pakistan, the previous research on enhanced production of bioethanol by

employing fed batch fermentation was carried out by Mukhtar et al., (2009) and he

determined the production of 8.3% (v/v) of bioethanol under optimized condition.

During current study, it was observed that the optimization of process enhanced the

ethanol production up to 10% (v/v); whereas the process shifting to fed-batch

fermentation utilizing same microbial strains enhanced the ethanol production up to

13.9%. Moreover, the fed batch fermentation process enhanced fermentation

efficiency and reduced process completion time. Therefore, it can be suggested that

fed batch fermentation is more efficient and economic process as compared to batch

fermentation and it should be adopted by sugar industries in Pakistan to get more

economic benefit.

PART B: BIOFUEL PRODUCTION FROM SUGARCANE BAGASSE

5.5. Pretreatments of Sugarcane

This study investigated various pretreatment strategies i.e. autohydrolysis and IL

([C4mim][OAc]) pretreatment of sugarcane bagasse, and comparatively evaluated

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more appropriate technique to increase enzymatic digestibility and bioethanol

production. Studies on various alkylimidazolium salts reveal that shorter alkyl chain

of [C2mim]+ imparts greater extent of saccharification with faster dissolution.

However, higher dissolution extent of [C2mim]+ does not benefit overall process of

pretreatment since losses in [C4mim]+-treated biomass were much less as compared to

[C2mim]+ pretreatment process. In terms of hemicellulose saccharification yield,

[C4mim]+ ILs perform better as hemicellulose is preserved in its polymeric and

recovered form after pretreatment (Karatzos et al., 2012). Due to these reasons,

[C4mim][OAc] (1- butyl-3-methyl imidazolium acetate) pretreatment was selected for

this study and compared with high temperature autohydrolysis.

5.6. Compositional Analysis of Sugarcane Bagasse

The chemical composition of untreated sugarcane bagasse was 37.7% glucan, 18.5%

xylan and 31.9% lignin that were comparable to previous reports (Pitarelo, 2007; Qiu

et al., 2012; Batalha et al., 2015). During autohydrolysis, hemicelluloses were

converted into soluble oligomers and monomers which degraded the intact

lignocellulosic structure. The release of hemicellulose also accompanied with partial

release of lignin fraction (mainly acid soluble lignin) (Lee et al., 2009). Comparative

to ionic liquids, autohydrolysis has shown more promising effect on dissolution of

hemicelluloses. It was inferred from the results that maximum xylan dissolution was

observed when autohydrolysis was carried out at high temperature condition i.e.

205ºC for 6 min. In comparison to autohydrolysis, ionic liquid exhibited less effect on

dissolution of hemicelluloses. Since hemicellulose forms a physical barrier which

prevents compact cellulose structure from enzymatic attack; autohydrolysis has been

considered as a good method to remove this barrier, thereby increasing enzymatic

cellulose digestion. Other studies also indicated that the degree of xylan dissolution

depends on severity of pretreatment condition (Boussarsar et al., 2009).

A significant increase in lignin content was observed in biomass after autohydrolysis

which might be attributed to the removal of significant amount of hemicellulose and

certain amount of cellulose while retaining most of the lignin (Qiu et al., 2012;

Vallejos et al., 2012; HU, 2014). Some researchers suggested that the increased lignin

content might be due to the repolymerization of polysaccharides degradation product

(such as furfural) and/or polymerization with lignin, which forms a lignin like

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material termed as pseudo-lignin (Li et al., 2007). The pseudo-lignin can also be

generated from carbohydrate without significant contribution from lignin, especially

under high severity pretreatment conditions (Sannigrahi et al., 2011). Previous

researches have shown that the major impacts of autohydrolysis on lignin were its

translocations and redistributions, resulting in formation of lignin droplets of various

morphologies on cell wall. The pretreatment that exceed critical phase transition

temperature of lignin allow it to expand and migrate to larger void, where it is

reshaped by aqueous environment into spherical droplet (Donohoe et al., 2008).

Lignin droplet can leave the cell matrix, move into solution and are relocated to the

surface of biomass. In addition, some of the dissolved lignin is re-precipitated onto

the surface of biomass, especially in a batch reactor, as the system is cooled (Jung et

al., 2010; HU, 2014; Pu et al., 2015).

During our current studies, the IL pretreatment lead to greater dissolution and removal

of lignin in bagasse than what was accomplished under comparable autohydrolysis

conditions. The dark brown color of the IL-treated sample, just after the onset of

reaction, showed its excellent ability to extract lignin from bagasse (Sant'Ana da Silva

et al., 2011). It has been previously reported that the ionic liquid cation interacts with

lignin through Π-Π interaction to help in lignin dissolution; however, complete

dissolution of lignin was difficult due to location of lignin within lignin–carbohydrate

complex and hydrophobicity (Qiu et al., 2012). Ionic liquid exhibited better effect on

lignin dissolution as compared to autohydrolysis thus assisting in enzymatic

accessibility but both pretreatment methods has limited effect on cellulose removal

because of its highly packed crystalline structures which are resistant to high

temperature (Qiu et al., 2012; Vallejos et al., 2012; Qiu and Aita, 2013). The increase

in cellulose content after autohydrolysis was attributed to the significant removal of

hemicelluloses. The effect of both pretreatments on lignocellulosics conversion was

further studied by FTIR and XRD analysis.

5.7. Structural Analysis of Sugarcane Bagasse

Attenuated total reflection-Fourier transform infrared spectroscopy (ATR–FTIR) of

untreated and pretreated bagasse was carried out. Previous studies showed that the

peaks at 3175-3490 cm-1

showed O-H stretching intra-molecular hydrogen bonds for

cellulose I (Kumar et al., 2014; Poletto et al., 2014; Sun et al., 2014). In present

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study, a reduction of this peak in autohydrolyzed samples showed the effectiveness of

severity on OH-stretching molecules. The band intensity at 3336 cm-1

after water

treated sample at 110°C represented cellulose hydrogen bonding and possible co-

crystallization due to pretreatment; however, comparatively lower absorption band

was observed from [C4mim][OAc] pretreated samples at same conditions. The peaks

associated to lignin structure are 1604 and 1510 cm-1

(Adapa et al., 2011; Kumar et

al., 2014), which showed during current study a remarkable decrease in lignin content

in autohydrolyzed and IL pretreated sample; however, some of the autohydrolyzed

samples showed an increase in peak which might be due to removal of large amount

of hemicelluloses which in turn increased the lignin content. Similar results were

reported by Kartoz et al., (2012) who demonstrated that the bagasse treated with

[C2mim][OAc] showed reduction in these peaks as compared to untreated bagasse.

During compositional analysis, an increase in lignin content was observed due to

removal of large amount of hemicellulose; however, the reduction in these specific

FTIR peaks cleared that some of the lignin structure was also removed during

pretreatment despite of its increase in overall composition. The band positions at 1730

and 1243 cm-1

are assigned to acetyl group of hemicelluloses (Adapa et al., 2011;

Kumar et al., 2014; Sun et al., 2014). During current study, the reduction in these

bands in all pretreated samples showed the removal of hemicelluloses component

from sugarcane bagasse. Similar results were found by Sun et al., (2014), which

showed the effect of various pretreatment strategies on removal of hemicellulose

associated peaks. Similarly, the reduction in peak at 1160 cm-1

was also observed

after both pretreatment during current study which was associated with hemicellulose

structure as reported by (Kumar et al., 2014). The bands associated with cellulose

structures are 1429, 1370, 1319, 1103 and 898 cm-1

(Karatzos et al., 2012; Kumar et

al., 2014). The band position at 1421 cm-1

is mainly due to CH2 scissor motion in

cellulose. In current study, the increase in band intensity at 1421 cm-1

and other

cellulose associated band can be attributed to increase in cellulose content due to

removal of lignin and hemicellulose content. During compositional analysis of

bagasse it was observed that removal of hemicellulose content resulted in increased

cellulose and lignin content. Similar results were studied by Sun et al., (2015) that the

cellulose band intensity was increased after pretreatments due to decrease in

hemicellulose content. Moreover, the present study showed that the spectra obtained

from autohydrolyzed samples showed clear peaks at 1103 cm-1

and a reduction of the

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amorphous band at 898 cm-1

, which demonstrated an increase in crystallinity. The

peak at 1103 cm-1

(referring to crystalline cellulose) was diminished after IL

pretreatment which is a clear indication of cellulose crystallinity. Similar results were

found in previous studies where various pretreatment strategies reduced crystallinity

after IL pretreatment as reported by Li et al., (2010); however, Sun et al., (2015)

reported an increase in crystallinity after acidic pretreatment. These results clearly

demonstrated that how both pretreatments effectively weaken the van der Waals

interaction between cell wall polymers (Li et al., 2010).

5.8.Crystallinity Measurement

The quantitative analysis of absorption spectrometry is based on the Beer-Bouguer-

Lambert law which implies that absorption band intensities are linearly proportional

to the concentration of each component (Adapa et al., 2011). During this research the

peak heights were used as quantitative parameter as stated by Adapa et al. (2011), and

the heights of the peaks were determined by measuring the difference between

maximum peak intensity and baseline (Adapa et al., 2011). Total crystallinity index

(TCI) and Lateral order index (LOI) was determined for all pretreated samples. The

term Lateral order index (LOI) was assigned to two ratios related to cellulose

structure were calculated i.e. 1424 cm-1

/898 cm-1

(Hurtubise and Krässig, 1960;

Spiridon et al., 2011; Qiu et al., 2012). It has been used by many previous researchers

to show the presence of cellulose I structure in cellulose material (Oh et al., 2005;

Spiridon et al., 2011). Total crystallinity index (TCI) was the term used by Nelson

and O’Connor (1964) for the ratio of 1371 cm-1

/2919 cm-1

, and used by various

researchers to evaluate the infrared crystallinity (IR) ratio (Nelson and O'Connor,

1964; Spiridon et al., 2011; Qiu et al., 2012). Thus, the higher values of both LOI and

TCI are indicative of the more ordered structure of cellulose and higher crystallinity

of biomass (Qiu et al., 2012). The order of crystallinity determined by LOI was 205ºC

> 190ºC > untreated bagasse > 110ºC > IL pretreated bagasse which also corroborated

with the CrI (crystallinity index) determined by XRD analysis. The TCI values of

different pretreated samples showed deviation from this order which can be attributed

to hydrocarbonate linear chain extractives which can be associated with 2900 cm-1

band intensity (Ornaghi et al., 2014), thus showed higher values in this specific band,

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decreasing the calculated total crystallinity value (Ornaghi et al., 2014; Poletto et al.,

2014). However, the evaluation of LOI was based on 898 cm-1

band, which is

associated with amorphous cellulose, so a higher intensity in this band indicates more

amorphous content (Ornaghi et al., 2014). These reduction in values showed that

during IL pretreatment process, crystalline structure of cellulose was converted into

amorphous cellulose (Li et al., 2010). Similar results were found by Spiridon et al.,

(2010) that the IL pretreatment reduced the crystallinity of cellulose as determined by

TCI and LOI thus it provided more surface area for enzyme attachment and

conversion. Qiu et al., (2012) pretreated the bagasse with [C2mim][OAC] which was

shown to have positive effect on reduction of crystallinity determined by measuring

TCI and LOI. Zhu et al., (2012) showed that LOI and TCI of sugarcane bagasse was

decreased when treated with NH4OH–H2O2+[Amim]Cl-pretreated (100°C for 1 h),

similarly, Kuo et al. (2009) also observed the decrease in TCI and LOI values when

sugarcane bagasse was treated with N-methylmorpholine-N-oxide (NMMO) at 100°C

for 7 h.

XRD analysis was performed to examine the cellulose crystallinity index of biomass

(Park et al., 2010). The slight increase in CrI in autohydrolyzed samples can be

attributed to removal of amorphous region i.e. hemicellulose, lignin or amorphous

cellulose and rearrangement of remaining components (Ruiz et al., 2011; Zhang et al.,

2012). Lei et al., (2013) reported that the dilute acid pretreatment was unable to break

cellulose hydrogen bonding but removed the amorphous components; thus increased

the crystallinity of biomass. The reduction in CrI of IL treated sample was observed,

which exhibited reduction in cellulose crystallinity in bagasse sample treated with

[C4mim][OAc]. Previous studies has been showed the crystallinity index of untreated

bagasse as 0.56 which was reduced to 0.24 after [C2mim][OAc] pretreatment (Qiu et

al., 2012). Li et al., (2010) and Saliva et al., (2011) also determined the reduction in

CrI after pretreatment of biomass with [C2mim][OAc]. It was suggested that anion

and cation in IL were responsible for disruption of cellulose structure. The cation

interacted with lignin though Π-Π interaction and hydrogen bonding whereas anionic

acetate acted as hydrogen bond acceptor that attacked the free hydroxyl group of

cellulose and deprotonated it, thus reduced cellulose crystallinity (Qiu and Aita, 2013;

Ninomiya et al., 2015). Hydrogen bonding in cellulose structure was disrupted and

replaced by another hydrogen bonding between cellulose hydroxyl and anions of ionic

PhD Thesis


liquid, thus it caused disruption and dissolution of cellulose structure and reduced its

crystallinity (Qiu and Aita, 2013).

5.9.Enzymatic Hydrolysis

The cellulose digestibility is considered as an important factor to select the most

efficient method of pretreatment. Some important factors which act as a barrier during

enzymatic hydrolysis of a biomass are lignin hindrance, hemicellulose content,

porosity, cellulose degree of polymerization and cellulose crystallinity. Variation in

glucose and xylose concentration released by enzymatic hydrolysis of sugarcane

bagasse pretreated with different strategies was observed and the major effect found

by autohydrolysis on lignocellulosic material was removal of hemicellulose which

increased the accessibility of enzymes to cellulosic content, thus higher concentration

of sugars were obtained with increase in pretreatment temperature (C. Li et al., 2010;

Batalha et al., 2015). Severity in pretreatment conditions showed profound effects on

enzymatic hydrolysis. It was observed that increased pretreatment temperature of

autohydrolysis enhanced glucose release after enzymatic hydrolysis but xylose

concentration was reduced because major amount of xylan has already been removed

during autohydrolysis. Among the autohydrolyzed samples, maximum glucose

concentration was obtained from the samples autohydrolyzed at 205ºC for 6 min;

whereas maximum xylose concentration under these conditions was obtained from

sample autohydrolyzed at 190°C for 10 min. The reduction in xylose concentration

released from bagasse autohydrolyzed at 205ºC could be attributed to removal of

large hemicellulose content during high temperature autohydrolysis but low xylose

concentration autohydrolyzed sample at 110ºC might be due to more lignin hindrance

(Qiu et al., 2012; Hongdan et al., 2013; Batalha et al., 2015). Enzymatic

saccharification of the ionic liquid pretreated bagasse (at 110ºC for 30 min) released

more glucose and xylose as compared to all autohydrolyzed samples which depicted

increased conversion of cellulose into glucose as compared to other pretreatments.

Ionic liquid exhibited good effect on removal of hemicellulose but compositional

analysis and FTIR data of pretreated bagasse revealed that the effect of ionic liquid on

hemicellulose was much lesser than autohydrolysis, thus released more xylose from

biomass after enzymatic hydrolysis.

PhD Thesis


Autohydrolysis was compared with ionic liquid pretreatment to determine kinetics of

enzymatic hydrolysis and cellulose digestibility, which showed increased kinetics i.e.

97.4% in IL-treated bagasse. Biomass pretreated with [C2mim][OAc] has been shown

cellulose digestibility up to 87% and 96% within 24 h of hydrolysis in various

previous studies (Li et al., 2010; Qiu et al., 2012). In case of autohydrolysis, the

limited enzymatic hydrolysis and cellulose digestibility can be attributed to

unmodified crystalline cellulosic structure. The loss in inter- and intra- molecular

hydrogen bonding during IL pretreatment resulted in amorphous cellulose and

provided an enhanced surface area leading to better enzyme accessibility and

increased binding sites in recovered cellulose fibers (Li et al., 2010; Qiu et al., 2012).

Higher hemicellulose digestibility from ionic liquid pretreated bagasse might be

attributed to minimal loss of initial xylan and delignification. Silva et al., (2011)

reported 75% xylan digestion when treated with [C2mim][OAc]. In autohydrolyzed

bagasse hemicellulose were not accessible to enzymes because most of the

hemicellulose was already been removed and rest of it was covalently linked with

lignin component (Karatzos et al., 2012).

Hydrolysis results clearly showed that soluble sugars released faster to a greater

extend in IL pretreated sugarcane bagasse than autohydrolysis pretreatment. However,

reduction in enzymatic loading, low cost ionic liquid and recovery of ionic liquid are

essential to promote the economy of bio-refineries and develop the optimal ionic

pretreatment (Li et al., 2010). It was also observed that total processing time to reach

60% cellulose digestibility was about 48 h with autohydrolysis but it was reached up

to 80% within 3 h with ionic liquid pretreatment. In comparison to autohydrolysis,

ionic liquid required low energy consumption, less processing time, lead to higher

glucose yield and it is convenient and environment friendly too. These advantages are

paramount in order to counterbalance higher costs associated with ionic liquids but it

also offers motivation to explore and develop this pretreatment technique in the

future.

5.10. Fermentation

Ethanol production from fermentation of pretreated samples of sugarcane bagasse

showed that different microbial strains have different abilities to produce ethanol from

PhD Thesis


available sugar. The maximum amount of ethanol was obtained from IL pretreated

sample when it was fermented with newly isolated MZ-4 strain. The difference in

ethanol production amount by different strains can be attributed to their difference in

tolerance against side products released during different pretreatment conditions

(Iwaki et al., 2013). Previously, it was reported that ILs have negative affect on

growth of microorganisms; and the toxic role of ILs on microbes can be attributed to

either cationic or anionic moiety (Frade and Afonso, 2010; Pham et al., 2010).

Presence of [C2mim][OAc] (up to 1% concentration ) in fermentation medium

improves the growth of S. cerevisiae and also increases ethanol production but its

higher concentrations has negative impact on yeast growth and fermentation process

(Ouellet et al., 2011; Mehmood et al., 2015). S. cerevisiae has the ability to utilize

both monomeric sugar and sucrose which make it an efficient microbe to be used in

variety of substrates (Badotti et al., 2008; Canilha et al., 2012). Other advantages

related to its use are its highest resistance against high ethanol concentration, inhibitor

resistance and its ability to consume significant amount of substrate in adverse

conditions. Unfortunately, S. cerevisiae lacks genes which could make it able to

assimilate xylose; however, to obtain optimal ethanol yields, conversion of

hemicellulose fraction is also essential (Canilha et al., 2012). Pitchia stipites is

considered as more efficient to ferment xylose into ethanol but its less production

with all pretreated samples can be attributed to its less efficient glucose utilization as

compared to S. cerevisiae (Krahulec et al., 2012). Moreover, it was also suggested in

previous studies that all symporters in P. stipites are competitively inhibited by

glucose molecules which makes it difficult to utilize both sugars simultaneously and

hinders the conversion of xylose into ethanol (Farwick et al., 2014).

PhD Thesis


Conclusions

In this study, different strategies were applied to enhance bioethanol yield from sugar

industry waste under optimal physicochemical conditions. The following findings were

concluded from the study:

For the production of bioethanol from sugarcane molasses by using batch

fermentation, a newly isolated strain MZ-4 was determined as the most efficient

strain in terms of bioethanol yield and fermentation efficiency.

In comparison, fed-batch fermentation process showed better yield and

fermentation efficiency as compared to batch fermentation.

During fed batch fermentation of sugarcane molasses, Lalvin EC-1118 showed

better bioethanol production than MZ-4 in terms of actual yield; however, MZ-4

strain showed less yield with improved fermentation efficiency. Therefore,

despite of lower ethanol yield, MZ-4 can be considered as a better strain in terms

of maximum sugar conversion in to bioethanol. Lalvin EC-1118 can be preferred

for maximum yield and to avoid time constraints, because more time is required

by strain MZ-4 to complete the fermentation process.

For the conversion of sugarcane bagasse into bioethanol, IL pretreatment showed

better effect on release of sugars as compared to high severity autohydrolysis

pretreatment, which can contribute in reduction of operational cost. Similarly, the

higher cellulose and xylan digestibility was also observed after IL pretreatment,

which might be attributed to removal of lignin and hemicelluloses along with

conversion of cellulose crystalline structure into amorphous, during pretreatment.

The reaction rate for the enzymatic hydrolysis was much faster for the IL

pretreated bagasse as compared to autohydrolyzed bagasse, which makes IL

pretreatment more favorable.

In comparison to autohydrolysis, the greater ethanol production was obtained

from IL pretreated bagasse, when it was fermented with strain MZ-4.

PhD Thesis


Future Prospects

A lot of work has already been done regarding production of bioethanol from yeast by

using cheaper substrates. However, still there are many problems to face while scaling up

the ethanol production in industrial scale.

Work on continuous fermentation should be done to overcome the inhibitory effect of

end product.

Genetically modified crops containing high sugar content can be used to enhance the

bioethanol production.

An effort to create genetically modified microorganisms (GMOs) to get higher

ethanol yield should be done, but they should have to certify as ―GRAS‖ (generally

recognized as safe) prior to use.

The process of simultaneous saccharification and co-fermentation should be tested on

IL pretreated bagasse by using those GMOs, which should be able to convert glucose

and xylose simultaneously.

Optimization of IL pretreatment and following fermentation process should be done

to enhance the production of ethanol.

Other pretreatment strategies on sugarcane bagasse should be tested to increase the

amount of released sugar.

Inhibitors released during this pretreatment should be determined and efforts should

be made to avoid their effects on biological processes.

This study should be scaled up and optimized to the pilot scale, so that best

determined strategies could be utilized on industrial scale at later stages.

PhD Thesis


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PhD Thesis


Annexes

PhD Thesis

161

Appendix A

A1: The Composition of WLN Media

Contents Amount (g/L)

Yeast extract 4.00

Trypton 5.00

Glucose 50.00

Potassium dihydrogen phosphate 0.55

Potassium chloride 0.425

Magnesium sulphate 0.125

Calcium chloride 0.125

Ferric chloride 0.0025

Manganese sulphate 0.0025

Bromocresol green 0.022

Agar 15.00

PhD Thesis

162

Appendix B

B1: Sequence for Strain MZ-4

ccccctccgtcctgggccccagtccctccgggggcgcccttattcagtgccatccctggaggggctgaaaagcgttcccaattt

gtaatgggcggacaaaatccatactcgtgtggggggcccccattaataggtttcctggtttttgagcgtgagacgcccctattggg

agcggccccaagtgccgggtcgtccgtttgaagaaaaaaggccggaggattggggcccgctgctttttgtctagtaaatgttgca

aacaaactcagcagaagtaa

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163

Appendices C

C.1: Calibration curve for reducing sugar analysis by DNS method

C.2: Calibration curve for ethanol determination by HPLC method

y = 0.2682x R² = 0.9977

0

0.5

1

1.5

2

2.5

0 2 4 6 8 10

O.D

(n

m)

Reducing Sugars (mg/ml)

y = 237812x R² = 0.998

0

2000000

4000000

6000000

8000000

10000000

12000000

14000000

0 10 20 30 40 50 60

HP

LC p

eak

are

a

Ethanol % (v/v)

PhD Thesis

164

C.3: Calibration curve for glucose determination by HPLC method

C.4: Calibration curve for xylose determination by HPLC method

y = 6.9543x R² = 0.9972

0

5

10

15

20

25

30

0 0.5 1 1.5 2 2.5 3 3.5 4

HP

LC p

eak

he

igh

t

Glucose Conc. (mg/ml)

y = 5.0696x R² = 0.998

0

0.5

1

1.5

2

2.5

3

3.5

4

4.5

0 0.2 0.4 0.6 0.8 1

HP

LC p

eak

he

igh

t

Xylose Conc. (mg/ml)

PhD Thesis

165

C.5: Calibration curve for galactose determination by HPLC method

C.6: Calibration curve for arabinose determination by HPLC method

y = 3.1419x R² = 0.9974

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0.9

0 0.05 0.1 0.15 0.2 0.25 0.3

HP

LC p

eak

He

igh

t

Galactose Conc. (mg/ml)

y = 57.175x R² = 0.9988

0

0.2

0.4

0.6

0.8

1

1.2

1.4

0 0.005 0.01 0.015 0.02 0.025

HP

LC p

eak

he

igh

t

Arabinose Conc. (mg/ml)

PhD Thesis

166

C.7: Calibration curve for mannose determination by HPLC method

y = 48.089x R² = 0.9993

0

0.5

1

1.5

2

2.5

0 0.01 0.02 0.03 0.04 0.05

HP

LC P

eak

he

igh

t

Mannose Conc. (mg/ml)

PhD Thesis

167

Appendices D

Bioethanol production from Sugarcane Molasses

D.1: Effect of pH on enhanced production of bioethanol by using Lalvin EC-

1118 and MZ-4

pH

Lalvin EC-1118 MZ-4

Ethanol

Productio

n (v/v)

S.D. Ethanol

Production

(v/v)

S.D.

3.0 6.4 ±0.124 7.2 ±0.124

3.5 7.3 ±0.081 8.2 ±0.081

4.0 9.6 ±0.163 9.6 ±0.163

4.5 9.8 ±0.081 10.1 ±0.081

5.0 8.9 ±0.163 10.2 ±0.081

5.5 8.3 ±0.081 9.5 ±0.163

6.0 8.1 ±0.081 8.2 ±0.124

S.D= Standard deviation

D.2: Effect of temperature on enhanced production of bioethanol by using


Temp

Lalvin EC-1118 MZ-4

Ethanol

Productio

n (v/v)

S.D. Ethanol

Production

(v/v)

S.D.

27 9.5 ±0.163 9.8 ±0.081

30 9.8 ±0.081 10.2 ±0.163

33 9.4 ±0.169 10.3 ±0.244

36 8.9 ±0.205 9.8 ±0.081

39 7.1 ±0.169 7.5 ±0.249


PhD Thesis

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D.3: Effect of Inoculum size on enhanced production of bioethanol by using


Inoculum

Size

Lalvin EC-1118 MZ-4

Ethanol

Productio

n (v/v)

S.D. Ethanol

Production

(v/v)

S.D.

2.5 9.5 ±0.163 10.0 ±0.163

5 9.8 ±0.081 10.3 ±0.081

7.5 10 ±0.081 10.4 ±0.163

10 9.7 ±0.169 10.5 ±0.163

12.5 9.6 ±0.124 10.2 ±0.081


D.4: Effect of Inoculum age on enhanced production of bioethanol by using


Inoculum

Age

(hours)

Lalvin EC-1118 MZ-4

Ethanol

Productio

n (v/v)

S.D. Ethanol

Production

(v/v)

S.D.

12 9.8 ±0.163 10.1 ±0.124

24 10 ±0.081 10.5 ±0.163

36 9.4 ±0.081 9.4 ±0.124

48 8.4 ±0.169 8.3 ±0.163


PhD Thesis

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D.5: Effect of nitrogen sources on enhanced production of bioethanol by

using Lalvin EC-1118 and MZ-4

Nitrogen

source

Concentrati

on

Lalvin EC-1118 MZ-4

Ethanol

Productio

n (v/v)

S.D Ethanol

Product

ion (v/v)

S.D

Urea 0.05% 10.1 ±0.081 10.5 ±0.163

0.10% 10.2 ±0.163 10.7 ±0.081

0.15% 9.4 ±0.124 9.6 ±0.169

Ammonium

Chloride

0.05% 10.1 ±0.081 10.6 ±0.081

0.10% 10.3 ±0.244 10.8 ±0.081

0.15% 10.4 ±0.081 9.9 ±0.286

Ammonium

Nitrate

0.05% 10.3 ±0.081 10.5 ±0.163

0.10% 9.4 ±0.047 10.6 ±0.081

0.15% 8.9 ±0.205 10.1 ±0.081

Ammonium

Sulfate

0.05% 10.1 ±0.081 10.5 ±0.163

0.10% 10.3 ±0.163 10.6 ±0.081

0.15% 10.4 ±0.081 10.7 ±0.163

Di-ammonium

phosphate

0.05% 10.3 ±0.081 10.5 ±0.081

0.10% 10.5 ±0.081 10.6 ±0.081

0.15% 10 ±0.081 9.9 ±0.249


PhD Thesis

170

D.6: Effect of chelating agent sources on enhanced production of bioethanol


Chelating

agents

Conc. Lalvin EC-1118 MZ-4

Ethanol

Production

(v/v)

S.D. Ethanol

Production

(v/v)

S.D.

EDTA 0.0025 10.5 ±0.163 10.7 ±0.163

0.005 10.5 ±0.081 10.7 ±0.081

0.01 10.6 ±0.163 10.8 ±0.163

0.02 10.6 ±0.081 10.8 ±0.081

0.04 10.7 ±0.163 10.9 ±0.163

0.08 10.4 ±0.081 10.2 ±0.047

0.16 9.8 ±0.169 9.36 ±0.047

0.32 9.2 ±0.188 8.8 ±0.081

Potassium

Ferrocyanide

0.0025 10.5 ±0.163 10.7 ±0.163

0.005 10.6 ±0.163 10.9 ±0.081

0.01 10.7 ±0.081 11.1 ±0.163

0.02 10.8 ±0.081 10.3 ±0.047

0.04 10.9 ±0.163 9.7 ±0.047

0.08 10.3 ±0.094 9.1 ±0.047

0.16 9.6 ±0.124 8.5 ±0.124

0.32 9.4 ±0.081 8.0 ±0.081

Sodium

Potassium

tartrate

0.0025 10.5 ±0.163 10.7 ±0.163

0.005 10.5 ±0.081 10.7 ±0.081

0.01 10.6 ±0.081 10.8 ±0.081

0.02 10.6 ±0.163 10.8 ±0.047

0.04 10.7 ±0.081 10.1 ±0.047

0.08 10.6 ±0.124 9.2 ±0.081

0.16 10.1 ±0.162 8.4 ±0.094

0.32 10.5 ±0.163 7.7 ±0.124


PhD Thesis

171

Appendices E

Ethanol Production from Sugarcane Bagasse

E.1.1. Glucose concentration released from untreated and autohydrolyzed samples


Time

(hours)

Untreated bagasse 190°C for 10 min 205°C for 6 min

Glucose

(mg/ml)

S.D. Glucose

(mg/ml)

S.D. Glucose

(mg/ml)

S.D.

0 0 ±0 0 ±0 0 0

3 0.33 ±0.027 1.05 ±0.049 1.55 ±0.015

6 0.45 ±0.045 1.35 ±0.019 2.02 ±0.075

12 0.46 ±0.039 1.55 ±0.083 2.49 ±0.088

24 0.46 ±0.014 1.88 ±0.011 2.98 ±0.067

48 0.47 ±0.014 2.20 ±0.044 3.39 ±0.067

72 0.47 ±0.014 2.37 ±0.061 3.53 ±0.148


E.1.2: Cellulose digestibility of untreated and autohydrolyzed samples during


Untreated control 190190°C for 10 min 205°C for 6 min

Time

(h)

Cellulose

digestibilit

y (%)

S.D. Cellulose

digestibili

ty (%)

S.

D

Cellulose

digestibili

ty (%)

S.D.

0 0 ±0 0 ±0 0 ±0

3 8.19 ±0.675 20.85 ±0.985 27.16 ±0.278

6 10.37 ±1.109 25.43 ±0.385 34.69 ±1.328

12 11.02 ±0.962 30.62 ±1.640 42.80 ±1.550

24 11.21 ±0.689 37.08 ±0.224 51.54 ±1.179

48 11.43 ±0.344 43.55 ±0.882 58.78 ±1.190

72 11.46 ±0.344 46.93 ±1.205 60.29 ±2.602


PhD Thesis

172

E.1.3: Xylose concentration released from untreated and autohydrolyzed samples



Time

(h)

Xylose

(mg/ml)

S.D. Xylose

(mg/ml)

S.D. Xylose

(mg/ml)

S.D.

0 0 ±0 0 ±0 0 ±0

3 0.003 ±0.002 0.25 ±0.003 0.13 ±0.011

6 0.01 ±0.001 0.29 ±0.002 0.15 ±0.004

12 0.029 ±0.000 0.32 ±0.011 0.18 ±0.004

24 0.03 ±0.001 0.38 ±0.014 0.21 ±0.0057

48 0.03 ±0.001 0.41 ±0.026 0.24 ±0.0057

72 0.03 ±0.002 0.426 ±0.035 0.24 ±0.002


E.1.4: Xylan digestibility of untreated and autohydrolyzed samples during



Time

(h)

Xylan

digestib

ility (%)

S.D. Xylan

digestib

ility (%)

S.D. Xylan

digestibili

ty (%)

S.D.

0 0 ±0 0 ±0 0 ±0

3 0.11 ±0.101 2.765 ±0.042 2.86 ±0.257

6 0.76 ±0.065 3.135 ±0.024 3.54 ±0.097

12 1.41 ±0.018 3.527 ±0.122 4.24 ±0.097

24 1.44 ±0.045 4.197 ±0.157 4.88 ±0.132

48 1.44 ±0.048 4.502 ±0.284 5.57 ±0.133

72 1.48 ±0.115 4.677 ±0.379 5.690 ±0.049


PhD Thesis

173

E.2.1: Glucose concentration released from untreated control, water treated control

and ionic liquid pretreated samples during enzymatic hydrolysis

Untreated control A-110°C for 30 min IL 110°C for 30 min

Time Glucose

(mg/ml)

S.D. Glucose

(mg/ml)

S.D. Glucose

(mg/ml)

S.D.

0 0 ±0 0 ±0 0 ±0

3 0.33 ±0.027 0.60 ±0.192 3.31 ±0.033

6 0.45 ±0.045 0.66 ±0.018 3.64 ±0.033

12 0.46 ±0.039 0.70 ±0.001 3.67 ±0.052

24 0.46 ±0.014 0.74 ±0.016 3.77 ±0.047

48 0.47 ±0.014 0.78 ±0.016 3.92 ±0.015

72 0.47 ±0.014 0.82 ±0.004 4.04 ±0.011


E.2.2: Cellulose digestibility from untreated control, water treated control and

ionic liquid pretreated samples during enzymatic hydrolysis


Time Cellulose

digestibility

(%)

S.D. Cellulose

digestibility

(%)

S.D. Cellulose

digestibility

(%)

S.D.

0 0 ±0 0 ±0 0 ±0

3 8.19 ±0.675 14.25 ±0.404 79.88 ±0.800

6 10.37 ±1.109 15.61 ±0.061 87.81 ±1.273

12 11.02 ±0.962 16.54 ±0.360 88.48 ±1.133

24 11.21 ±0.689 17.46 ±0.369 90.92 ±0.362

48 11.46 ±0.344 18.48 ±0.066 94.59 ±0.276

72 11.46 ±0.344 19.31 ±0.561 97.44 ±1.476


PhD Thesis

174

E.2.3: Xylose digestibility from untreated control, water treated control and ionic

liquid pretreated samples during enzymatic hydrolysis


Time Xylose

(mg/ml)

S.D. Xylose

(mg/ml)

S.D. Xylose

(mg/ml)

S.D.

0 0 ±0 0 ±0 0 ±0

3 0.003 ±0.002 0.092 ±0.012 0.924 ±0.002

6 0.016 ±0.001 0.111 ±0.012 1.073 ±0.005

12 0.029 ±0.000 0.124 ±0.004 1.164 ±0.002

24 0.030 ±0.001 0.135 ±0.007 1.270 ±0.004

48 0.030 ±0.001 0.170 ±0.001 1.365 ±0.009

72 0.031 ±0.002 0.178 ±0.014 1.401 ±0.006


E.2.4: Xylan digestibility from untreated control, water treated control and ionic

liquid pretreated samples during enzymatic hydrolysis


Time Xylan

digestibility

(%)

S.

D

Xylan

digestibility

(%)

S.D. Xylan

digestibility

(%)

S.D.

0 0 ±0 0 ±0 0 ±0

3 0.119 ±0.101 4.280 ±0.599 65.12 ±0.198

6 0.762 ±0.065 5.170 ±0.574 75.63 ±0.399

12 1.418 ±0.014 5.800 ±0.205 81.99 ±0.187

24 1.441 ±0.048 6.320 ±0.341 89.48 ±0.318

48 1.441 ±0.048 7.911 ±0.079 96.16 ±0.684

72 1.489 ±0.115 8.321 ±0.651 98.67 ±0.476


PhD Thesis

175

E.3.1: Bioethanol Production from Untreated control by various yeast strains

Strains Ethanol (mg/g-substrate) S.D.

Uvaferm-43 6.02 ±0.11

Lalvin EC-1118 12.21 ±0.25

MZ-4 15.52 ±0.05

Pitchia stipitis 10.67 ±0.11


E.3.2.Bioethanol Production from bagasse autohydrolyzed at 190°C by various

yeast strains


E.3.3: Bioethanol Production from bagasse autohydrolyzed at 205°C by various

yeast strains



Uvaferm-43 56.33 ±0.74

Lalvin EC-1118 66.02 ±0.65

MZ-4 56.86 ±0.15

Pitchia stipilis 50.09 ±1.04


Uvaferm-43 68.55 ±0.75

Lalvin EC-1118 69.42 ±0.36

MZ-4 70.92 ±0.09


PhD Thesis

176

E.3.4: Bioethanol Production from bagasse autohydrolyzed at 110°C by various

yeast strains


Uvaferm-43 13.84 ±0.142

Lalvin EC-1118 28.42 ±0.122

MZ-4 16.20 ±0.983



E.3.5: Bioethanol Production from IL pretreated bagasse at 110°C by various yeast

strains


Uvaferm-43 77.00 ±0.269

Lalvin EC-1118 59.96 ±0.707

MZ-4 78.78 ±0.943



PhD Thesis

177

Appendix F

STATISTICAL ANALYSIS FOR BIOETHANOL PRODUCTION FROM

SUGARCANE MOLASSES

F.1: Analysis of variance for the effect of pH on enhanced production of bioethanol


Yeast

Strains

Sum of

Squares

df Mean

Square

F Sig.

Lalvin

EC-1118

Between

Groups

26.426 6 4.404 215.093 .000

Within

Groups

.287 14 .020

Total 26.712 20

MZ-4 Between

Groups

22.319 6 3.720 166.206 .000

Within

Groups

.313 14 .022

Total 22.632 20

F.2: Analysis of variance for the effect of temperature on enhanced production of


Yeast

strains

Sum of

Squares

df Mean

Square

F Sig.

LalvinE

C1118

Between

Groups

13.409 4 3.352 83.808 .00

Within Groups .400 10 .040

Total 13.809 14

MZ4 Between

Groups

15.127 4 3.782 77.705 .00

Within Groups .487 10 .049

Total 15.613 14

df= degree of freedom

PhD Thesis

178

F.3: Analysis of variance for the effect of inoculum size on enhanced production of


Yeast

Strains

Sum of

Squares

df Mean

Square

F Sig.

LalvinEC

-1118

Between

Groups

.420 4 .105 4.145 .031

Within

Groups

.253 10 .025

Total .673 14

MZ4 Between

Groups

.444 4 .111 3.964 .035

Within

Groups

.280 10 .028

Total .724 14

df= degree of freedom; Sig.= P value

F.4: Analysis of variance for the effect of inoculum age on enhanced production of

bioethanol by using Lalvin MZ-4

Yeast

Strains

Sum of

Squares df

Mean

Square F Sig.

LalvinE

C1118

Between

Groups 4.363 3 1.454 56.290 .000

Within

Groups .207 8 .026

Total 4.569 11

MZ4 Between

Groups 8.516 3 2.839 89.640 .000

Within

Groups .253 8 .032

Total 8.769 11


PhD Thesis

179


bioethanol by using Lalvin EC-1118


Nitrogen

Source

Sum of

Squares

df Mean

Square

F Sig.

Urea Between

Groups

1.802 2 .901 28.964 .001

Within

Groups

.187 6 .031

Total 1.989 8

Ammonium

Chloride

Between

Groups

1.236 2 .618 12.930 .007

Within

Groups

.287 6 .048

Total 1.522 8

Ammonium

Nitrate

Between

Groups

.420 2 .210 10.500 .011

Within

Groups

.120 6 .020

Total .540 8

Ammonium

Sulfate

Between

Groups

.060 2 .030 1.000 .422

Within

Groups

.180 6 .030

Total .240 8

Diammonium

phosphate

Between

Groups

.696 2 .348 9.206 .015

Within

Groups

.227 6 .038

Total .922 8

PhD Thesis

180


bioethanol by using MZ-4



Nitrogen

Source

Sum of

Squares

df Mean

Square

F Sig.

Urea Between

Groups .949 2 .474 19.409 .002

Within

Groups .147 6 .024

Total 1.096 8

Ammonium

Chloride

Between

Groups .140 2 .070 1.909 .228

Within

Groups .220 6 .037

Total .360 8

Ammonium

Nitrate

Between

Groups 2.722 2 1.361 53.261 .000

Within

Groups .153 6 .026

Total 2.876 8

Ammonium

Sulfate

Between

Groups .140 2 .070 3.500 .098

Within

Groups .120 6 .020

Total .260 8

Diammonium

Phosphate

Between

Groups .380 2 .190 19.000 .003

Within

Groups .060 6 .010

Total .440 8

PhD Thesis

181

F.7: Analysis of variance for the effect of chelating agents on enhanced production

of bioethanol by using Lalvin EC-1118

Chelating

agents

Sum of

Squares

df Mean

Square

F Sig

.

EDTA Between

Groups

5.056 7 .722 23.427 .000

Within

Groups

.493 16 .031

Total 5.550 23

Potassium

Ferrocyanide

Between

Groups

6.240 7 .891 38.204 .000

Within

Groups

.373 16 .023

Total 6.613 23

Sodium

potassium

tartarate

Between

Groups

2.660 7 .380 17.208 .000

Within

Groups

.353 16 .022

Total 3.013 23


PhD Thesis

182

F.8: Analysis of variance for the effect of chelating agents on enhanced production

of bioethanol by using MZ-4

Chelating

agents

Sum of

Squares

df Mean

Square

F Sig

.

EDTA Between

Groups

12.905 7 1.844 94.140 .000

Within

Groups

.313 16 .020

Total 13.218 23

Potassium

Ferrocyanide

Between

Groups

27.827 7 3.975 238.514 .000

Within

Groups

.267 16 .017

Total 28.093 23

Sodium

potassium

tartarate

Between

Groups

30.467 7 4.352 307.227 .000

Within

Groups

Total .227 16 .014


PhD Thesis

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F.9: Analysis of variance for the effect of fed batch fermentation on enhanced

production of bioethanol by using Lalvin EC-1118

Sum of

Squares

df Mean

Square

F Sig.

Between

Groups

28.772 6 4.795 3.622 .013

Within

Groups

27.805 21 1.324

Total 56.577 27


F10: Analysis of variance for the effect of fed batch fermentation on enhanced

production of bioethanol by using MZ-4

Sum of

Squares

df Mean

Square

F Sig.

Between

Groups

21.528 6 3.588 3.065 .026

Within

Groups

24.585 21 1.171

Total 46.113 27


PhD Thesis

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Appendix G

Statistical Analysis for Bioethanol Production from Bagasse

G.1.1: Analysis of variance for the glucose concentration released from untreated

and autohydrolyzed samples during enzymatic hydrolysis

Sum of

Squares

df Mean

Square

F Sig.

Between

Groups

20.170 4 5.043 833.1

70

.000

Within

Groups

.030 5 .006

Total 20.201 9


PhD Thesis

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G.1.2: Post hoc Tukey HSD multiple comparisons for the glucose concentration released from untreated and pretreated

bagasse samples during enzymatic hydrolysis

(I)

Pretreatme

nt

(J)

Pretreatment

Mean Difference (I-

J)

Sig. 95% Confidence Interval

Lower

Bound

Upper Bound

Untreated

Control

Autohydrolysis-190C -1.90938* .000 -2.2215 -1.5973

Autohydrolysis-205 C -3.06535* .000 -3.3774 -2.7533

Water treated- 110C -.35547* .030 -.6675 -.0434

IL- 110 C -3.57406* .000 -3.8861 -3.2620

Autohydro

lysis-190C

Untreated Control 1.90938* .000 1.5973 2.2215

Autohydrolysis-205 C -1.15597* .000 -1.4680 -.8439

Water treated- 110C 1.55392* .000 1.2418 1.8660

IL- 110 C -1.66467* .000 -1.9768 -1.3526

Autohydro

lysis-205 C

Untreated Control 3.06535* .000 2.7533 3.3774

Autohydrolysis-190C 1.15597* .000 .8439 1.4680

Water treated- 110C 2.70988* .000 2.3978 3.0220

IL- 110 C -.50871* .007 -.8208 -.1966

Water

treated-

110C

Untreated Control .35547* .030 .0434 .6675

Autohydrolysis-190C -1.55392* .000 -1.8660 -1.2418

Autohydrolysis-205 C -2.70988* .000 -3.0220 -2.3978

IL- 110 C -3.21859* .000 -3.5307 -2.9065

IL- 110 C Untreated Control 3.57406* .000 3.2620 3.8861

Autohydrolysis-190C 1.66467* .000 1.3526 1.9768

Autohydrolysis-205 C .50871* .007 .1966 .8208

Water treated- 110C 3.21859* .000 2.9065 3.5307

*. The mean difference is significant at the 0.05 level.

PhD Thesis

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G.2.1: Analysis of variance for the xylose concentration released from untreated and

pretreated bagasse samples during enzymatic hydrolysis

Sum of

Squares

df Mean

Square

F Sig.

Between

Groups

2.388 4 .597 1990.823 .000

Within

Groups

.001 5 .000

Total 2.389 9


PhD Thesis

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G2.2: Post hoc-Tukey HSD multiple comparisons for the xylose concentration released from untreated and pretreated

bagasse samples during enzymatic hydrolysis

(I)

Pretreatment

(J) Pretreatment Mean

Difference

(I-J)

Std. Error 95% Confidence Interval

Lower

Bound

Upper

Bound

Untreated

Control

Autohydrolysis-190C -.40369* .01732 -.4732 -.3342

Autohydrolysis-205 C -.21721* .01732 -.2867 -.1477

Water treated- 110C -.14763* .01732 -.2171 -.0782

IL- 110 C -1.36986* .01732 -1.4393 -1.3004

Autohydrolysi

s-190C


Autohydrolysis-205 C .18648* .01732 .1170 .2559

Water treated- 110C .25606* .01732 .1866 .3255

IL- 110 C -.96618* .01732 -1.0356 -.8967

Autohydrolysi

s-205 C



Water treated- 110C .06958* .01732 .0001 .1390

IL- 110 C -1.15265* .01732 -1.2221 -1.0832

Water

treated- 110C



Autohydrolysis-205 C -.06958* .01732 -.1390 -.0001

IL- 110 C -1.22224* .01732 -1.2917 -1.1528

IL- 110 C Untreated Control 1.36986* .01732 1.3004 1.4393

Autohydrolysis-190C .96618* .01732 .8967 1.0356

Autohydrolysis-205 C 1.15265* .01732 1.0832 1.2221

Water treated- 110C 1.22224* .01732 1.1528 1.2917


PhD Thesis

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G.3.1. Analysis of variance for the production of bioethanol from pretreated

bagasse by using various yeast strains

Sum of

Squares df

Mean

Square F Sig.

Between

Groups 24820.451 4 6205.113 178.043 .000

Within Groups 1219.809 35 34.852

Total 26040.260 39


PhD Thesis

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G.3.2. Post hoc tukey HSD multiple comparisons for the production of bioethanol from pretreated bagasse by using various

yeast strains

(I) Pretreatment (J) Pretreatment

Mean

Difference

(I-J) S.D Sig.

95% Confidence Interval

Lower

Bound Upper Bound

Untreated

control

Water treated 110C -9.41500* 2.95177 .023 -17.9015 -.9285

IL-110C -58.35875* 2.95177 .000 -66.8453 -49.8722

Autohydrolysis-190 C -46.21875* 2.95177 .000 -54.7053 -37.7322

Autohydrolysis 205C -58.51000* 2.95177 .000 -66.9965 -50.0235

Water

treated 110C

Untreated control 9.41500* 2.95177 .023 .9285 17.9015

IL-110C -48.94375* 2.95177 .000 -57.4303 -40.4572

Autohydrolysis-190 C -36.80375* 2.95177 .000 -45.2903 -28.3172


IL-110C Untreated control 58.35875* 2.95177 .000 49.8722 66.8453

Water treated 110C 48.94375* 2.95177 .000 40.4572 57.4303

Autohydrolysis-190 C 12.14000* 2.95177 .002 3.6535 20.6265

Autohydrolysis 205C -.15125 2.95177 1.000 -8.6378 8.3353

Autohydroly

sis-190 C

Untreated control 46.21875* 2.95177 .000 37.7322 54.7053

Water treated 110C 36.80375* 2.95177 .000 28.3172 45.2903

IL-110C -12.14000* 2.95177 .002 -20.6265 -3.6535


Autohydroly

sis 205C

Untreated control 58.51000* 2.95177 .000 50.0235 66.9965

Water treated 110C 49.09500* 2.95177 .000 40.6085 57.5815

IL-110C .15125 2.95177 1.000 -8.3353 8.6378

Autohydrolysis-190 C 12.29125* 2.95177 .002 3.8047 20.7778


PhD Thesis

189

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