220
MICROENCAPSULATION OF CLOSTRIDIUM ACETOBUTYLICUM CELLS AND UTILISATION OF SAMANEA SAMAN LEAF LITTER FOR THE PRODUCTION OF BIOBUTANOL SWETA RATHORE NATIONAL UNIVERSITY OF SINGAPORE 2013
MICROENCAPSULATION OF CLOSTRIDIUM ACETOBUTYLICUM
CELLS AND UTILISATION OF SAMANEA SAMAN LEAF LITTER
FOR THE PRODUCTION OF BIOBUTANOL
SWETA RATHORE
NATIONAL UNIVERSITY OF SINGAPORE
2013
MICROENCAPSULATION OF CLOSTRIDIUM ACETOBUTYLICUM
CELLS AND UTILISATION OF SAMANEA SAMAN LEAF LITTER FOR
THE PRODUCTION OF BIOBUTANOL
SWETA RATHORE
(B.Sc. (Pharm), Mumbai University)
A THESIS SUBMITTED
FOR THE DEGREE OF DOCTOR OF PHILOSOPHY
DEPARTMENT OF PHARMACY
NATIONAL UNIVERSITY OF SINGAPORE
2013
Declaration
I hereby declare that this thesis is my original work and it has been written by
me in its entirety. I have duly acknowledged all the sources of information
which have been used in the thesis. This thesis has also not been submitted for
any degree in any university previously.
ACKNOWLEDGEMENTS
I consider this as the most important page of my entire thesis as I list the
names of all the people who have, in some way or the other, helped me reach
the end of the scientific adventure that I ventured four years back. First and
foremost, I would like to thank my supervisors, Associate Professor Chan Lai
Wah and Associate Professor Paul Heng Wan Sia, for their attentive
supervisor and invaluable guidance. This thesis would not have been possible
without their encouragement and support.
I am also grateful to National University of Singapore for providing me the
opportunity and infrastructure to carry out my research work. Special thanks
to the laboratory technologists, Mdm Teresa Ang, Ms Yong Sock Leng and
Mdm Wong Mei Yin for providing technical and logistic assistance from time
to time. I am thankful to my fellow GEANUS friends, past and present as well
as the FYP students, Alvin, Jeanette and Eileen for helping with a part of this
project.
And last but not the least; I would like to express my heartfelt gratitude to the
pillars of my life, my family. Their patience and support has motivated to face
all the challenges in the four years with self-belief and positive attitude.
Overall, this PhD journey has been an enriching experience inculcating in me
to have a broader outlook towards science as well as life.
Sweta Rathore
2013
i
CONTENTS
SUMMARY………………………………………………………………...viii
LIST OF TABLES ........................................................................................... x
LIST OF FIGURES ....................................................................................... xii
I. INTRODUCTION ........................................................................................ 2
A. Biofuel ................................................................................................ 2
A.1 Biobutanol .................................................................................. 2
B. Biobutanol production ......................................................................... 3
B.1 Clostridium acetobutylicum ......................................................... 4
B.2 ABE fermentation ........................................................................ 5
B.3 Morphological changes in Cl. acetobutylicum during
ABE fermentation ........................................................................ 7
B.4 Limitations of the conventional ABE batch fermentation
process ......................................................................................... 8
C. Strategies to overcome limitations of ABE fermentation .................. 10
C.1 Solvent recovery ........................................................................ 10
C.2 Genetic/metabolic engineering .................................................. 12
C.3 Advanced fermentation techniques ............................................ 14
D. Cell immobilisation ........................................................................... 15
D.1 Immobilisation of solventogenic clostridia ................................ 16
ii
D.2 Limitations of conventional cell immobilisation methods used
in ABE fermentation................................................................... 18
E. Microencapsulation as a cell immobilisation technique .................... 19
E.1 Techniques used for microencapsulation of microbial cells ...... 20
E.2 Polymers used for microencapsulation ...................................... 23
F. Alternative fermentation substrates .................................................... 28
F.1 Samanea saman tree (rain tree) .................................................. 29
F.2 Structure of lignocellulosic substrate ......................................... 31
F.3 Pretreatment of lignocellulosic substrate ................................... 34
F.4 Types of pretreatment................................................................. 35
F.5 Enzymatic hydrolysis of lignocellulosic substrate ..................... 37
F.6 Strategies for detoxification of acid hydrolysate ........................ 38
II. HYPOTHESES AND OBJECTIVES ..................................................... 42
III. EXPERIMENTAL .................................................................................. 47
A. Materials ............................................................................................ 47
A.1 Model microorganism ................................................................ 47
A.2 Growth media ............................................................................ 47
A.3 Fermentation medium ................................................................ 48
A.4 Encapsulating polymer and chemicals ...................................... 48
A.5 Chemicals for assay of butanol by gas chromatography
iii
–mass spectrometry .................................................................... 48
A.6 Lignocellulosic substrate .......................................................... 49
A.7 Cellulolytic enzyme .................................................................. 49
A.8 Chemicals used in assay of reducing sugars ............................. 49
A.9 Chemicals used for dilute acid coupled with heat treatment
of S. saman leaf litter……………………………….…..........49
A.10 Chemicals used for measuring the filter paper units (FPU)
activity of Accellerase® 1500 ................................................. 50
A.11 Chemicals used for detoxification of acid hydrolysate of
S. saman leaf litter ................................................................... 50
B. METHODS........................................................................................ 51
B.1 Preparation of growth media ...................................................... 51
B.2 Cultivation of Cl. acetobutylicum ATCC 824 ........................... 51
B.2.1 Revival of Cl. acetobutylicum ATCC 824 ........................... 51
B.2.2 Determination of suitable media for the growth
of Cl. acetobutylicum ATCC 824 ........................................ 51
B.2.3 Determination of suitable anaerobic set-up for the growth
of Cl. acetobutylicum ATCC 824 ........................................ 52
B.2.4 Determination of growth curve and morphology
of Cl. acetobutylicum ATCC 824 .......................................... 53
iv
B.2.5 Preparation of spore stock culture of Cl. acetobutylicum
ATCC 824 ............................................................................ 54
B.2.6 Optimisation of heat shock treatment (HST) conditions
for the revival of Cl. acetobutylicum ATCC 824 spores ...... 55
B.2.7 Preparation of standardised inoculum of vegetative cells
of Cl. acetobutylicum ATCC 824 ........................................ 56
B.3 Production of microspheres by emulsification method .................. 57
B.3.1 Optimisation of production of gellan gum microspheres .... 58
B.3.2 Characterisation of the microspheres .................................. 62
B.4 Study of emulsification process on viability of
Cl. acetobutylicum ATCC 824 vegetative cells/spores ................. 63
B.5 Method development for the assay of butanol
by gas chromatography-mass spectrometry (GC-MS) ................... 63
B.6 Fermentation studies using Cl. acetobutylicum ATCC 824 cells ... 66
B.7 Determination of viable count of cells liberated from
microspheres into the fermentation medium .................................. 69
B.8 Comparison of reusability between free (non-encapsulated)
cells and encapsulated cells of Cl. acetobutylicum ATCC 824 ...... 70
B.9 Pretreatment of S. saman leaf litter ................................................ 70
B.10 Assay of fermentable sugars by DNS method .............................. 74
v
B.11 Determination of filter paper activity of Accellerase® 1500 ....... 76
B.12 Enzymatic hydrolysis of pretreated S. saman leaf litter ............... 77
B.13 Detoxification of acid hydrolysate of S. saman leaf litter ............ 78
B.14 Fermentation of detoxified leaf hydrolysate by
Cl. acetobutylicum ATCC 824 ..................................................... 79
B.15 Statistical analysis ......................................................................... 80
IV. RESULTS AND DISCUSSION .............................................................. 82
PART ONE ..................................................................................................... 82
A. Cultivation of Cl. acetobutylicum ATCC 824 .................................... 82
A.1 Suitable media for the growth of Cl. acetobutylicum ATCC 824
………………………………………………………………….83
A.2 Suitable set-up for the growth of Cl. acetobutylicum
ATCC 824 .................................................................................. 86
A.3 Growth curve of Cl. acetobutylicum ATCC 824 in RCM .......... 89
A.4 Morphological changes in Cl. acetobutylicum ATCC 824
cells during different phases of growth ..................................... 92
A.5 Optimisation of heat shock treatment for the revival
of Cl. acetobutylicum ATCC 824 spores ................................... 94
B. Optimisation of microsphere production using Design
of Experiments (DoE) ...................................................................... 96
vi
B.1 Influence of the variables on size .......................................... 100
B.2 Influence of the variables on span ......................................... 101
B.3 Influence of the variables on aggregation index .................... 102
B.4 Model equations and model adequacy ................................... 102
B.5 Optimisation of formulation and process parameters in the
production of microspheres with the desired properties ........ 107
C. Effect of emulsification process on viability of Cl. acetobutylicum
ATCC 824 vegetative cells and spores ........................................... 111
D. Microencapsulation of Cl. acetobutylicum ATCC 824
spores by emulsification method .................................................... 114
E. Optimisation of gas chromatography-mass spectrometry
conditions for the assay of butanol ................................................. 115
F. Fermentation using free (non-encapsulated) cells of
Cl. acetobutylicum ATCC 824 ...................................................... 117
F.1 Influence of glucose on fermentation efficiency .................. 118
F.2 Influence of inocula age ....................................................... 120
F.3 Influence of inocula size ...................................................... 121
G. Fermentation using encapsulated spores of Cl. acetobutylicum
ATCC 824 ..................................................................................... 124
H. Cell leakage from gellan gum microspheres ................................. 127
vii
H.1 Contribution by liberated cells to butanol production ........... 128
I. Reusability of free (non-encapsulated) vegetative cells/spores
and encapsulated spores of Cl. acetobutylicum ATCC 824 ........... 135
PART TWO .................................................................................................. 143
A. Potential of Samanea saman leaf litter as a source of
fermentable sugars for biobutanol production ............................... 143
A.1 Recovery of total fermentable sugars from S. saman leaves .. 144
A.2 Determination of filter paper activity of Accellerase® 1500 . 145
A.3 Pretreatment of S. saman leaf litter ........................................ 147
A.4 Detoxification of acid hydrolysate of S. saman leaf litter ...... 164
A.5 Fermentation of detoxified leaf hydrolysate by free
(non- encapsulated) and encapsulated cells of
Cl. acetobutylicum ATCC 824 ................................................ 167
V. CONCLUSIONS ..................................................................................... 173
VI. REFERENCES ...................................................................................... 176
VII. LIST OF PUBLICATIONS ................................................................ 200
viii
SUMMARY
The purpose of the present study was to provide insights on applicability of
microencapsulation using gellan gum, as a cell immobilisation method for
Clostridium acetobutylicum ATCC 824 cells for biobutanol production.
Secondly, an investigation on the use of leaf litter from Samanea saman tree,
as a lignocellulosic substrate for biobutanol production, was attempted. The
combination of these methods were aimed to address the issues of low butanol
yield and high production cost of biobutanol production
The factors affecting the production of gellan gum microspheres by
emulsification technique were investigated using full factorial design,
followed by derivation of optimised process conditions. The viability of Cl.
acetobutylicum ATCC 824 cells was adversely affected by the emulsification
process. The spore form was more suitable and successfully encapsulated in
gellan gum microspheres using optimised process conditions. Encapsulated
spores were revived by heat shock treatment at 90 °C for 10 min prior to use
in fermentation. The microspheres could be easily recovered from the
fermentation media and reused up to five cycles of fermentation. In contrast,
the free (non-encapsulated) cells could be used for two cycles only. The
microspheres remained intact throughout repeated use. The fermentation
efficiency of the encapsulated spores was lower than that of free (non-
encapsulated) cells during the first fermentation cycle. This was attributed to
lag time for revival of the spores and acclimatisation of the cells to the
microenvironment. In addition, presence of the encapsulating polymer matrix
ix
also caused impairment of mass transfer, prolonging the fermentation time to
achieve maximum butanol yield. The fermentation efficiency of the
encapsulated spores was however much higher than that of the free cells in
subsequent cycles. Significant cell leakage from the microspheres was
observed at the end of the fermentation process. The microspheres served as
nurseries for the generation of new cells. Both encapsulated and liberated cells
contributed to butanol production.
The potential of S. saman leaf litter, as a readily available lignocellulosic
substrate for biobutanol production, was explored in the second part of the
project. Due to the resistant structure of any lignocellulosic substrate,
pretreatment of the substrate is prerequisite. The pretreatment methods
investigated were milling, hydrothermal treatment and dilute acid coupled
with heat treatment. It was found that milling alone or in combination with
hydrothermal treatment was inefficient. However, dilute acid coupled with
heat treatment could recover substantial quantities of sugar from the leaf litter.
This process was further optimised by the response surface methodology.
Various detoxification methods for the pretreated leaf litter were also
investigated. Using sodium hydroxide neutralisation, the acid hydrolysate was
effectively detoxified and could be used as a fermentation substrate for
biobutanol production by both free and encapsulated Cl. acetobutylicum
ATCC 824 cells.
x
LIST OF TABLES
Table 1 Physical properties of butanol and other fuels .................................. 4
Table 2 Summary of various cell immobilisation methods employed
in ABE fermentation process ......................................................... 17
Table 3 HST optimisation of Cl. acetobutylicum ATCC 824 ...................... 55
Table 4 Coded and uncoded values of the two independent factors
in the optimisation of microencapsulation process ........................ 59
Table 5 Response variables used in the 24
full factorial design .................. 60
Table 6 Different combinations of parameters investigated in the
optimisation of fermentation by free vegetative cells
of Cl. acetobutylicum ATCC 824 .................................................. 67
Table 7 Different combinations of parameters investigated
in the optimisation of fermentation by encapsulated spores
of Cl. acetobutylicum ATCC 824 .................................................. 68
Table 8 Coded and uncoded values of the two independent factors
in the optimisation of dilute acid coupled with heat treatment ...... 72
Table 9 Preparation of different dilutions of glucose for standard
calibration curve............................................................................ 76
Table 10 Viable count of Cl. acetobutylicum ATCC 824 in
different cultivation broths ............................................................ 85
Table 11 Effects of different incubation set-ups on the cells growth
of Cl. acetobutylicum ATCC 824 ................................................. 88
Table 12 Viable count of Cl. acetobutylicum ATCC 824 spores
xi
revived by different heat shock treatment conditions .................... 96
Table 13 Factorial design matrix employed in the optimisation study
for the microencapsulation process ................................................ 98
Table 14 Coefficient estimate, sum of squares and their respective
p-values for the three responses ................................................... 106
Table 15 Effect of emulsification process on the viability of vegetative
cells and spores of Cl. acetobutylicum ATCC 824 ....................... 112
Table 16 Results from the fermentation optimisation studies of free
(non-encapsulated) cells of Cl. acetobutylicum ATCC 824 ......... 123
Table 17 Results from the fermentation optimisation studies of
encapsulated spores of Cl. acetobutylicum ATCC 824 ................ 126
Table 18 Experimental design used for the optimisation of dilute
acid coupled with heat treatment along with the values of the
response variables…………………………………………….…155
Table 19 ANOVA table for yield and recovery of fermentable sugars
in dilute acid coupled with heat treatment ...... …………………..157
xii
LIST OF FIGURES
Figure 1 Scanning electron microscopic image of vegetative cells
of Cl. acetobutylicum ....................................................................... 5
Figure 2 Biochemical pathway of ABE fermentation..................................... 6
Figure 3 Morphological changes in Cl. acetobutylicum ................................. 8
Figure 4 Chemical structures of (a) acetylated gellan gum and
(b) deacetylated gellan gum ........................................................... 28
Figure 5 Different components of the S. saman tree: (a) leaves,
(b) pods, (c) leaf litter and (d) flowers .......................................... 31
Figure 6 Structure of lignocellulose ............................................... ………. 32
Figure 7 Chemical structures of (a) cellulose, (b) hemicellulose and
(c) lignin………………………………………………….………32
Figure 8 Pretreatment of lignocellulose ...................................................... 35
Figure 9 Schematic diagram of quantification of viable cells
by spread plate method ........................................................ ……54
Figure 10 Production of gellan gum microspheres using emulsification
method ......................................................................................... 61
Figure 11 Cl. acetobutylicum ATCC 824 colonies on RCM agar after
24 h of incubation at 37 °C .......................................................... 85
Figure 12 Cl. acetobutylicum ATCC 824 colonies on TGM agar after
48 h of incubation at 37 °C .......................................................... 85
Figure 13 Growth of Cl. acetobutylicum ATCC 824 in (a) RCM agar
and (b) RCM broth incubated under anaerobic
xiii
conditions maintained by Anaerogen™ ....................................... 89
Figure 14 Cultivation of Cl. acetobutylicum ATCC 824 in RCM broth
at 37 °C: (a) growth curve and (b) optical density of culture…. . 91
Figure 15 Relationship between optical density and viable count of
Cl. acetobutylicum ATCC 824 culture in RCM broth…..……....91
Figure 16 Gram staining of Cl. acetobutylicum ATCC 824 cells in
(a) exponential, (b) stationary and (c) decline phase
of growth………………………………………………………...93
Figure 17 Response surface plots of the effects of (a) temperature
and concentration of gellan gum on size, (b) concentration
of gellan gum and stirring speed on size, (c) stirring speed and
HLB on span and (d) concentration of gellan gum and stirring
speed on aggregation index…………………………………….104
Figure 18 Correlation between observed and predicted values for
(a) size, (b) span and (c) aggregation index of microspheres….110
Figure 19 Photographs of (a) blank gellan gum microspheres and
(b) gellan gum microspheres loaded with Cl. acetobutylicum
ATCC 824 spores prepared using the optimised
microencapsulation method…………………………………….115
Figure 20 Calibration plot for estimation of butanol concentration in
fermentation medium………………………………………......117
Figure 21 Photograph of liberated Cl. acetobutylicum ATCC 824
cells from gellan gum microspheres .......................................... 127
Figure 22 Viability profiles of Cl. acetobutylicum ATCC 824 cells
xiv
liberated from gellan gum microspheres and fermentation
profile of encapsulated and liberated cells ................................. 129
Figure 23 Viability and fermentation profiles of free Cl. acetobutylicum
ATCC 824 cells (equivalent to the number of liberated cells
from the microspheres during the course of fermentation) ........ 133
Figure 24 Photographs of microspheres with Cl. acetobutylicum
ATCC 824 cells at the periphery of gellan gum microspheres .. 133
Figure 25 Photographs of gellan gum microspheres recovered
from fermentation after (a) 24 h, (b) 48 h, (c) 72 h,
(d) 96 h, (e) 120 h and (f) 144 h of fermentation ........................ 134
Figure 26 Plot of fermentation profile of free cells vs. encapsulated
cells in first fermentation cycle .................................................. 139
Figure 27 Photographs of gellan gum microspheres recovered
from fermentation medium after (a) 1 cycle, (b) 2 cycles,
(c) 3 cycles, (d) 4 cycles and (e) 5 cycles of fermentation ........ 142
Figure 28 Calibration curve of glucose ...................................................... 146
Figure 29 Plot of enzyme concentration vs. glucose concentration ........... 147
Figure 30 Leaf litter of S. saman: (a) before milling, (b) after using
hammer mill, (c) followed by disintegrator mill ....................... 150
Figure 31 Relationship between sugar recovery and Accellerase®
1500 dose .................................................................................. 151
Figure 32 Effect of hydrothermal pretreatment on sugar recovery
from S. saman leaf litter before and after enzyme addition ...... 152
Figure 33 Sugar recovery from milled S. saman leaf litter subjected to
xv
1.0 %, w/w acid for different treatment times ........................ 154
Figure 34 Surface plots for effects of treatment time and acid
concentration on (a) sugar yield and (b) sugar recovery .......... 159
Figure 35 Contour plot for optimisation of the dilute acid coupled
with heat treatment of milled S. saman leaf litter……………160
Figure 36 Comparison of (a) total sugar recovery from S. saman leaf
litter after both dilute acid coupled with heat treatment and
enzyme hydrolysis (b) total sugar recovery due to enzymatic
hydrolysis alone after different dilute acid coupled with
heat treatment conditions …………………………..……...…163
Figure 37 Butanol yield achieved from the acid hydrolysate of
S. saman leaf litter subjected to different detoxification
methods .................................................................................... 165
Figure 38 Fermentation of detoxified acid hydrolysate by free
and encapsulated Cl. acetobutylicum ATCC 824 cells ............ 169
Figure 39 Viability profile of free and encapsulated cells of
Cl. acetobutylicum ATCC 824 during fermentation of
detoxified acid hydrolysate of S. saman leaf litter ................... 170
1
INTRODUCTION
2
I. INTRODUCTION
A. Biofuel
Most of the current global energy requirements are met by fossil fuels, such as
petroleum oil, coal and natural gas, which originate from deceased organisms
that lived several million years ago (Weiland, 2010). Due to growing energy
demands, the global consumption of these limited fossil fuel reserves has
increased tremendously (Finley, 2012). In addition, burning of fossil fuels has
caused a net increase in carbon dioxide levels, resulting in global warming
effect (Dürre, 2007). These factors have played a crucial role in the increased
interest to produce biofuels as an alternative to fossil fuels (García et al., 2011;
Lynd et al., 2008; Srirangan et al., 2012). A biofuel is defined as any liquid or
gaseous transportation fuel, originating from a biological source (Giampietro
et al., 1997). Unlike fossil fuels, they are renewable and cleaner sources of
energy owing to their carbon neutral attribute as the raw materials used to
produce the biofuel consume as much carbon dioxide as the biofuel contributes
during its combustion (Demirbas, 2005). Examples of biofuels include
biodiesel, bioethanol, biomethanol and biobutanol (Fatih Demirbas et al.,
2011). Amongst these, only biodiesel and bioethanol have been
commercialised (Hess, 2006).
A.1 Biobutanol
Butanol, acetone, ethanol and isopropanol are naturally formed by a number of
solventogenic clostridia from fermentation of various biomass raw materials
3
like molasses, whey permeate and corn (Ezeji et al., 2007a; Lee et al., 2008;
Qureshi et al., 2008). The term biobutanol is used to indicate that the butanol
is derived from fermentation instead of petrochemical processes. Due to its
similarity to gasoline, biobutanol has shown to be a promising biofuel (Table
1). Compared to ethanol and methanol, butanol can be blended at higher
concentrations with gasoline for use in standard vehicle engines and is well-
suited to current vehicle and engine technologies (Campos-Fernández et al.,
2012; Dürre, 2007; Lee et al., 2008). The low vapour pressure of butanol
makes it less volatile and more prone to combustion (Lütke-Eversloh and
Bahl, 2011). It can be transported using the existing fuel distribution
infrastructure and is less susceptible to separation in the presence of water than
other biofuel-gasoline blends (Lee et al., 2008). Another major advantage of
biobutanol is the wide range of feedstock that can be utilised for its production
(Kumar and Gayen, 2011). According to a recent report, biobutanol
production ranged from 10 - 12 billion pounds per year, with a value of US$ 7
- 8.4 billion (Lee et al., 2008). Improvement in biobutanol production will
escalate the market demand further.
B. Biobutanol production
The production of solvents by strains of solventogenic clostridia is known as
“Acetone Butanol Ethanol fermentation” or “ABE fermentation” (Zverlov,
2006). ABE fermentation, using corn and molasses, was the second largest
industrial fermentation in the early part of 20th century (Cheng et al., 2012;
Dürre, 2007). It eventually became obsolete as the cost of fermentation
4
feedstocks increased and more efficient and cheaper petrochemical processes
for butanol production were available (García et al., 2011). However, with
depleting fossil fuels, as well as the advancement in biotechnological
processes and development of innovative fermentation process technologies,
the interest in fermentative butanol production has gained momentum once
again (Kumar and Gayen, 2011).
Table 1 . Physical properties of butanol and other fuels
Properties Butanol Gasoline Methanol Ethanol
Boiling point (°C) 117.7 27-221 65 78.5
Miscibility with water immiscible immiscible miscible miscible
Explosive limits (vol. %
in air)
1.7-12 1-8 6-31 3.3-19
Energy density (MJ/L) 29.2 32 19.6 16
Energy content
(BTU/gal)
110000 115000 84000 76000
Air-fuel ratio 11.1 14.6 6.4 9.0
Heat of vaporisation
(MJ/kg)
0.43 0.36 1.2 0.92
Adapted from Dürre, 2007; García et al., 2011; Kumar and Gayen, 2011; Lee
et al., 2008
B.1 Clostridium acetobutylicum
Amongst the different strains of solventogenic clostridia, Cl. acetobutylicum
ATCC 824 is reported to display superiority in biobutanol production (Lee et
5
al., 2008). Hence, it has been extensively used in both the investigation and
production of biobutanol. Cl. acetobutylicum is a Gram-positive anaerobic
bacterium that can ferment a wide variety of carbon substrates, such as
glucose, xylose, pentose and starch, to industrially useful solvents such as
acetone, butanol and ethanol (Dürre, 2007; Jones and Woods, 1986). This
bacterium has peritrichous flagella for motility and produces sub-terminal
endospores. Vegetative cells of Cl. acetobutylicum are straight rods of 2.4 -
4.7 microns by 0.6 - 0.9 microns in size (Smith and Hobbs, 1974) (Figure 1).
The spores are oval and resistant to adverse environmental factors, such as
heat, desiccation and aerobic conditions.
Figure 1. Scanning electron microscopic image of vegetative cells of Cl.
acetobutylicum
B.2 ABE fermentation
Much work has been conducted to understand the ABE fermentation that Cl.
acetobutylicum undergoes. A typical feature of the ABE fermentation is its
biphasic nature (Lee et al., 2008). ABE fermentation consists of an acidogenic
6
phase, followed by a solventogenic phase (Figure 2). During the acidogenic
phase, which corresponds to the exponential growth phase, acetic acid, butyric
acid and lactic acid are produced as major products from the metabolised
carbon source (Sukumaran et al., 2011).
Figure 2. Biochemical pathway of ABE fermentation
The synthesis of acids has been found to be essential for cell growth and
metabolism (Ezeji et al., 2010). However, these acidic products cause a
gradual decline in the pH of the culture medium, resulting in the cells
switching from the acidogenic phase to the solventogenic phase. A threshold
value of 60 mmol/L of acid has been found to trigger phase shift to
solventogenesis (Maddox et al., 2000; Zheng et al., 2009b). During the
second phase of the fermentation, which corresponds to the stationary phase of
the bacterium, acids are reassimilated and used in the production of acetone,
7
butanol and ethanol along with concomitant increase in the pH of the culture
medium (Dürre, 2007). It has been suggested that the uptake of acid during
solventogenic phase functions as a detoxification process in response to the
accumulation of acidic end products (Long et al., 1984). Depending on the
strain and inoculum size, as well as the substrate used, it takes 48 - 144 h to
complete batch fermentation and the final total concentration of solvents
produced is in the range of 12 to 20 g/L of fermentation medium (Lee et al.,
2008). The solvents can be separated from the fermentation medium by
distillation. Solvent ratios vary according to the strain and fermentation
conditions, with a ratio of 3:6:1 (acetone: butanol: ethanol) being typical in
ABE fermentation (Qureshi and Maddox, 2005; Sukumaran et al., 2011).
B.3 Morphological changes in Cl. acetobutylicum during ABE
fermentation
The cells of Cl. acetobutylicum exist in different morphological structures
during the course of fermentation (Long et al., 1984) (Figure 3). The
vegetative cells in the exponential stage are rod-shaped and predominantly
involved in acid formation. As the cells enter the stationary phase, they begin
to swell and accumulate reserve material in the form of granulose. These cells
in this phase are known as clostridial cells, which subsequently form the
forespore cells (mother cells containing the endospores). The cells in the
stationary phase are involved in conversion of acids to solvents. Finally, the
endospore is released from the forespore cells. When provided with favourable
conditions, these spores can be revived into vegetative form.
8
Figure 3. Morphological changes in Cl. acetobutylicum
B.4 Limitations of the conventional ABE batch fermentation process
Production of biobutanol faces some major limitations which have hindered its
large scale application. Inherent problems associated with ABE fermentation
include solvent toxicity, low product concentrations and volumetric
productivity, high cost of product recovery, complex metabolic pathways and
the high cost of substrates (Chauvatcharin et al., 1998; Dürre, 1998).
Solvent toxicity is one of the most critical limitations of ABE fermentation
(Ezeji et al., 2003). It has been found that the cellular metabolism of the
solventogenic clostridia diminishes when the total solvent concentration
reaches around 20 g/L (Lee et al., 2008; Qureshi and Blaschek, 2001; Woods,
1995). In addition, due to solvent toxicity, cell viability is decreased which
9
further reduces the overall productivity of the fermentation process. Butanol
has been found to be the most toxic amongst the three solvents produced in
ABE fermentation due to its lipophilic nature (Lee et al., 2008). It disrupts the
phospholipid components of the cell membrane, resulting in increased
membrane fluidity (Bowles and Ellefson, 1985; Liu and Qureshi, 2009).
Butanol exposure of as low as 10 g/L has been shown to increase the
membrane fluidity by almost 20 - 30 % (Liu and Qureshi, 2009). Increase in
membrane fluidity affects various membrane-related cellular functions such as
preferential solute transport, glucose uptake, maintenance of the proton motive
force (or maintenance of intracellular pH) and intracellular ATP level
(Moreira et al., 1981). These observations have been confirmed in a number
of laboratory studies, and it has been shown that the addition of 7 - 13 g/L of
butanol to cultures growing on hexose sugars resulted in a 50 % inhibition of
growth (Jones and Woods, 1986). Recovery of the relatively little butanol
produced, by distillation is energy intensive and increases the overall cost of
the process.
Apart from the low yields and productivity due to solvent toxicity, the high
cost of the fermentation substrates for ABE fermentation is another area of
concern. Traditionally, food crops or food by-products like corn, potatoes,
maize and molasses were used as fermentation substrates (Jones and Woods,
1986). Increase in the demand and price of these food crops has hindered the
large scale economical production of biobutanol.
10
C. Strategies to overcome limitations of ABE fermentation
Various approaches have been employed to overcome the limitation of solvent
toxicity, such as in situ solvent recovery, strain improvement by
genetic/metabolic manipulation, improvement of butanol tolerance using
different fermentation modes such as fed-batch fermentation and continuous
fermentation and utilisation of immobilised cells (Sukumaran et al., 2011). In
addition, alternative fermentation substrates derived from lignocellulose-based
feedstock and waste materials have also been utilised so as to reduce the
overall cost of the fermentation process (Papoutsakis, 2008).
C.1 Solvent Recovery
The conventional method of solvent recovery from ABE fermentation medium
is distillation (Dürre, 2007). However it is usually energy intensive and
involves high operation costs due to low solvent concentrations obtained
during ABE fermentation (Qureshi et al., 2005). In order to overcome these
problems, in addition to solvent toxicity, many in situ solvent recovery
techniques for butanol removal have been investigated, such as liquid-liquid
extraction, perstraction and gas stripping (Ezeji et al., 2004; Groot et al., 1984;
Ishii et al., 1985). The in situ recovery methods are able to improve the solvent
yield and productivity of the ABE fermentation process. However, they
require high capital costs apart from the inherent limitations associated with
each method (Qureshi and Blaschek, 2001).
11
Liquid-liquid extraction technique utilises a water-immiscible organic solvent
to extract the solvents produced (i.e. acetone, butanol and ethanol) from the
ABE fermentation medium. The extractant is mixed with the fermentation
medium. Being more soluble in the organic phase, the solvents partition into
the organic extractant. The extracted solvents are then isolated by distillation.
Examples of common extractants used include decanol and oleyl alcohol.
Though liquid-liquid extraction is a simple technique, it faces serious
problems of toxicity of the extractant to the cells and emulsion formation
(Qureshi, 1992; Takriff et al., 2008). Thus, finding an extracting solvent with
suitable distribution coefficient, low cost and low toxicity is difficult.
Perstraction is a modification of liquid-liquid extraction method in which the
fermentation medium and the extractant are separated by a membrane. This
membrane is impermeable to the extractant but permeable to the solvent of
interest. Preferential diffusion of butanol occurs across the membrane, leaving
behind other fermentation intermediates in the aqueous fermentation medium
(Qureshi and Maddox, 2005). Separation of the aqueous fermentation phase
from the organic extractant phase eliminates or drastically reduces problems
such as extractant toxicity, emulsion formation and phase dispersion (Ezeji et
al., 2007a). However, fouling and clogging of the membranes have been
reported as the major limitations of this method (Dürre, 2007). Furthermore,
the membrane acts as a physical barrier that can limit the rate of butanol
extraction (Ezeji et al., 2007a).
12
In gas stripping technique, either nitrogen or the fermentation gases (carbon
dioxide and hydrogen) are sparged into the fermentation medium. The
formation and bursting of the gas bubbles cause the surrounding fermentation
liquid to vibrate and the gases capture the volatile butanol. The gases are then
separated from the fermentation medium and butanol isolated by condensation
(Ezeji et al., 2007a; Zheng et al., 2009b). The gases are recycled back to the
fermentation medium to capture more butanol. This process continues until all
the sugar in the fermentation medium is utilised. Gas stripping is an effective
solvent recovery method as the bacterial cells are not harmed and the products
can be easily recovered with low energy consumption (Ezeji et al., 2003).
However, foam formation during gas stripping may pose a problem (Ezeji et
al., 2005).
C.2 Genetic/metabolic engineering
Genetic/metabolic engineering of solventogenic clostridia aims to increase
butanol (solvent) tolerance of the organism, extend substrate utilisation range
and allow selective production of butanol instead of mixed acids/solvents
production (Dürre, 2007; Ezeji et al., 2007a; Huang et al., 2010).
Genetic mutation has been carried out by either spontaneous alteration,
exposure of wild type strain to chemical mutagens or high butanol
concentrations (Harris et al., 2001; Mermelstein et al., 1993). Different
mutagenic agents, such as hydrogen peroxide, nalidixic acid, metronidazole,
ethyl methanesulfonate, N-methyl-N-nitro-N-nitrosoguanidine and UV
irradiation have been used to induce mutation in solventogenic clostridia
13
(Ezeji et al., 2007a). A mutant strain of Cl. acetobutylicum ATCC 824 was
developed by exposing the original strain to n-butanol. The developed strain
had higher butanol tolerance (121% higher) over the native strain (Lin and
Blaschek, 1983). In another study, treatment of parent strain with N-methyl-N-
nitro-N-nitrosoguanidine, ethyl methane sulphonate and UV radiation yielded
a novel mutated strain (MEMS-7) which possessed better substrate (molasses)
utilisation property and produced 20 % higher butanol yield than the parent
strain (Syed et al., 2008).
Several other organisms such as Escherichia coli were also employed as hosts
for introduction of butanol producing genes so as to circumvent the problems
associated with the growth and cultivation of clostridial species (Nielsen et al.,
2009; Zheng et al., 2009b). However, the solvent yield obtained from these
organisms was found to be relatively low.
Clostridial species often have a complex physiology that is not well-
understood and genetic tools and strategies for improving the productivity of
these species are still under development (Dürre, 2011; Lütke-Eversloh and
Bahl, 2011; Patakova et al., 2012). In spite of several attempts to improve the
industrial strains by mutation and genetic manipulations, the highest butanol
titre achieved so far was between 19 to 20 g/L using the strain C. beijerinckii
BA101 (Formanek et al., 1997; Qureshi et al., 2008).
14
C.3 Advanced fermentation techniques
Conventionally, batch fermentation is the preferred mode of fermentation due
to its simple and stable operation, high efficiency and low risk of
contamination (Cardona and Sánchez, 2007; Sukumaran et al., 2011).
However, the productivity attainable in a batch reactor is generally low (0.5
g/L/h) due to low cell viability, product inhibition effects as well as downtime
for harvesting, cleaning, sterilising, and re-filling the reactor (Ezeji et al.,
2006; García et al., 2011). Moreover, a low viable cell concentration
(typically, less than 4 g/L) is achieved in batch fermentation (Ezeji et al.,
2007b; Qureshi and Blaschek, 2001). This is attributed to the formation of
metabolic products during the fermentation which can adversely affect cell
viability. Higher cell densities can be achieved using advanced techniques
such as cell immobilisation or cell recycle systems (Ezeji et al., 2006; García
et al., 2011; Qureshi and Maddox, 1987). Cell immobilisation refers to the
physical confinement of whole cells to a certain defined region of space while
preserving their activity for repeated or continuous use (Karel et al., 1985). In
a cell recycle system, the cells and the fermentation products are first
separated using a filter and then the cells are returned to the fermentor
(Tashiro et al., 2005). The separation of the cells from the toxic metabolic
products in the fermentation medium allows attainment of high viable cell
densities compared to conventional batch cell fermentation (Kleman and
Strohl, 1992). Utilisation of immobilised cell cultures can increase the reactor
15
productivity about 40-50 times compared to batch reactors utilising free cells
(Ezeji et al., 2004).
D. Cell immobilisation
The pioneering work in cell immobilisation was carried out by (Chibata, 1979)
using immobilised microbial cell preparation for the continuous production of
L-aspartic acid. Since then, cell immobilisation has been employed in a wide
range of biotechnological applications in different industries, such as food
production, agriculture, pest control and biofuels (Amiet-Charpentier, 1999;
Badr et al., 2001; Häggström and Molin, 1980; Kailasapathy, 2002; Reid et al.,
2007). The utilisation of immobilised microbial cells in various
biotechnological fermentation processes was found to be advantageous over
the use of free cells (Cassidy et al., 1996).
Various techniques have been investigated for the purpose of microbial cell
immobilisation. These include adsorption or attachment of cells to an inert
substrate, self-aggregation by flocculation or using cross-linking agents and
entrapment or encapsulation using polymers (Jen et al., 1996; Kourkoutas et
al., 2004). Amongst these, immobilisation by adsorption and
entrapment/encapsulation are the most commonly used techniques.
Adsorption of microbial cells utilises the natural ability of cells to adhere onto
solid supports to form biofilms which can exist as a single layer or multilayers
of cells (Kourkoutas et al., 2004; Rezaee et al., 2008). For microbial cells that
do not adhere naturally, methods involving chemical cross-linking by
16
glutaraldehyde, silanisation onto silica support and metal oxide chelation can
be employed (Karel et al., 1985).
Cell immobilisation by encapsulation involves entrapping or coating microbial
cells with a continuous film of polymeric material to produce capsules
permeable to nutrients, gases and metabolites for cell growth and survival
(Ding and Shah, 2009; John et al., 2011). Based on the size of the capsules
produced, the encapsulation technique can be classified as macroencapsulation
and microencapsulation (John et al., 2011). In macroencapsulation, the
capsules produced have size ranging from a few millimetres to centimetres
(Gentile et al., 1995; John et al., 2011). On the other hand, the capsules
produced by microencapsulation have size range of 1-1000 microns (Byrd et
al., 2005; Heidebach et al., 2012).
D.1 Immobilisation of solventogenic clostridia
Solventogenic clostridia have also been immobilised using various systems for
the production of butanol. Many of these systems are based on immobilisation
of the cells either by adsorption or by entrapment/ encapsulation method
(Table 2).
17
Table 2. Summary of various cell immobilisation methods employed in ABE fermentation process
Strain Cell immobilisation method Carbon source Butanol yield
(g/L)
Productivity
(g/L/h)
References
Cl. acetobutylicum
ATCC 55025
Adsorption on fibrous matrix Glucose +
butyric acid
5.1
4.6
Huang et al., 2004
Cl. beijerinckii
LMD 27.6
Entrapment in calcium
alginate beads
Lactose 1.43 1.0 Schoutens et al.,
1985
Cl. beijerinckii
LMD 27.6
Entrapment in calcium
alginate beads
Glucose 2.4 0.8 Schoutens et al.,
1985
Cl. beijerinckii
BA 101
Adsorption on brick Glucose 8.1 16.2 Lienhardt et al.,
2002
Cl. saccharobutylicum
NCP 262
Adsorption on bone char Lactose 4.1 4.1 Qureshi and
Maddox, 1987
Cl. acetobutylicum
ATCC 824
Entrapment in calcium
alginate beads
Glucose 3.81 0.16 Häggström and
Molin, 1980
18
D.2 Limitations of conventional cell immobilisation methods used in ABE
fermentation
Although studies have shown that immobilised cells perform better than free
cells, the butanol yields are still on the lower side. These low yields can be
attributed to the inherent limitations of the respective cell immobilisation
method. For instance, extensive cell leakage from the support and insufficient
protection of the cells from harsh environmental conditions are often observed
with the adsorption method (Núñez and Lema, 1987). In addition, damage to
the cells may occur when they are exposed to the chemical cross-linking
agents used.
Cell immobilisation by entrapment/encapsulation method can overcome some
of the shortcomings of adsorption method. However, it has a significant
drawback of mass transfer limitation that needs to be addressed. Most of the
studies done using solventogenic clostridia are based on macroencapsulation
technique which forms beads larger than 1 mm. (Badr et al., 2001; Häggström
and Molin, 1980; Tripathi et al., 2010). Various problems due to mass transfer
limitations have been associated due to the large size of the beads. For
instance, low cell viability has been reported at the centre of the larger beads
after a relatively short operation time due to depletion in the nutrient diffusion
at a depth of more than 300-500 microns as well as accumulation of toxic
metabolites in the centre (McLoughlin, 1994). In addition, the hypoxic
conditions in the centre of larger beads caused cell death (Christenson et al.,
1993).
19
Many studies have reported that the optimum size of beads for efficient mass
transfer is in the size range of 0.1 - 1 mm (Christenson et al., 1993; Guiseley,
1989; Ogbonna et al., 1991). Smaller size beads permit high cell
concentrations within the beads by allowing efficient diffusion of nutrients,
oxygen as well as metabolites. In addition, they have been found to be more
mechanically robust than macrocapsules (Uludag et al., 2000). In spite of the
advantages of microencapsulation over macroencapsulation, there has been
little work done to explore the use of microencapsulation technique to
immobilise Cl. acetobutylicum cells.
E. Microencapsulation as a cell immobilisation technique
Microencapsulation has been applied to microbial cell immobilisation in order
to overcome the drawbacks encountered with other cell immobilisation
techniques such as cell leakage and contamination, in adsorption technique
and low mechanical stability and mass transfer limitations, in
macroencapsulation technique (Park and Chang, 2000).
The confinement of microbial cells in microspheres offers protection against
both mechanical as well as environmental stresses while maintaining growth
and metabolic activities for extended periods of time (Uludag et al., 2000).
Owing to their relatively small size, the microspheres have a larger specific
surface area for diffusion of nutrients into the microspheres and diffusion of
metabolites out of the microspheres. In the fermentation industry, besides the
above advantages, microencapsulation allows easy separation of cells and
20
minimizes cell wash out (Tan et al., 2011; Ylitervo et al., 2011). It is possible
to reuse the encapsulated microbial cells for continuous operation for
prolonged period of time due to constant cell regeneration within the
microspheres (Tan et al., 2011). Improved protection of cells from substrate
and end-product inhibition has been reported in numerous studies apart from
the decrease in undesirable process effects (Park and Chang, 2000).
Depending on the type of application, microbial cells can be encapsulated for
the purpose of isolation, protection and/or controlled release (Albertini et al.,
2010; Sultana et al., 2000). Moreover, the immobilised cells can be packed
into a column and the fermentation medium can be introduced from the top or
the packed immobilised cells can be placed in the fermenting medium. This
set-up can be used for both batch and continuous fermentation.
E.1 Techniques used for microencapsulation of microbial cells
Various techniques for microencapsulation of microbial cells have been
investigated over the past few years. Some of the techniques used include
extrusion, coacervation, spray drying and emulsification (de Vos et al., 2009).
The choice of techniques employed is dependent on the encapsulating
material, the type of microorganism and the desired microsphere properties.
The selected method should be able to produce microspheres with desired
physical/chemical attributes while causing minimal damage to cell integrity
and viability. It should also be easy to scale-up with acceptable processing
costs.
21
E.1.1 Extrusion
Extrusion is a commonly employed technique for the microencapsulation of
microbial cells (Ozer et al., 2008). In this method, a polymeric solution is
mixed with the microbial cells and extruded through a nozzle or orifice, to
form droplets which harden by contact with a cross-linking agent, cooling or
combination of both.
The major advantages of extrusion method are its simplicity, low cost, and
mild conditions that enable high cell viability. However, it has some
drawbacks, such as difficulty to form microspheres of size less than 500
microns due to the viscosity of the polymer and limited availability of suitable
nozzle size (Reis et al., 2006). Moreover, rapid cross-linking and hardening at
the surfaces of the microspheres delay the movement of cross-linking ions into
the inner core, resulting in less stable microspheres (Liu et al., 2002). In
addition, though, microspheres are conveniently produced at laboratory-scale;
the scaling-up of the process is difficult and involves high processing costs
(Burgain et al., 2011).
E.1.2 Coacervation
Microencapsulation using coacervation involves separation of one or more
polymers (coacervate) from the initial polymer solution, induced by varying
the pH, temperature or composition of the solution. The coacervate surrounds
the core material (e.g. cells dispersed in the polymer solution) resulting in
formation of microspheres (Gouin, 2004; Nihant et al., 1995; Oliveira et al.,
22
2007). If required, cross-linking agents can be used for strengthening the
microspheres. The coacervation method offers advantages that include high
encapsulation efficiency and ease of controlled liberation of encapsulated cells
by mechanical stress and temperature or pH changes (Oliveira et al., 2007).
However, high costs and the need to control various critical processing
conditions limit its usefulness (John et al., 2011).
E.1.3 Spray drying
Spray drying technique involves atomization of polymer dispersion into a
drying gas, followed by rapid evaporation of the solvent (Zhao et al., 2008).
The microspheres formed are collected as dry powder. Compared to other
techniques, spray drying offers the attractive advantage of producing
microspheres in a relatively simple continuous operation (Gouin, 2004).
However, when applied on a large scale, the high installation and operational
costs and considerable area occupied by the equipment present as major
limitations of the process (John et al., 2011). Besides, the range of polymers
that can be used for encapsulation is rather limited (Gouin, 2004). In addition,
the use of high air inlet temperature can lead to an excessive evaporation and
result in cracks in the polymeric membrane as well as loss of cell viability
(Brun-Graeppi et al., 2011).
E.1.4 Emulsification
Emulsification involves the dispersion of the cell/polymer suspension
(dispersed phase) in an oil or organic phase (continuous phase) with the aid of
23
surfactant(s) (Ding and Shah, 2009; Ozer et al., 2008). The mixture is
homogenised using a mechanical stirrer to form a water-in-oil emulsion.
Microspheres form by congelation of the dispersed phase through cooling or
the addition of a cross-linking agent into the emulsion. The microspheres are
harvested by filtration or centrifugation. Microencapsulation by emulsification
requires strict control of a number of parameters, such as concentration of
polymer, type and amount of surfactants, stirring speed as well as type and
concentration of cross-linking agent to obtain microspheres with desired
characteristics and encapsulation efficiency (Heng et al., 2003; Shah and
Ravula, 2000; Wan et al., 1994; Yang et al., 2000).
Emulsification can be utilised to produce microspheres of size below 300
microns, which is difficult to achieve with extrusion method due to clogging
of the orifice (Burgain et al., 2011). The shear forces used in emulsification
method disperse the cells heterogeneously within the microsphere (Rabanel et
al., 2009). The main drawback of the emulsification technique is the potential
toxicity of organic solvents to the cells. Although oils are less toxic than
organic solvents, removal of oil from the microspheres may be more difficult
compared to organic solvents. Moreover, its use in emulsification also
increases the overall cost of process (Krasaekoopt et al., 2004).
E.2 Polymers used for microencapsulation
Selection of an appropriate encapsulating polymer for microencapsulation of
cells is a crucial step. The microspheres produced should be mechanically
robust so as to preserve the viability and long-term culture of the microbial
24
cells (Tan et al., 2011). Both synthetic and natural water-soluble polymers
have been used for microencapsulation of microbial cells (Groboillot et al.,
1994; John et al., 2011). Though synthetic polymers offer higher mechanical
strength and better chemical stability, natural polymers are preferred over their
synthetic counterparts as the formulation of microspheres using natural
polymers usually requires milder process and is less harmful to cell viability
(Rosevear, 1984). In addition, microspheres produced using water-soluble
polymers are more permeable to low molecular weight nutrients and
metabolites, thus providing optimal conditions for the functioning of
immobilised microbial cells.
E.2.1 Alginate
Alginate is a frequently used polymer for the microencapsulation of cells
mainly because of its mild gelling conditions, simple gelling process with
divalent cations such as calcium cations and excellent biocompatibility and
biodegradability properties (Champagne et al., 2000; Green et al., 1996). It is
obtained from the cell walls of brown seaweed composed of D-mannuronic
and L-guluronic acids joined by glycosidic linkages. Factors affecting the gel
strength and stability, and consequently the activity of the encapsulated cells
include the type of alginate, alginate concentration, cross-linking agent
concentration and cell/alginate ratio (Albarghouthi et al., 2000; Vandenberg et
al., 2001).
25
E.2.1.1 Limitations of alginate as an encapsulating polymer
Microspheres or beads prepared using alginate have found to be susceptible to
chelating agents, substances present in microbial growth media as well as
metabolites produced during fermentation (Tan et al., 2011; Thu et al., 1996).
The presence of these substances can lead to disruption of the gel matrix and
subsequent release of the encapsulated cells into the fermentation medium.
This hinders the long term operation of the immobilised cells for continuous or
repeated batch fermentation. Alginate matrices have also been shown to be
susceptible to digestion by enzymes produced by many types of
microorganisms (Thu et al., 1996). Another drawback of alginate is its thermal
instability. It has been reported that alginates in solution degrade significantly
at temperatures around 60 to 120 °C and undergo extensive decomposition
when exposed to temperature above 250 °C (Holme et al., 2003). Thus
sterilisation of alginate solution by autoclaving is not ideal. Other sterilisation
methods employing ethylene oxide and gamma-irradiation also result in chain
depolymerisation, loss of viscosity and reduced gel strength (Thu et al., 1996).
Sterilisation of alginate solution by filtration will not cause polymer
degradation. However, this method of sterilisation is less preferred as it
requires aseptic technique and may not be feasible for high viscosity alginates.
E.2.2 Gellan gum
Gellan gum is another type of natural biocompatible polysaccharide produced
by the bacterium, Sphingomonas elodea. It is of commercial importance
because of its diverse applications in the food industry and biotechnology
26
(Shimazaki and Ogino, 1996). Gellan gum is water-soluble at temperature
above 90 °C and forms a transparent gel upon cooling to approximately 30 °C,
at sufficiently high polymer concentrations (Milas et al., 1990). The gelation
process is also affected by the presence of cations (Bajaj et al., 2007). There
are two main types of gellan gum, the acetylated and the deacetylated gellan
gum. Acetylated gellan gum is obtained by partial esterification of the native
gellan gum at the 3-linked-D-glucosyl residue with partial L-glyceric ester
substitution at C-2 of the 3-linked glucose and/or substitution of an acetyl
group at C-6 of the same residue (Figure 4a). The presence of acetyl groups in
gellan gum causes a massive change in the thermal stability of the double
helix structure of the molecule and affects its ability to form cation-mediated
aggregates (Morris et al., 1996). Deacetylated gellan gum is produced by
removing the aforementioned substituents with alkali (Figure 4b). It produces
aqueous solutions of low viscosity at high temperature and forms strong gels
upon cooling or in the presence of cations (Bajaj et al., 2007).
Rheological properties of gellan gum solution are strongly influenced by the
presence of cations because the latter promote association of helices into
cation-mediated aggregates which stabilise the gel. Divalent cations are able to
form more heat resistant gels than monovalent cations because the electric
charges of divalent cations are larger and shield the electrostatic repulsion of
the carboxyl groups in the molecules more effectively (Miyoshi and Nishinari,
1998). In addition, the junction zones formed by divalent cations have higher
thermal stability compared to monovalent cations (Moslemy et al., 2002).
27
Amongst the divalent cations, calcium ions are preferred as they form very
stable gels that are only reversible to liquid state when subjected to heat
treatment above 100 °C (Miyoshi and Nishinari, 1998). Only a small amount
of calcium ions is sufficient to induce gellan gum gelation to form a
homogeneous macroscopic network.
Unlike alginate, gellan gum is able to withstand the high temperature of the
autoclaving process without significant loss of gel strength (Yuguchi et al.,
2002). Gellan gum is also highly resistant to enzymatic breakdown (Chilvers
and Morris, 1987). As gellan gum is resistant to degradation by most
microorganisms, it is advantageous for encapsulation of microbial cells. In
addition, the acid-resistant gellan gum is useful for the encapsulation of
microbial cells for application in processes that involve acidic conditions
(Yuguchi et al., 2002). In spite of its many advantages, immobilisation of
solventogenic clostridia using gellan gum has not been investigated.
28
Figure 4. Chemical structures of (a) acetylated gellan gum and (b)
deacetylated gellan gum
F. Alternative fermentation substrates
In order to make the process of biobutanol production economically viable,
utilisation of cost effective fermentation substrates is required. Lignocellulosic
materials have been used as biomass feedstocks for biofuel production. These
materials are available as forestry wastes, municipal solid wastes, agricultural
and food wastes (Nigam and Singh, 2011). Utilisation of these sustainable
substrates is both economically viable and environmentally friendly. The
ability of saccharolytic clostridia to utilise many different carbohydrates is
(a)
(b)
29
advantageous for the utilisation of these cheap and renewable lignocellulosic
feedstocks (Blaschek, 1999).
Considerable research efforts have been put into the optimisation of the ABE
fermentation using various lignocellulosic substrates, such as corn, wheat
straw, barley straw, and switchgrass (Cheng et al., 2012; Ezeji et al., 2006;
Pfromm et al., 2010; Qureshi et al., 2007; Qureshi et al., 2010). Apart from
these, there exist numerous other lignocellulosic materials in nature which
have yet to be explored as biomass feedstock for biobutanol production. The
municipal waste that can be utilised as a potential lignocellulosic substrate for
biobutanol production is the leaf litter from Samanea saman tree, commonly
known as the rain tree.
F.1 Samanea saman tree (rain tree)
Samanea saman tree is a lofty tree, belonging to the leguminosae family, with
a massive umbrella-shaped and wide-spreading crown. The rain tree is
commonly planted in tropical regions for shade. The components of this tree
have been used for various purposes apart from providing shade. For example,
the fruits, wood and pods of the tree have been used as animal fodder, as well
as substrate for biofuels (Datt et al., 2008; Geeta et al., 1990; Jashimuddin et
al., 2006). The barks, leaves and fruits are known to contain compounds that
are active against a variety of ailments, such as colds, headache, enteritis,
diarrhoea and diabetes (Ferdous et al., 2010; Marles and Farnsworth,1995).
However, the S. saman leaves have not been investigated as a substrate for
30
biobutanol production and their relative degradability compared to other
biomass substrates is not known.
The leaf litter of S. saman tree is readily available throughout the year. The
removal of this waste leaf litter is either done manually or mechanically. The
collected leaves are then either used as manure, animal feed or destroyed in
incinerators by the local authorities. The disposal of the leaf litter is laborious
and costly, and the incineration process, if carried out in the open, releases
harmful gases such as carbon dioxide into the environment (Demirbas, 2005).
Finding an economical alternative use of the abundant leaf litter is thus much
to be desired.
It has been reported that at least 20 - 50 % of the S. saman leaf litter is
composed of lignocellulosic substances (Chanda et al., 1993; Datt C et al.,
2008). It also contains an appreciable amount of different sugars such as
xylose, arabinose and glucose (Chanda et al., 1993). Owing to the presence of
substantial amounts of fermentable sugars, the S. saman leaf litter offers the
potential for use as a substrate in biobutanol production. However, the
successful utilisation of any lignocellulosic substrate for biobutanol production
requires selection and optimisation of pretreatment conditions specific to the
substrate so as to release the fermentable sugars from the tightly woven
structure of its lignocellulosic matrix (Chandra et al., 2007). Subsequently, the
pretreated substrate needs to be detoxified to remove toxic substances
produced during the pretreatment process which may hinder the fermentation
process.
31
Figure 5. Different components of the S. saman tree: (a) Leaves, (b) pods,
(c) leaf litter and (d) flowers
F.2 Structure of lignocellulosic substrate
Lignocellulosic substrate is composed of three major components, namely,
cellulose, hemicellulose and lignin (Chen et al., 2004) (Figure 6). It also
contains variable amounts of other components, such as pectin, proteins and
minerals.
(d) (c)
(b) (a)
32
Figure 6. Structure of lignocellulose
Cellulose is made up of tightly packed polymeric chains of glucose (Figure
7a). It has a high crystallinity index that is responsible for its water insolubility
and resistance to depolymerisation (Sunkyu et al., 2010). Hemicellulose is a
branched polymer of glucose or xylose, substituted with arabinose, galactose,
fructose, mannose or glucuronic acid (Figure 7b). The hemicellulose binds to
cellulose microfibrils via hydrogen bonds to produce a network that provides
the structural backbone of the plant cell wall. Lignin is made up of phenyl
propanoid units and its main function is to impart support and strength to the
plant (Figure 7c). Reducing or fermentable sugars can be obtained from
cellulose and hemicellulose components of the matrix, also known as the
holocellulosic fraction, via enzymatic hydrolysis.
33
Figure 7. Chemical structures of (a) cellulose (b) hemicellulose and (c)
lignin
(a)
(b)
(c)
34
F.3 Pretreatment of lignocellulosic substrate
Factors such as the crystallinity of cellulose, limited accessible surface area
and protection of cellulose by lignin and hemicellulose impede enzymatic
hydrolysis of the holocellulose fraction of lignocellulose (Chang and
Holtzapple., 2000). Pretreatment is thus needed to modify the structural and
compositional barriers present in the lignocellulosic biomass and thereby
improve the efficiency of enzymatic hydrolysis and fermentable sugar yields
from holocellulosic fraction as shown in Figure 8 (Mosier et al., 2005).
Various physical and chemical pretreatment methods are usually employed,
either alone or in combination. It is important to judiciously select
pretreatment method/s for a given lignocellulosic substrate which is efficient,
can be scaled-up easily and forms little harmful by-products while preserving
the chemical integrity of the sugars (Chang and Holtzapple, 2000; Mosier et
al., 2005). In addition, it is crucial that the selected pretreatment method is
cost-efficient as it is one of the most costly steps of the conversion process
from biomass to fermentable sugars.
35
Figure 8. Pretreatment of lignocellulose
F.4 Types of pretreatment
Milling or size reduction, a mechanical pretreatment method, is often the first
step of pretreatment. It reduces particle size as well as crystallinity of the
lignocellulosic substrate (Schell and Harwood, 1994). Enzymatic hydrolysis is
principally initiated on the amorphous portions of cellulose. Thus, it is widely
accepted that by reducing the particle size and crystallinity of the substrate, the
fraction exposed to enzymatic hydrolysis is increased, without altering the
composition of the substrate (Cadoche and López, 1989). In addition, milling
also makes material handling easier in subsequent processing steps.
Apart from milling, several physical, chemical and biological pretreatment
methods and their combinations have been developed for making the
lignocellulosic substrates suitable for enzymatic hydrolysis and subsequent
fermentation. Some of these methods include ammonia fibre explosion, steam
36
explosion and acid and alkaline hydrolysis (El-Zawawy et al., 2011; Fox et al.,
1989; Holtzapple et al., 1994; Ruiz E et al., 2008). For economic reasons,
pretreatments such as acid hydrolysis and steam explosion, which solubilize
the hemicellulose components and increase cellulose accessibility, are
commonly preferred (Belkacemi et al., 1991; Zheng et al., 2009a).
Acid hydrolysis has been carried out for a variety of lignocellulosic substrates
and has been shown to be a promising method (Xu and Hanna, 2010; Zhang et
al., 2013; Zheng et al., 2009a). Both concentrated and dilute acids have been
employed for hydrolysing the highly complex lignocellulosic substrate (Sun
and Cheng, 2002). It enhances enzymatic hydrolysis by hemicellulose
removal, disruption of the lignin structure and deconstruction of cellulose,
enabling sugar recovery (Lenihan et al., 2010). Dilute acid coupled with heat
treatment (acid concentration of 0.5 - 5 %, w/w) at high temperature (120–160
°C) can hydrolyse hemicellulose to simple sugars (xylose, arabinose, and other
sugars) and acids (acetic, glucuronic), which are water-soluble (Lenihan et al.,
2010). On the other hand, concentrated acid pretreatment is operated at lower
temperatures such as 40 °C, and the sugar yield obtained is usually higher
compared to that of dilute acid coupled with heat treatment (Taherzadeh and
Karimi, 2007). Although concentrated acids are more effective than dilute
acids but they are also more toxic, corrosive and hazardous, requiring
corrosion-resistant reactor (Banerjee et al., 2010). Moreover, since a large
amount of acid is used, an acid recovery step is usually required which further
escalates the cost of the procedure. On the other hand, the amount of acid used
37
in dilute acid coupled with heat treatment is much less and thus acid recovery
is not necessary (Esteghlalian et al., 1997). Hence, dilute acid coupled with
heat treatment is a more common and cost-effective pretreatment method for
various lignocellulosic substrates.
F.5 Enzymatic hydrolysis of lignocellulosic substrate
Following pretreatment, the lignocellulose substrates are subjected to
hydrolysis by cellulolytic enzymes to release fermentable sugars for
biobutanol production. The cellulolytic enzymes, also known as cellulases, are
suite of several enzymes such as cellobiohydrolases (CBH I and II),
endoglucanases (EG I and III), and β-glucosidase, which together act
synergistically to degrade cellulose to glucose (Wilson, 2009). They are
produced by various fungal species (e.g. Trichoderma reesei and Aspergillus
sp.) and bacterial species (e.g. Cl. thermocellum and Cellulomonas flavigena)
(Lo et al., 2011; Suto and Tomita, 2001). These enzymes are highly specific
natural catalysts.
Enzymes constitute a significant cost in the bioconversion of a lignocellulosic
substrate and therefore the optimisation of the total amount of the enzyme
used is important. The amount of enzyme needed is dependent on the type of
substrate. For example, substrates with a high lignin composition may require
higher enzyme loadings due to non-productive adsorption of enzymes to the
lignin portion (Palonen et al., 2004). Thus, the optimal enzyme loadings have
to be identified for maximum efficiency of the enzymatic hydrolysis of the
lignocellulosic substrate.
38
F.5.1 Accellerase® 1500
Accellerase® 1500 is a commercial cellulase preparation composed of
combination of various enzymes such as exoglucanase, endoglucanase, hemi-
cellulase and β-glucosidase (Grande and Domínguez de María, 2012). It is
produced with a genetically modified strain of Trichoderma reesei. It is known
to efficiently and synergistically hydrolyse lignocellulosic biomass into
fermentable sugars. It has been well-documented that cellulases are inhibited
by cellobiose, a disaccharide produced by partial hydrolysis of cellulose, while
β-glucosidase is inhibited by glucose (Wilson, 2009). For this reason,
Accellerase® 1500 contains higher levels of β-glucosidase activity than other
commercial cellulases available, to ensure almost complete conversion of
cellobiose to glucose. Accellerase® 1500 appears as a brown liquid and has a
pH of 4.6 - 5.0. The optimum temperature and pH for best enzyme activity
have been reported to be 50 - 65 °C and 4.0 - 5.0 respectively (Balsan et al.,
2012).
F.6 Strategies for detoxification of acid hydrolysate
The acid hydrolysis of lignocellulosic substrates produces various degradation
products derived from sugars and lignin such as furfural, hydroxyfurfural,
acetic acid, levulinic acid, vanillin and phenolic compounds, apart from the
fermentable sugars (Mussatto and Roberto, 2004). These degradation products
can adversely affect the subsequent steps of enzymatic hydrolysis. In addition,
some of the degradation compounds are toxic to the bacterial cells and may
39
inhibit their growth and metabolism (Taherzadeh et al., 2000). Thus, it is
necessary to detoxify or remove these compounds from the acid hydrolysates.
A number of biological, physical and chemical detoxification methods have
been proposed to transform the toxic compounds into inactive forms or to
reduce their concentration. The selection of any detoxification method is
dependent on the type of lignocellulosic substrate, as well as the species of
microorganism used for fermentation (Mussatto and Roberto, 2004; Palmqvist
and Hahn-Hägerdal, 2000). Among the biological detoxification methods,
treatment with enzymes such as peroxidase and laccase for the removal of
phenolic monomers and phenolic acids has been investigated (Jönsson et al.,
1998). Acetic acid, furfural and benzoic acid derivatives were removed from
the hydrolysate by the treatment with enzymes produced by Trichoderma
reesei (Palmqvist and Hahn-Hägerdal, 2000). The most commonly used
physical detoxification method involved vacuum evaporation of the volatile
compounds in the acid hydrolysate such as acetic acid, furfural and vanillin
(Palmqvist and Hahn-Hägerdal, 2000; Parajó et al., 1998). Chemical
detoxification of lignocellulosic hydrolysates involves pH adjustment of the
hydrolysate using alkali and acid (Roberto et al., 1991). Another chemical
detoxification method, known as overliming, is also commonly employed
(Martinez et al., 2001). In this method, calcium hydroxide is used to increase
the pH of the acid hydrolysate to 9 - 10, followed by readjustment to 5.5 with
sulphuric acid. Overliming has been reported to result in better fermentability
40
than pH adjustment using sodium hydroxide, due to the precipitation of toxic
compounds (Van Zyl et al., 1991).
Detoxification steps increase the overall costs of the production process (Li et
al., 2011; Liu, 2011). Hence, it is crucial to carefully evaluate the need of
removing the inhibitors. The detoxification method should be inexpensive,
easy to integrate into the process, and able to remove inhibitors selectively
without any significant loss of fermentable sugars.
41
HYPOTHESES AND OBJECTIVES
42
II. HYPOTHESES AND OBJECTIVES
A. Hypotheses
Immobilisation of solventogenic clostridia by adsorption or entrapment has
been employed as a technique to overcome the frequently encountered
problems in biobutanol production namely, solvent toxicity, low viable cell
densities and low yields and productivity. However, due to problems such as
limited cell protection, mass transfer limitation and low viable cell densities,
butanol yield and productivity obtained with these techniques are low.
Microencapsulation by emulsification technique has been shown to be a
promising cell immobilisation strategy used in other fields as it enables the
formation of microspheres in the size range of 0.1 - 1mm. These small-sized
microspheres have high surface area to volume ratio, thereby allowing
efficient mass transfer of nutrients and metabolites.
It was hypothesised that microencapsulation of Cl. acetobutylicum ATCC 824
cells in gellan gum microspheres by emulsification, would provide the cells
with a protective barrier against solvent toxicity during fermentation and thus
maintain the viability of the cells. The large surface area to volume ratio
would enable efficient mass transfer of nutrients and metabolites and thus
improve butanol yield. Furthermore, the microspheres could be easily
recovered and reused for successive batch fermentation cycles. It was also
postulated that the spore form of the bacterium would be more amenable for
encapsulation by emulsification method owing to its hardy characteristics.
43
Another major deterrent in the large scale production of biobutanol is the high
cost of the fermentation substrates. Various alternative substrates for
biobutanol production have been investigated with variable degree of success.
Samanea saman or rain tree, commonly planted for shade, generates huge
amount of leaf litter daily. The utilisation of the leaf litter as a potential
lignocellulosic substrate is yet to be explored.
It was hypothesised that selection and optimisation of suitable pretreatment
conditions and subsequent enzymatic hydrolysis will enable recovery of
available sugars in S. saman leaf litter. Selection of suitable detoxification
methods will be required to make the pretreated leaf litter amenable for
bacterial fermentation. Lastly, it was postulated that encapsulation of Cl.
acetobutylicum ATCC 824 cells will enable better fermentability of the S.
saman leaf litter owing to the protective effect of the polymer matrix against
inhibiting compounds present in the pretreated leaf litter as well as solvents
formed during fermentation.
44
B. Objectives
The objectives of this study were:
Part One
1. To determine the optimal growth conditions for Cl. acetobutylicum ATCC
824 with respect to growth medium, incubation temperature and anaerobic
growth set-up and heat shock treatment for the revival of spores.
2. To optimise the process of microencapsulation by emulsification method to
produce microspheres in the desired size range and minimum span and
aggregation index.
3. To investigate the effect of emulsification method on the viability of Cl.
acetobutylicum ATCC 824 vegetative cells and spores.
4. To optimise the microencapsulation of Cl. acetobutylicum ATCC 824 cells
for the purpose of biobutanol production.
Part Two
1. To compare the fermentation efficiency between free and encapsulated Cl.
acetobutylicum ATCC 824 cells for biobutanol production.
2. To evaluate the stability of gellan gum microspheres during the
fermentation process.
3. To compare the reusability between free and encapsulated Cl.
acetobutylicum ATCC 824 cells in the biobutanol production in repeated
batch fermentation.
45
Part Three
1. To investigate different pretreatment methods for S. saman leaf litter.
2. To investigate different detoxification methods for pretreated S. saman leaf
litter.
3. To utilise detoxified pretreated S. saman leaf litter for fermentation by free
and encapsulated cells of Cl. acetobutylicum ATCC 824.
46
EXPERIMENTAL
47
III. Experimental
A. Materials
A.1 Model microorganism
Clostridium acetobutylicum ATCC 824, in freeze-dried form, was purchased
from American Type Culture Collection (ATCC, Virginia, USA). The culture
was maintained as a spore suspension in sterile distilled water at 4 °C.
A.2 Growth media
Reinforced Clostridial Medium (RCM broth, Acumedia, Michigan, USA) was
used as the growth medium for Cl. acetobutylicum ATCC 824. One litre of
this medium consisted of 3.0 g yeast extract, 10.0 g beef extract, 10.0 g
peptone, 5.0 g glucose, 5.0 g sodium chloride, 1.0 g soluble starch, 0.5 g
cysteine hydrochloride, 3.0 g sodium acetate, and 0.5 g agar, adjusted to pH
of 6.8 ± 0.2. Thioglycollate medium (TGM broth Oxoid, Hampshire,
England) was also investigated as a growth medium for the cultivation of Cl.
acetobutylicum ATCC 824. One litre of this medium consisted of 5.0 g yeast
extract, 15.0 g tryptone, 5.5 g glucose, 0.5 g sodium thioglycollate, 2.5 g
sodium chloride, 0.5 g L-cysteine, 0.001 g resazurin and 0.75 g agar, adjusted
to pH of 7.1 ± 0.2. The corresponding solid agar media consisted of the broth
with 1.5 %, w/v of purified agar (Oxoid, Hampshire, England).
48
A.3 Fermentation medium
RCM broth supplemented with glucose monohydrate (Merck, Darmstadt,
Germany) was used as the fermentation medium.
A.4 Encapsulating polymer and chemicals
Gellan gum (CP Kelco, San Diego, USA) was used as the encapsulating
polymer. Isooctane (analytical grade, Merck, Darmstadt, Germany) was used
as the continuous phase of the test emulsion, with Tween 80 (Merck,
Darmstadt, Germany) and Span 80 (Sigma Aldrich, Missouri, USA) as the
emulgents. Calcium chloride dihydrate (Merck, Darmstadt, Germany) was
used as the cross-linking agent.
A.5 Chemicals for assay of butanol by gas chromatography-mass
spectrometry
1-Butanol (spectroscopic grade, Merck, Darmstadt, Germany) was used for the
preparation of standard solutions in the assay of butanol. 1-Propanol
(spectroscopic grade, Merck, Darmstadt, Germany) was used as the internal
standard. 2-ethyl 1-hexanol (synthesis grade, Merck, Darmstadt, Germany)
was used to extract butanol from the fermentation medium. Purified helium
gas (99.99%, 200 bar, Soxal, Singapore) was used as the carrier gas for GC
MS analysis.
49
A.6 Lignocellulosic substrate
Leaf litter collected from the S. saman tree was used as the biomass feedstock.
The leaves were first separated from the stalks and twigs and washed with
water to remove any soil debris. The washed leaves were dried overnight in
the oven at 60 °C.
A.7 Cellulolytic enzyme
Commercial cellulase complex, Accellerase® 1500 (Genencor, California,
USA) was used as the cellulolytic enzyme in the experiments.
A.8 Chemicals used in assay of reducing sugars
Assay of the reducing sugars was carried out using the dinitro salicylic acid
(DNS) method. Glucose (Merck, Darmstadt, Germany) was used as the
standard for preparation of glucose calibration plot. Other chemicals used
were dinitrosalicylic acid (Sigma Aldrich, Missouri, USA), sodium potassium
tartrate (Merck, Darmstadt, Germany), sodium metasulphite (Merck,
Darmstadt, Germany), phenol (Alfa Aesar, Lancashire, England), and sodium
hydroxide (Merck, Darmstadt, Germany).
A.9 Chemicals used for dilute acid coupled with heat treatment of S.
saman leaf litter
Sulphuric acid (98 %, BDH Chemicals Ltd, Poole, England) was used to
prepare different concentrations of dilute acid for the pretreatment of S. saman
50
leaf litter. Sodium hydroxide (Merck, Darmstadt, Germany) was used to adjust
the pH of the acid hydrolysate.
A.10 Chemicals used for measuring the filter paper units (FPU) activity of
Accellerase® 1500
Citric acid (Merck, Darmstadt, Germany), dinitrosalicylic acid (Sigma
Aldrich, Missouri, USA), sodium potassium tartrate (Merck, Darmstadt,
Germany), sodium metasulphite (Merck, Darmstadt, Germany), phenol (Alfa
Aesar, Lancashire, England) and sodium hydroxide (Merck, Darmstadt,
Germany) were used to determine the FPU activity of the commercial enzyme,
Accellerase® 1500. Whatman filter paper no. 1 (Whatman, Kent, England)
was used as the substrate in the FPU assay.
A.11 Chemicals used for detoxification of acid hydrolysate of S. saman
leaf litter
Sodium hydroxide (Merck, Darmstadt, Germany) and calcium hydroxide
(Merck, Darmstadt, Germany) were used as detoxifying agents for the acid
hydrolysates of the leaf litter.
51
B. METHODS
B.1 Preparation of growth media
A specific amount of powdered medium was weighed and dispersed in
distilled water. The volume was made up to a specific volume. Sterilisation of
the media was performed by autoclaving at 121 °C at 15 psi for 20 min.
B.2 Cultivation of Cl. acetobutylicum ATCC 824
B.2.1 Revival of Cl. acetobutylicum ATCC 824
The freeze-dried culture of Cl. acetobutylicum ATCC 824 was revived by
inoculation in RCM broth in a tightly capped bottle, which was then incubated
at 37 °C for 48 h. Loopfuls of the turbid broth were streaked on RCM agar
media, which were incubated under anaerobic conditions using gas pack
systems (Anaerogen™, Oxoid, Hampshire, England). After 48 h of incubation,
single colonies were observed on agar plates. Pure subcultures were then
prepared from these single colonies using the most appropriate media and
incubation conditions determined.
B.2.2 Determination of suitable media for the growth of Cl.
acetobutylicum ATCC 824
A single colony derived from the revived culture was each inoculated into
RCM and TGM broths as well as the agar media respectively. The inoculated
samples were then incubated under anaerobic conditions using gas pack
system at room temperature, 25 °C and 37 °C. They were examined for cell
52
growth after different time intervals of the incubation period. The suitable
medium and incubation temperature were those that could achieve maximum
cell growth in minimum time. Each experiment was carried out in triplicate.
B.2.3 Determination of suitable anaerobic set-up for the growth of Cl.
acetobutylicum ATCC 824
Different set-ups for the maintenance of anaerobic conditions were
investigated for the growth of Cl. acetobutylicum ATCC 824. Plate cultures of
Cl. acetobutylicum ATCC 824 were incubated under anaerobic conditions
using gas pack systems, Anaerogen™ (Oxoid, Hampshire, England) and
Campygen™ (Oxoid, Hampshire, England) respectively. Broth cultures were
incubated in loosely capped bottles under anaerobic conditions or in tightly
capped bottles under normal atmospheric conditions. The ratio of liquid to air
in broth culture was 3:1 and the cultures were kept static. Experiments were
conducted in triplicate. The extent of cell growth in broth cultures under the
respective incubation conditions was determined after 48 h of incubation using
spread plate method described in section B.2.4.1. In the case of plate cultures,
presence or absence of colonies was observed. The set-up which could give
the maximum cell growth after 48 h of incubation was selected as the suitable
one.
53
B.2.4 Determination of growth curve and morphology of Cl.
acetobutylicum ATCC 824
Clostridium acetobutylicum ATCC 824 was cultivated in 300 mL of suitable
cultivation broth under pre-determined optimal conditions. Samples were
taken at 6, 12, 18, 24, 30, 36, 48 and 54 h to determine the viable count by
turbidimetric measurements using a UV-VIS spectrophotometer (U-1900,
Hitachi, Japan) at 600 nm. The spectrophotometric readings were calibrated
against known viable counts determined by the spread plate method (described
in section B.2.4.1). From the results, a plot of log viable count against time
was constructed. The different phases of growth of Cl. acetobutylicum ATCC
824 were determined from the plot and the corresponding morphological
features of the bacterium were examined using an optical microscope
(Olympus Corporation, Tokyo, Japan). Experiments were conducted in
triplicate.
B.2.4.1 Determination of viable count of Cl. acetobutylicum ATCC 824 by
spread plate method
The viable count of sample was determined using the spread plate method
(Figure 9, Messer et al., 1999). Serial dilutions of the sample were prepared. A
0.1 mL volume of each dilution was then spread onto suitable agar medium,
which was then incubated at 37 °C under suitable anaerobic conditions. Total
number of colonies developed at the end of 48 h of incubation was determined
using a colony counter. It was assumed that each viable cell in the culture
54
formed an individual colony. The experiment was performed in triplicate. The
viable count of the sample was calculated as follows:
Viable count = Average number of colonies per dilution X dilution factor
(CFU/mL)
Figure 9. Schematic diagram of quantification of viable cells by spread
plate method
B.2.5 Preparation of spore stock culture of Cl. acetobutylicum ATCC 824
The bacterial culture was grown on RCM agar plates anaerobically at 37 °C
for 72 h. The plates were removed from anaerobic conditions after 72 h and
stored at room temperature for 9 days, thus exposing the culture to normal
atmospheric conditions. For the preparation of the spore stock culture, the
colonies from the plates were scraped using a sterile spreader and washed with
sterile water. The washings were collected and centrifuged at 10,000 rpm for
10 min. The pellet formed was washed thrice with sterile water and then
suspended in a specific volume of sterile water. The spore count was estimated
55
as described in section B.2.5.1. The resultant spore suspension was stored at 4
°C for future use.
B.2.5.1 Determination of viable spore count of Cl. acetobutylicum ATCC
824 spore suspension
Serial dilutions of the spore suspension were prepared. Heat shock treatment
was given to each of these dilutions in order to revive the spores (Gould,
1971). The viable spore count was then determined by the spread plate method
described in section B.2.4.1.
B.2.6 Optimisation of heat shock treatment (HST) conditions for the
revival of Cl. acetobutylicum ATCC 824 spores
A volume of 0.1 mL of spore suspension containing 107 spores/mL was
prepared as described in section B.2.5. The spore suspension was added to a
20 mL of RCM broth. The mixture was then subjected to different heat shock
treatment conditions by placing the tubes containing the suspension in water
baths maintained at desired temperature (Table 3).
Table 3 . HST optimisation of Cl. acetobutylicum ATCC 824
Temperature ( °C) Time (min)
70 3, 5, 10, 15
80 3, 5, 10, 15
90 3, 5, 10, 15
100 3, 5 , 10, 15
56
Each treated spore suspension was then incubated at 37 °C under anaerobic
conditions. Controls with no heat shock treatment were also prepared and
incubated under same conditions as test samples. Presence or absence of
turbidity in the broth samples was observed after 48 h of incubation. Viable
count of revived spores was estimated by spread plate method (section
B.2.4.1). Each experiment was performed in triplicate.
B.2.7 Preparation of standardised inoculum of vegetative cells of Cl.
acetobutylicum ATCC 824
A specific volume of spore suspension of Cl. acetobutylicum ATCC 824 was
given heat shock treatment using optimised conditions. A volume of 0.1 mL of
the revived spore suspension was then spread onto RCM agar, which was then
incubated anaerobically at 37 °C. After 48 h of incubation, numerous single
colonies were observed on the plates. Around 5-6 single colonies were used to
inoculate 20 mL RCM broth in a bottle. The bottle was capped tightly to
simulate an anaerobic environment, and incubated at 37 °C for 24 h. One mL
of the resultant broth culture was introduced into 19 mL of fresh RCM broth in
a bottle which was tightly capped and incubated at 37 °C for 24 h to produce
standardized 24-hour broth culture. Cell count of the standardized culture of
Cl. acetobutylicum ATCC 824 was estimated by optical density and spread
plate method.
57
B.3 Production of microspheres by emulsification method
A modified emulsification method was used in this study (Wan et al., 1992).
The schematic diagram of the procedure used is shown in Figure 10. Fifty
grams of gellan gum solution was used as the aqueous phase. It was prepared
by rehydrating gellan gum in the required quantity of distilled water at 80 °C
for 20 min under mild agitation using a magnetic stirrer (Lee Hung Scientific
Pvt Ltd, Singapore). Thereafter, the solution was sterilised by autoclaving at
121 °C at 15 psi for 20 min. It is necessary to point out that sterilisation was
required only for encapsulation of Cl. acetobutylicum ATCC 824 cells and not
for this study, where blank microspheres were produced. However, slight
change in viscosity of the gellan gum solution was previously observed after
autoclaving which could affect the properties of the microspheres. Hence the
gellan gum solution was sterilised in the production of both blank and cell-
loaded microspheres. The organic phase was composed of 75 g of isooctane
with span 80. The organic phase was added to the aqueous phase placed in a
water bath at 50 °C and the mixture stirred at 500 rpm using a mechanical
stirrer (Eurostar digital, Selangor, Malaysia) for 10 min. Tween 80 solution
was then added to the above dispersion and stirring continued for another 5
min. The temperature of the water bath was then reduced to 15 °C to induce
congelation of gellan gum to form microspheres. The dispersion was stirred
for 10 min, followed by the addition of 20 g of 25 %, w/w calcium chloride as
cross-linking agent to further stabilise the gellan gum matrix. Stirring was
continued for another 15 min and the dispersion kept in a shaker water bath at
58
room temperature for 1 h to allow the microspheres to cure. The microspheres
were harvested by filtration in vacuo using a Buchner funnel lined with filter
paper, (Whatman No. 54, GE Healthcare Life Sciences, New Jersey, USA),
and washed thrice with 20 mL sterile water. In the production of microspheres
consisting of cells or spores, the latter was added to the gellan gum solution
before emulsification. Also, the organic phase was sterilised by filtration.
B.3.1 Optimisation of production of gellan gum microspheres
The influence of parameters known to affect the formation and properties of
microspheres, namely concentration of polymer, HLB of the surfactant blend,
temperature used for emulsification and stirring speed on the size, span and
aggregation index of the microspheres was studied using a 24 full factorial
design (Forghani et al. 2013). Each factor was varied over three levels: the
high level (+1), the intermediate point (0) and the low level (−1). Other factors
such as concentration of cross-linking agent, volume of organic phase and
stirring time were set at fixed values. The total number of runs was 35. The
order of the tests was randomised to avoid any uncontrolled systematic error.
The coded and actual values of these variables along with their low and high
levels and the response variables are shown in Tables 4 and 5 respectively.
The coded mathematical model for 24 factorial designs can be given as
Y = X0 + X1A + X2B + X3C + X4D + X5AB + X6AC + X7AD + X8BC +
X9BD + X10CD (1)
59
where Y is the response, X0 is the constant, Xi represents the other regression
coefficients and A, B, C, D stand for concentration of gellan gum, HLB of
surfactant blend, temperature of emulsification and stirring speed,
respectively. Individual factors represent main effect while any combination of
two factors represents interaction effect. The goodness of fit of the model,
coefficient estimates and R2 values were checked to assess the suitability of
the model in describing the responses adequately. The significant and
insignificant terms were assessed based on their respective p-values from the
ANOVA table.
Table 4. Coded and uncoded values of the two independent factors in the
optimisation of microencapsulation process
Independent variable Unit Coded value Actual value
Gellan gum concentration (A) %, w/v -1 1.0
0 1.5
+1 2.0
HLB of surfactant blend (B) NA -1 8
0 9
+1 10
Emulsification temperature (C) °C -1 40
0 50
+1 60
Stirring speed (D) rpm -1 400
0 600
+1 800
60
Table 5 Response variables used in the 24
full factorial design
Response Name Units
Y1 Size microns
Y2 Span NA
Y3 Aggregation index percent
Three dimensional surface plots were constructed using Design-Expert
software to study significant main and interaction effects. The optimal
condition for preparation of microspheres was determined using the equations
derived from the model, taking into consideration the contributions of all the
factors and the surface plots. To validate the chosen experimental design and
model equations, optimum test condition was selected and production of
microspheres using the optimum condition and three other random runs were
carried out. The response values determined experimentally were
quantitatively compared with corresponding predicted values. Close match
between experimental and predicted values indicates good correlation between
the predicted model and experimental results.
61
Figure 10. Production of gellan gum microspheres using emulsification method
50 °C 50 °C 50 °C 15 °C 15 °C Curing for 1 hour Congealation Cross-linking Emulsification Gellan gum
solution
Organic phase: 75 g of
isooctane + span 80 20 g of calcium chloride
solution (25 %, w/w)
15 min 10 min 5 min 10 min 5 min
5 g of tween 80
solution
62
B.3.2 Characterisation of the microspheres
The size and size distribution of hydrated microspheres were determined by
laser diffractometry (LS 230, Coulter Corporation, California, USA).
Measurements were made in triplicate for each batch. Particle size distribution
was determined by the span value which was calculated as follows:
Span =
(2)
where D(V90), D(V10) and D(V50) are volume size diameters at 90, 10 and
50 % of the cumulative volume, respectively. A smaller span value indicates a
narrower size distribution.
The morphology and aggregation index of the hydrated microspheres was
determined by microscopic examination using a light microscope (BX61-TRF,
Olympus, Tokyo, Japan) connected to an image analyser (DXC-390P 3CCD,
Sony, Tokyo, Japan). A total of at least 300 microspheres mounted in water on
a microscope slide was evaluated using the Image Pro-express 6.3 Software
(Media Cybernetics, Maryland, USA).
The total number of discrete and aggregated microspheres was determined.
The degree of aggregation was calculated as follows:
Aggregation index = umber of aggregated microspheres
Total number of microspheres (discrete aggregated) X 100 (3)
63
B.4 Study of emulsification process on viability of Cl. acetobutylicum
ATCC 824 vegetative cells/spores
In order to select a suitable stage of growth of Cl. acetobutylicum ATCC 824
for microencapsulation, it was necessary to study the effect of the
emulsification process on the viability of the vegetative cells and spores. A 50
g suspension of Cl. acetobutylicum ATCC 824 cells containing approximately
107 vegetative cells/mL in exponential phase of growth was subjected to the
emulsification process (without gellan gum) as described in section B.3. The
viable counts of Cl. acetobutylicum ATCC 824 vegetative cells before and
after the emulsification process were determined by the spread plate method
(section B.2.4.1). The results were presented in terms of log reduction i.e. the
difference between the logarithmic values of viable cell count before and after
the emulsification process. Each experiment was carried out in triplicate. The
above procedure was repeated with 50 g of spore suspension containing
approximately 107 spores/mL. The spores were revived by heat shock
treatment using pre-determined conditions before viable count determination
by spread plate method.
B.5 Method development for the assay of butanol by gas
chromatography-mass spectrometry (GC-MS)
The analysis of butanol was done using GC-MS (GC-MS QP 2010, Shimadzu
Corp., Kyoto, Japan) equipped with an autosampler (AOC 20i). The GC-MS
was operated with an interface temperature of 220 °C, and an ionization
source temperature of 240 °C. The mass spectrometer was tuned before
64
analysis using PFTBA (perfluorotributylamine). The solvent delay, before the
MS filament was turned on, was set to 1 min. Chromatographic separation was
achieved using a polar wax column, BP-wax 20 (0.25 µm film thickness, 30m
× 0.25 mm I.D., SGE, Texas, USA). Helium with a minimum purity of 99.99
% was used as the carrier gas. Samples were injected in split mode. The mass
spectrometer was operated in the positive ion electron impact (EI) mode. The
mass spectra were obtained at an ionizing energy of 70 eV. Quantification was
carried out in the SCAN mode of the mass spectrometer. Optimisation of the
analytical parameters was carried out by varying the column oven temperature
program, gas flow-rate, gas flow-pressure and split-ratio. The selection of the
best conditions for analysis was based on the shape and resolution of the
analyte peak. A good peak can be defined as a narrow peak with good
separation from other components of the sample.
B.5.1 Extraction of butanol from fermentation medium by liquid-liquid
extraction
A 2 mL volume of fermentation sample was first centrifuged at 10,000 rpm for
10 min at 20 °C. A 1 mL volume of the supernatant was added to an equal
volume of 2-ethyl 1-hexanol. The mixture was vortexed (Fisons WhirliMixer,
Haverhill, England) for 5 min and centrifuged (RA-50J, Kubota 1720, Osaka,
Japan) at 6,000 rpm for 5 min at 20 °C. The upper organic phase was
separated and assayed for butanol by GC-MS using the predetermined
optimised conditions. The butanol assay was carried out in triplicate.
65
B.5.2 Preparation of butanol calibration curve
Fermentation media containing different concentrations of 1-butanol (0 - 15
g/L) were prepared. The sample aliquots were first passed through 0.45
microns membrane filter, followed by extraction using 2-ethyl 1-hexanol and
assay by GC-MS as described in sections B.5.1. Each experiment was repeated
3 times and the results averaged. The calibration plot of the ratio between the
peak areas against the concentration of butanol was constructed using GC-MS
Lab Solutions software.
B.5.3 Assay of butanol produced in the fermentation medium
An aliquot sample from the fermentation medium was centrifuged (RA-50J,
Kubota 1720, Osaka, Japan) at 6,000 rpm for 10 min at 20 °C and the
supernatant passed through 0.45 microns membrane filter. This was followed
by the extraction according to the procedure, described in section B.5.1.
Butanol yield was then calculated by extrapolating the peak area of the sample
in the butanol calibration curve (section B.5.2).
B.5.4 Calculation of extraction efficiency of 2-ethyl 1-hexanol
Fresh fermentation media, containing 5, 10 and 15 g/L of 1-butanol
respectively, were prepared. Butanol was then extracted and assayed
according to the procedures described in sections B.5.1 and B.5.3. The
concentration of butanol was determined using the calibration curve. The
amount of butanol recovered was compared to the corresponding butanol
66
added to the fermentation medium. The extraction efficiency was calculated as
follows:
(4)
B.6 Fermentation studies using Cl. acetobutylicum ATCC 824 cells
B.6.1 Preparation of fermentation media
RCM broth of appropriate concentration was sterilised by autoclaving at 121
°C and 15 psi for 20 min. The glucose (substrate) solution was sterilised
separately by filtration to prevent caramelization. It was then mixed with the
previously sterilised RCM broth in appropriate amounts to obtain the desired
glucose concentration in RCM broth
B.6.2 Optimisation of fermentation by free (non-encapsulated) vegetative
cells of Cl. acetobutylicum ATCC 824
A standardised inoculum of Cl. acetobutylicum ATCC 824 containing
approximately 107 cells/mL was used to inoculate the 100 mL of fermentation
medium contained in 200 mL Schott bottles in a shaker water bath maintained
at 37 °C. The effects of various factors on the fermentation efficiency were
investigated (Table 6). The bottles were loosely capped during fermentation to
allow the escape of fermentation gases. Samples were withdrawn aseptically at
regular intervals for the determination of cell density, pH, butanol yield and
productivity. Butanol productivity (g/L/h) was calculated as the butanol yield
67
in g/L divided by the fermentation time in h. All experiments were carried out
in triplicate. Suitable conditions were derived based on the combination of
factors that resulted in maximum butanol yield and productivity.
Table 6. Different combinations of parameters investigated in the
optmisation of fermentation by free vegetative cells of Cl. acetobutylicum
ATCC 824
Run Inocula size
(%, v/v)
Glucose concentration
(%, w/v)
Inocula age
(h)
Code
1 10 6 24 F-10-6-24
2 10 8 24 F-10-8-24
3 10 6 48 F-10-6-48
4 10 8 48 F-10-8-48
5 15 6 24 F-15-6-24
6 15 8 24 F-15-8-24
7 15 6 48 F-15-6-48
8 15 8 48 F-15-8-48
B.6.3 Optimisation of fermentation by encapsulated spores of Cl.
acetobutylicum ATCC 824
Standardised inocula of Cl. acetobutylicum ATCC 824 spores (107 viable
spores/mL) was encapsulated in gellan gum microspheres using the optimised
set of conditions obtained in section B.3.1. The encapsulated spores were then
introduced into the fermentation medium and revived using different heat
shock treatment that involved heating the medium at high temperature. This
also ensured anaerobiosis of the fermentation medium by expelling any
68
dissolved oxygen through heating the medium. The effects of the following
factors on the amount of butanol produced by encapsulated Cl. acetobutylicum
ATCC 824 spores were determined (Table 7).
Table 7. Different combinations of parameters investigated in the
optmisation of fermentation by encapsulated spores of Cl. acetobutylicum
ATCC 824
Run Inocula size
(%, v/v)
Glucose
concentration
(%, w/v)
HST Code
1 10 6 90 °C, 15 min S-10-6-9015
2 15 6 90 °C, 15 min S-15-6-9015
3 10 8 90 °C, 10 min S-10-8-9010
4 15 6 90 °C, 10 min S-15-6-9010
5 15 8 90 °C, 10 min S-15-8-9010
6 10 6 90 °C, 10 min S-10-6-9010
7 15 8 90 °C, 15 min S-15-8-9015
8 10 8 90 °C, 15 min S-10-8-9015
All experiments were carried out using 100 mL of fermentation medium in
200 mL Schott bottles in a shaker water bath maintained at 37 °C. The bottles
were loosely capped during fermentation to allow the escape of fermentation
gases. Experiments were performed in triplicate. Samples were withdrawn
aseptically at regular intervals for the determination of pH of the medium,
morphology of the microspheres, butanol yield and productivity. Suitable
conditions were derived based on the combination of factors that resulted in
maximum butanol yield and productivity.
69
B.6.4 Measurement of optical cell density
The cell density of the fermentation samples was determined by measuring the
optical density (OD) of the cell suspension using a UV-VIS spectrophotometer
(UV-1900, UV-VIS spectrophotometer, Hitachi, Japan) at 600 nm. The
calibration curve prepared in section B.2.4 was used to determine the cell
count from the measured optical density.
B.6.5 Measurement of fermentation medium pH
After measurement of the optical density, fermentation samples were
centrifuged (RA-50J, Kubota 1720, Japan) at 16, 000 rpm for 5 min at 20 °C.
The clear supernatant was then used for pH measurement using a pH test strip
(Whatman, Kent, England).
B.7 Determination of viable count of cells liberated from microspheres
into the fermentation medium
The viability of the Cl. acetobutylicum ATCC 824 cells liberated from the
microspheres was determined at the end of each fermentation cycle by the
spread plate method as described in section B.2.4.1. Each experiment was
conducted in triplicate.
B.7.1 Examination of physical stability of microspheres
Gellan gum microspheres were recovered from the fermentation medium at
the end of each fermentation cycle. The physical appearance of the
microspheres was observed using a light microscope (BX61-TRF, Olympus,
70
Tokyo, Japan) and software program (Image Pro-Express 6.3 Software, Media
Cybernetics, Maryland, USA).
B.8 Comparison of reusability between free (non-encapsulated) cells and
encapsulated cells of Cl. acetobutylicum ATCC 824
The free (non-encapsulated) Cl. acetobutylicum ATCC 824 cells were
recovered at the end of the fermentation cycle by centrifugation at 10,000 rpm
for 10 min at 20 °C. The microspheres, consisting of encapsulated cells, were
recovered by filtration in vacuo using a Buchner funnel lined with filter paper,
(Whatman No. 54, GE Healthcare Life Sciences, New Jersey, USA). The free
cells and microspheres were then introduced into fresh fermentation medium
respectively. The same procedure was repeated till no more butanol could be
detected in the fermentation medium. For each fermentation cycle, aliquots
were withdrawn aseptically from the fermentation medium at specific time
intervals for the determination of viable cell count, butanol yield and stability
of the gellan gum microspheres. Each experiment was conducted in triplicate.
B.9 Pretreatment of S. saman leaf litter
B.9.1 Mechanical pretreatment (milling) of S. saman leaf litter
The dried leaves were first chopped in a hammer mill (Fitz mill, Illinois,
USA), followed by pulverization in a disintegrator mill (Laboratory mill,
Essex, England). The milled leaves (300 - 1000 microns) were stored in a
sealed plastic bag at room temperature and kept in a desiccator till required.
71
B.9.2 Hydrothermal pretreatment of S. saman leaf litter
Milled leaves were dispersed in 20 mL of distilled water to give a final
substrate concentration of 3.33 %, w/w. The dispersion was then subjected to
high temperature (121 °C) and pressure (15 psi) for 30, 60 and 90 min
respectively (Hideno et al. 2012). At the end of the pretreatment, the samples
were cooled to room temperature and filtered using a Buchner funnel lined
with filter paper, (Whatman No. 54, GE Healthcare Life Sciences, New Jersey,
USA) to separate the pretreated solids from the liquid. The filtrate was then
assayed for reducing sugars by the DNS method (as described in section
B.10.3).
B.9.3 Dilute acid coupled with heat treatment of S. saman leaf litter
Milled leaves were dispersed in 20 mL of 1 %, w/w sulphuric acid to give a
final substrate concentration of 3.33 %, w/w. The dispersion was then
autoclaved at 121 °C and 15 psi for 30, 60 and 90 min respectively (Sun and
Cheng, 2005). At the end of the acid hydrolysis, filtration was carried out to
separate the solid fractions from the acid hydrolysate. The pH of the
hydrolysate was adjusted to neutrality using 10 NaOH. The fermentable sugar
concentration of the hydrolysate was then determined using the DNS method
(section B.10.3). The dilute acid coupled with heat treatment was carried out
in triplicate.
72
B.9.3.1 Optimisation of dilute acid coupled with heat treatment of S.
saman leaf litter
The dilute acid coupled with heat treatment was optimised using response
surface methodology (RSM) with a 32
full factorial experimental design
(Periyanan and Natarajan, 2013). The experimental design and statistical
analysis were performed using the software, Design Expert 7.1.3 (Stat-ease
Inc., MN, USA). The independent variables chosen were acid concentration
(0.5, 1.0 and 2.0 %, w/w) and treatment time (30, 60 and 90 min) as shown in
Table 8 and the response variables were fermentable sugar yield and percent
sugar recovery. Three levels for each factor were chosen, resulting in 9 runs
that were carried out in triplicate. The order of runs was fully randomised.
Table 8. Coded and uncoded values of the two independent factors in the
optimisation of dilute acid coupled with heat treatment
Independent variable Unit Levels
-1 0 +1
A (Acid concentration) %, w/w 0.5 2.75 5.0
B (Treatment time) min 30 60 90
The model equation used to describe the effect of the independent variables on
the response variable is as follows:
73
Y = BA2B
2AB (5)
where Y is the response variable, A and Bareprocess variables and
arecoefficients.
The effects of the independent variables on the two responses were analysed
using response surface plots and contour plots. In addition, ANOVA was
performed to determine the statistical significance of the model with p value
set at 0.05.
Sugar yield was defined as the total sugar quantity obtained from a specific
quantity of leaves. The percent recovery of fermentable sugars was calculated
as follows:
Sugar recovery =
(6)
where X is the quantity of fermentable sugars obtained and Y is the total
amount of fermentable sugars available (i.e. sugar content) in the same weight
of substrate. The fermentable sugar content of the substrate was determined
using the method described in section B.10.3
B.9.3.2 Determination of fermentable sugar content in S. saman leaf litter
The procedure for determination of the fermentable sugar content present in S.
saman leaf litter was adapted from the National Renewable Energy
Laboratory’s ( REL) Laboratory Analytical Procedures (LAP) for the assay
of structural carbohydrates (Sluiter et al., 2008). This assay method involved a
two-step acid hydrolysis procedure. In the first step, 300 mg of the milled
74
leaves was hydrolysed with 3 mL of 72 %, w/w sulphuric acid for 60 min at
30 °C. The resultant mixture was diluted to 4 %, w/w acid and the second
hydrolysis step was carried out at 121 °C for another 60 min. The pH of the
hydrolysate was then neutralised using 1 N NaOH and the supernatant used for
the determination of total amount of fermentable sugar content by the DNS
method (section B.10.3).
B.10 Assay of fermentable sugars by DNS method
B.10.1 Preparation of citrate buffer
A quantity of 210 g of citric acid monohydrate was dissolved in 710 mL of
distilled water to give a 1M citrate buffer. The pH of the solution was adjusted
to 4.3 using approximately 50-60 g of NaOH. The final volume was made up
to 1000 mL with distilled water. For the preparation of 0.05 M citrate buffer,
the 1 M citrate buffer was appropriately diluted. The pH of 0.05 M citrate
buffer was 4.8.
B.10.2 Preparation of DNS reagent
An amount of 10.6 g of dinitrosalicylic acid was weighed and dispersed in
1416 mL of distilled water, followed by the addition of sodium hydroxide
(19.8 g). The mixture was stirred to obtain a clear solution. Approximately
306 g of sodium potassium tartrate was then added. The function of sodium
potassium tartrate salt was to prevent oxygen from dissolving in the solution
as this might affect the reaction. Further, 7.6 mL phenol and 8.3 g of sodium
metabisulfite were added to the above solution to intensify the colour of the
75
reaction and stabilize the reaction respectively. The prepared mixture was then
clarified by filtration before use.
B.10.3 Assay of fermentable sugars
The fermentable sugar contents of the test samples were determined using the
dinitrosalicylic acid (DNS) method (Miller, 1959). An appropriate amount of
DNS reagent was added to a known amount of the test sample and the mixture
heated at 90 °C for 5 min for colour development. The mixture was then
centrifuged to collect the supernatant, which was appropriately diluted for
spectrophotometric measurement (UV-1900, UV-VIS spectrophotometer,
Hitachi, Japan) at 540 nm. Glucose standards were also prepared and assayed
in accordance with the above procedure to obtain a calibration plot for
quantification of fermentable sugars in the test samples (section B.10.4). The
assays were carried out in triplicate.
B.10.4 Preparation of standard calibration curve for glucose
A stock solution containing 10 mg/mL of anhydrous glucose in 0.05 M citrate
buffer was prepared. Known dilutions (Table 9) were prepared as standards.
To each dilution, 3 mL of DNS solution was added and the mixture heated at
90 °C for 5 min. The solution was cooled and appropriately diluted to measure
the absorbance value at 540 m for each dilution using UV-VIS
spectrophotometer. A standard calibration curve was then constructed by
plotting amount of glucose against absorbance at 540 nm.
76
Table 9. Preparation of different dilutions of glucose for standard
calibration curve
Glucose stock
solution (mL)
0.05 M citrate
buffer (mL)
Dilution Conc of glucose
(mg/mL)
1 0 - 10
1 0.5 1:1.5 6.7
1 1.0 1:2 5.0
1 2.0 1:3 3.3
1 4.0 1:5 2.0
B.11 Determination of filter paper activity of Accellerase® 1500
The filter paper activity of Accellerase® 1500 was determined using the
method described by Ghose, 1987. In this method, the substrate used for
measuring the filter paper activity of Accellerase® 1500 was approximately
50 mg of Whatman No. 1 filter paper (1.0 mm X 6.0 mm strip). A 0.5 mL of
each enzyme dilution was added to 1 mL of 0.05 M citrate buffer in test tubes,
which were placed in a water bath at 50 °C. One filter paper strip was then
added to each test tube and incubated at 50 °C for 60 min. Three dilutions of
Accellerase® 1500 in 0.05 M citrate buffer were prepared such that at least
one dilution released slightly more and another slightly less than 2.0 mg
(absolute amount) of glucose in the reaction conditions. Apart from the test
solutions, enzyme blank (consisting of 1.5 mL of citrate buffer and a filter
paper strip), substrate blank (consisting of 1.0 mL of citrate buffer and 0.5 mL
of diluted enzyme) and control (consisting of 1.5 mL of citrate buffer) were
also incubated at 50 °C for 60 min. At the end of 60 min, the reaction was
77
stopped by the addition of 3 mL of DNS reagent. The sugar content in all the
tubes were then determined using the DNS method (section B.10.3). The
enzyme blank reading was subtracted from each reading to account for sugars
present inherently in the enzyme preparation. A plot of glucose liberated
against enzyme concentration was constructed and the enzyme concentration
required to release 2.0 mg of glucose extrapolated from the graph.
The FPU of the enzyme was then calculated as follows:
FPU (units/mL) =
(7)
B.12 Enzymatic hydrolysis of pretreated S. saman leaf litter
The milled leaves (without dilute acid coupled with heat treatment) were
subjected to enzymatic hydrolysis using the Accellerase® 1500 at doses
ranging from 7-35 FPU/g of substrate. A specific quantity of the milled leaves
was dispersed in 0.05 M citrate buffer (pH 4.8) and incubated with the
different doses of enzyme at 50 °C for 48 h. For leaves subjected to
hydrothermal or acid pretreatment, the solids were first separated from the
liquid hydrolysate by filtration in vacuo, washed thoroughly with distilled
water and dried at 60 °C. The solids were then dispersed in 0.05 M citrate
buffer and then subjected to enzymatic hydrolysis with an enzyme dose of 20
FPU/g substrate using similar incubation conditions for untreated leaf litter.
At the end of the enzymatic hydrolysis, the fermentable sugar concentration
78
was determined using the DNS method (section B.10.3). The enzymatic
hydrolysis was carried out in triplicate.
B.13 Detoxification of acid hydrolysate of S. saman leaf litter
The acid hydrolysate was separated from the solids by filtration in vacuo. The
clear filtrate was then subjected to different detoxification methods so as to
make it amenable for bacterial fermentation. The pH of the filtrate was noted
before and after detoxification.
B.13.1 Treatment of the acid hydrolysate with sodium hydroxide
The pH of acid hydrolysate was adjusted to 5-6 using 10 N NaOH. The
mixture was stirred thoroughly and allowed to stand for 30 min. Reducing
sugar content of the filtrate was determined by DNS method (section B.10.3)
B.13.2 Treatment of the acid hydrolysate with sodium hydroxide and high
temperature
The clear supernatant obtained (section B.13.1) was further subjected to
heating at 90 °C for 30 min using rotoevaporation. The solution was allowed
to cool to room temperature and the reducing sugar content of the filtrate was
determined by DNS method (section B.10.3)
B.13.3 Treatment of the acid hydrolysate with calcium hydroxide
(overliming)
The pH of the acid hydrolysate was adjusted to 10.5 by the addition of
Ca(OH)2 . The resultant mixture was stirred thoroughly and allowed to stand
79
for 30 min. The pH of the filtrate was adjusted to 5‐6 using hydrochloric acid
before use as fermentation substrate. Reducing sugar content of the filtrate was
determined by DNS method (section B.10.3)
B.13.4 Treatment of the acid hydrolysate with calcium hydroxide
(overliming) and high temperature
The solution obtained (section B.13.3) was further subjected to heating at 90
°C for 30 min using rotoevaporation. The solution was allowed to cool to
room temperature and the reducing sugar content of the filtrate was
determined by DNS method (section B.10.3)
B.14 Fermentation of detoxified leaf hydrolysate by Cl. acetobutylicum
ATCC 824
The efficiency of the various detoxification methods was evaluated by
measuring the fermentability of the detoxified acid hydrolysate of S. saman
leaf litter. The RCM broth was supplemented with the filtration-sterilised
detoxified hydrolysate. The resultant fermentation medium was inoculated
with Cl. acetobutylicum ATCC 824 cell suspension containing 107
cells/mL or
an equivalent number of encapsulated spores of Cl. acetobutylicum ATCC
824. Fermentation was carried out at 37 °C for 144 h. Samples were
withdrawn at intervals for determination of residual glucose concentration,
butanol yield and productivity. Experiments were performed in triplicate.
80
B.15 Statistical analysis
One-way ANOVA and t-test were carried out on the results using Minitab 14
(Minitab Inc, Pennsylvania, USA). Independent sample t test was used to
compare two sample sets. One-way ANOVA was used to compare more than
two sample sets with Tukey’s test as the post hoc analyses. The level of
significance in this study was set at p < 0.05 unless otherwise stated. The
factorial design and response surface optmisation studies were performed
using the statistical software Design Expert 7.1.3 (Stat-ease Inc., Minneapolis,
USA).
81
RESULTS AND DISCUSSION
82
IV. RESULTS AND DISCUSSION
PART ONE
A. Cultivation of Cl. acetobutylicum ATCC 824
It is important to determine the suitable conditions for growth of the test
microorganism before any study is initialised. Various factors such as
composition of growth medium, pH, and incubation conditions have an effect
on the growth of bacteria (Adamberg et al., 2003; Mayo and Noike, 1996;
Monot et al., 1982). Most bacteria belonging to clostridium species grow
within the pH range of 6 - 7, but some grow at pH below 4 or above 8
(Blaschek, 1999; Gibson and Roberts, 1986; Johnson, 2009; Keis et al., 2001).
The growth temperatures recommended for clostridium species range from 25
– 37 °C (Blaschek, 1999; Johnson, 2009). (In addition, special considerations
are needed for providing suitable anaerobic environment for the growth of
clostridial cells.
The selection and optimisation of suitable conditions for the growth of Cl.
acetobutylicum ATCC 824 was thus carried out with respect to media and
incubation conditions. This was followed by the determination of the growth
curve of the bacterium and microscopic examination of the morphological
changes during different phases of growth. Finally, the conditions for the
revival of the bacterial spore were optimised.
83
A.1 Suitable media for the growth of Cl. acetobutylicum ATCC 824 cells
An ideal cultivation medium for the growth of clostridial cells should consist
of a mixture of organic and inorganic nitrogen sources, sugars, proteins and
mineral salts (Bahl et al., 1986; Monot et al., 1982). In addition, clostridium
species require low redox potential for cellular metabolism and growth. Hence
the media should have reducing agents to maintain the requisite low redox
potential.
In this study, Cl. acetobutylicum ATCC 824 was grown in two different types
of media, namely reinforced clostridial medium (RCM) and thioglycollate
medium (TGM). The two media are examples of complex media composed of
various essential nutrients for the growth of the bacterium. In addition, RCM
and TGM also contain reducing agents necessary for the maintenance of
anaerobic conditions i.e. cysteine hydrochloride and sodium thioglycollate
respectively. RCM is recommended by ATCC for the cultivation of
clostridium species. In terms of cost, RCM is more expensive than TGM. In
order to examine the possibility of reducing the cost of the overall process, the
cheaper medium (TGM) was also explored as an alternate cultivation medium
to cultivate Cl. acetobutylicum ATCC 824.
A continuous streak of dense bacterial colonies was apparent on RCM agar
plate after 24 h of incubation while TGM agar plates showed relatively sparse
colonies after 48 h of incubation (Figures 11 and 12). In the case of broth
cultures, turbidity was apparent in RCM broth after 20 h of incubation while
TGM broth showed growth only after 28 h. The average viable count of the
84
RCM broth culture was markedly higher than that of the TGM broth (Table
10). Longer incubation period of 72 h did not offer any significant advantage
to the bacterial growth. It has been reported that pH, incubation temperature,
and presence or absence of certain media components can alter the growth of
clostridial species (Gibson and Roberts, 1986). Although both RCM and TGM
have been used for the growth and cultivation of anaerobic bacteria, they have
significantly different compositions. In the case of RCM, peptone, beef extract
and yeast extract provided the amino acids, nitrogen and vitamins while
glucose provided the carbon needed for bacterial growth. The osmotic balance
of the medium was maintained by sodium chloride and soluble starch
detoxified metabolic by-products. A low concentration of agar was also
present to minimize diffusion of oxygen into the medium. The pH of the final
medium was in the range of 6.6 - 7.0 which was maintained by sodium
acetate. The combination of these ingredients enabled a rapid growth of the
Cl. acetobutylicum ATCC 824 cells. On the other hand, for TGM, tryptone,
yeast extract and glucose served as the nitrogen, vitamin and carbon sources,
respectively. It did not contain soluble starch or beef extract. The final pH of
TGM was also higher (6.9 - 7.3) than that of RCM and there were no buffering
agents. Thus, TGM was less suitable than RCM for the growth of Cl.
acetobutylicum ATCC 824, which also suggested that the bacterium was a
relatively fastidious organism. Based on the preliminary findings, RCM was
selected as the cultivation medium for the rest of the studies.
85
Table 10. Viable count of Cl. acetobutylicum ATCC 824 in different cultivation broths
Cultivation broth Average number of colony forming units per mL (CFU/mL)
After 24 h After 48 h After 72 h
RCM 3.46 (± 0.25) × 107 1.45 (± 0.38) × 10
7 1.07 (± 0.17) ×10
7
TGM No evident turbidity 3.18 (± 1.12) × 103 1.65 (± 1.05) ×10
3
Figure 11. Cl. acetobutylicum
ATCC 824 colonies on RCM agar
after 24 h of incubation at 37 °C
Figure 12. Cl. acetobutylicum
ATCC 824 colonies on TGM agar
after 48 h of incubation at 37 °C
86
In addition, it was found that Cl. acetobutylicum ATCC 824 grew only when
the agar plates and broth cultures were incubated at 37 °C. It has been reported
that certain functional bacterial enzymes required for essential biochemical
reactions could only be activated around at 37 °C (Madigan et al., 2000). This
probably accounted for the lack of cell growth at low incubation temperatures.
Overall, it was concluded that incubation temperature and type of media were
critical factors affecting the growth of Cl. acetobutylicum ATCC 824.
A.2 Suitable set-up for the growth of Cl. acetobutylicum ATCC 824
The clostridium species are known to be strict anaerobes, implying that they
can grow only in the absence of atmospheric oxygen since most of them lack
defences against reactive oxygen (Finegold et al., 2002; Johnson, 2009).
Oxygen interferes with the vital enzyme systems and with nucleic acids of the
clostridium species (O'Brien and Morris, 1971). However, few reports have
stated that some species in this genus can be microaerophilic. For instance, the
vegetative cells of Cl. bifermentans could survive several hours to oxygen
exposure (Jayasinghearachchi et al., 2010).
Various methods for generating anaerobic or microaerophilic conditions for
the growth of Cl. acetobutylicum had been investigated. The methods included
the use of anaerobic glove boxes, flushing the media with nitrogen gas and use
of reduced media (Johnson, 2009; Lee et al., 1985; Lepage et al., 1987; Stim-
Herndon et al., 1996).
87
Gas generating sachets are compact systems capable of generating suitable
conditions for the growth of anaerobic bacteria. The active components of
these gas generating systems may consist ascorbic acid or activated carbon.
When placed in a sealed jar, gas generating sachets produce carbon dioxide
and simultaneously absorb the oxygen in the jar. Thus, a suitable environment
is generated for the growth of anaerobic microorganisms.
In this study, two types of gas generating systems, Campygen™ and
Anaerogen™ were investigated. The Anaerogen™ sachet reduces the oxygen
level in the jar to below 1 % within 30 min while producing about 9 - 13 %
carbon dioxide content (Ruangrungrote et al., 2008). The Campygen™ sachet
creates an environment containing 5 % oxygen, 10 % carbon dioxide, and 85
% nitrogen. Unlike other gas generating systems, Campygen™ and
Anaerogen™ do not produce hydrogen gas, thereby eliminating the risk of
explosion.
It was observed that Anaerogen™ in a sealed desiccator could provide
adequate anaerobic conditions for the growth of Cl. acetobutylicum ATCC
824 in both solid and liquid media (Table 11). The former showed bacterial
colonies within 36 h of incubation while the latter showed turbidity along with
gas bubbles, due to fermentation, after 20 h of incubation (Figures 13a and
13b). o growth was observed in the media incubated with Campygen™. The
broth cultures could grow well in tightly closed universal bottle without any
gas pack system. On the other hand, no growth was seen in plates and bottles
88
incubated with Campygen™ indicating its inefficiency in generating sufficient
anaerobic conditions for the growth of Cl. acetobutylicum ATCC 824.
It was thus concluded that Cl. acetobutylicum ATCC 824 could grow only in
very low oxygen environments, when the oxygen concentrations were less
than 1 % as attained by Anaerogen™. The viable count in tightly closed
bottles was almost the same as that obtained with Anaerogen™ (Table 11).
The cultivation medium, RCM, contained a reducing agent, cysteine
hydrochloride, capable of reducing the redox potential of the media. The
results implied that the combination of tightly closed bottles and a reducing
agent in the cultivation medium was sufficient in providing the anaerobic
conditions necessary for the growth of Cl. acetobutylicum ATCC 824. Hence,
in subsequent studies, agar cultures were incubated using Anaerogen™ while
broth cultures were incubated in tightly closed bottles.
Table 11. Effects of different incubation set-ups on the cells growth of Cl.
acetobutylicum ATCC 824
Incubation set-up Average number of colony forming units per mL
(CFU/mL) after 48 h of incubation
Anaerogen™ 4.5 (± 0.29) × 107
Campygen™ Nil
Tightly capped
bottles
2.78 (± 0.61) ×107
89
Figure 13. Growth of Cl. acetobutylicum ATCC 824 in (a) RCM agar and
(b) RCM broth incubated under anaerobic conditions maintained by
Anaerogen™
A.3 Growth curve of Cl. acetobutylicum ATCC 824 in RCM
Clostridium acetobutylicum ATCC 824 was inoculated in 300 mL of RCM
broth and incubated at 37 °C under anaerobic conditions maintained by
Anaerogen™. The viable count and optical density (at 600 nm) of the culture
was determined at regular time intervals, from the first time point of 6 h.
When a particular organism is introduced into a cultivation medium, growth of
the organism takes some time as it prepares for synthesis of enzymes and
DNA for cell division. This is known as the lag phase. It was not reflected in
the growth curve obtained (Figure 14a). The first time point (6 h) chosen in
the determination of the growth curve was too late to detect the lag phase. This
also indirectly showed that the cells were readily revived under the optimal
incubation conditions employed. Following the lag phase, the growth of the
bacteria increased exponentially. The highest cell count was reached at around
(a) (b)
90
18 h. Thus, the growth duration of 6 - 18 h denoted the exponential phase of
growth of Cl. acetobutylicum ATCC 824. There was no significant increase in
viable cell count observed from 18 – 42 h. This phase was the stationary phase
of bacterial growth where the number of new cells produced was equally
matched by cell death. The stationary phase could be attributed to the
depleting nutrients and accumulation of harmful metabolites. Following the
stationary phase, there was a decline in viable cell count which continued till
the end of incubation and sampling time (54 h). This decline phase
corresponded to the accumulation of high concentration of harmful
metabolites in the cultivation medium.
The optical density (indicated by absorbance at 600 nm) increased
correspondingly with cell count in the exponential phase of growth (Figure
14b). The optical density continued to increase, albeit to a smaller extent, even
when the culture entered the stationary phase. This could be due to the
formation of swollen clostridial cells during the stationary phase. These cells
are known to increase in mass due to accumulation of granulose as reserve
nutrient source (Jones et al., 1982). Apart from the decrease in viable cell
count during the decline phase, there was also lysis of the dead cells. The latter
accounted for the corresponding decrease in optical density during the decline
phase of the culture.
The relationship between the optical density and log CFU/mL aided in
understanding the growth characteristics of the bacterium. This relationship
could be expressed by the linear equation y = 0.28x - 0.921 (R2
= 0.974)
91
(Figure 15). This equation was employed in subsequent studies to approximate
the concentration of viable count of the sample from its optical density which
could be easily measured.
Figure 14. Cultivation of Cl. acetobutylicum ATCC 824 in RCM broth at
37 °C: (a) growth curve and (b) optical density of culture
Figure 15. Relationship between optical density and viable count of Cl.
acetobutylicum ATCC 824 culture in RCM broth
0
2
4
6
8
10
12
0 20 40 60
Lo
ga
rith
mic
nu
mb
er o
f
cell
s/m
L
Incubation time (h)
y = 0.28x - 0.921
R² = 0.974
0
0.5
1
1.5
2
2.5
0 2 4 6 8 10 12
OD
at
600 n
m
Logarithmic number of cells/ mL
(a)
0
0.5
1
1.5
2
2.5
0 20 40 60
OD
at
60
0 n
m
Incubation time (h)
(b)
92
A.4 Morphological changes in Cl. acetobutylicum ATCC 824 cells during
different phases of growth
There was a sequential change in the morphology of Cl. acetobutylicum
ATCC 824 with time. The cells also exhibited age-associated variability in
Gram staining characteristics (Johnson, 2009; Jones et al., 1982). The cells in
the exponential phase were predominantly Gram-positive rods (Figure 16a).
The cells in this phase of growth are known to be associated with acid
formation (Jones et al., 1982). In the stationary phase, both Gram-positive and
Gram-negative rods were found (Figure 16b). In addition, a large number of
the cells appeared swollen with granulose. Conversion of acids to solvents has
been reported to be correlated with the appearance of swollen cigar-shaped
clostridial cells characterized by accumulation of granulose (Jones and Woods,
1986). Many endospore-containing cells (i.e. forespores) as well as free spores
were also observed. The endospores appeared oval and sub-terminal. Cells in
the decline phase were mostly Gram-negative rods (Figure 16c).
93
Figure 16. Gram staining of Cl. acetobutylicum ATCC 824 cells in (a)
exponential, (b) stationary and (c) decline phase of growth
94
A.5 Optimisation of heat shock treatment for the revival of Cl.
acetobutylicum ATCC 824 spores
Clostridia produce spores as a defence mechanism to withstand unfavourable
environmental conditions. These spores are very resistant to heat, radiation,
drying and chemicals (Johnson, 2009). Under favourable environmental and
nutritional conditions, spores can undergo a revival process which involves
activation, germination and outgrowth of the spores to form vegetative cells.
Heat shock treatment is a commonly used method for revival of clostridial
spores (Tashiro et al. 2005, Noomtim and Cheirsilp, 2011). Apart from heat
shock treatment, other methods reported in the literature for the revival of
bacterial spores include the use of chemicals such as calcium chloride, chloral
hydrate, dimethyl sulphoxide, and dimethyl formamide (Lee and Ordal, 1963).
L-alanine and a combination of L-asparagine, D-glucose and D-fructose have
also been reported to trigger sporulation in Bacillus subtilis (Moir and Smith,
1990). It is possible that the chemical agents used for spore revival may
interfere with the fermentative ability of the revived cells as well as affect
subsequent assay of butanol during GCMS. Thus, heat shock treatment was
selected for the revival of clostridial spores owing to its simplicity and
effectiveness.
Heat shock treatment involves subjecting the spores to conditions of high
temperature for a brief period. Heat activates the dormant spores, thus
initiating its metabolic activity (Mitchell, 1997; Sauer et al., 1995) Increase in
95
the metabolic activity of the spores results in loss of resistance to
environmental stress. The spore coat is ruptured or absorbed, followed by
swelling of the endospore and its development into a fully functional
vegetative bacterial cell.
Different conditions of heat shock treatment have been used for the revival of
clostridial spores owing to the fact that heat resistance of clostridial spores is a
function of species (Sarathchandra et al., 1977). In this study, different
combinations of high temperature and time were investigated. The condition
that was able to give maximum spore revival in the shortest time was deemed
to be optimal for the heat shock treatment.
No growth was seen for tubes incubated without heat shock treatment,
implying that heat shock treatment was necessary to revive the spores (Table
12). Maximum spore revival was obtained when the spores were exposed to
80 °C for 10 min prior to incubation at 37 °C. Very little or no cell growth was
observed at 70 °C, indicating that lower temperature was ineffective in
reviving the spores. It has been reported that harsh heat shock treatment
conditions may destroy the spores of certain species (Sarathchandra et al.,
1977). This observation could be used as an explanation for the low viable
counts obtained for spores revived using high temperatures. The optimal
condition determined (80 °C for 10 min) was employed to revive the spores in
subsequent studies.
96
Table 12. Viable count of Cl. acetobutylicum ATCC 824 spores revived by
different heat shock treatment conditions
B. Optimisation of microsphere production using Design of Experiments
(DoE)
The microencapsulation process used in this study was based on a modified
emulsification method developed by (Wan et al., 1992). Many studies have
been carried out on the encapsulation of drugs by the emulsification technique
consisting of an aqueous gel forming solution in an immiscible phase, usually
composed by an organic solvent or oil (Aghbashlo et al., 2012). High shear
forces are utilised to enable emulsification and subsequent entrapment of the
drug in the polymeric material used (Belyaeva et al., 2004; Heng et al., 2003).
The droplets formed during emulsification process are stabilised by the
addition of surfactant having suitable hydrophilic-lipophilic balance (HLB)
Temperature Viable count of revived cells (CFU/mL)
3 min 5 min 10 min 15 min
70 °C No growth No growth 2.2(±0.99)
×103
1.0 (± 0.34)
×103
80 °C No growth 4.79 (±1.09)
×105 3.26(±0.33)
×106
1.03 (±1.56)
×106
90 °C 1.99 (±0.55)
×104
2.04 (±0.23) ×
104
3.99 (± 0.29)
×105
2.77 (±0.47)
×105
100 °C 1.26 (±0.28)
×103
1.77 (±0.32) ×
103
1.06 (±0.45)
×103
1.15 (±0.37)
×103
Control No growth
97
value (Wan et al., 1994). Microencapsulation of clostridial cells by
emulsification technique has yet to be investigated as a cell immobilisation
method. In this study, the feasibility of microencapsulation of Cl.
acetobutylicum ATCC 824 cells in gellan gum microspheres using
emulsification method was evaluated.
The selection and optimisation of various process and formulation parameters
such as the type and concentration of polymer, type and amount of surfactants,
stirring speed, processing temperature and concentration of congealing agent
during microencapsulation are crucial for the development of robust and well-
formed microspheres (Heiskanen et al., 2012).
Factorial designs are useful for evaluating treatment variations, thus providing
greater precision in estimating the overall main and interaction effects of
different factors (Fontana et al., 2000). A 24 full factorial design was used to
study the effects of different formulation and process variables viz.
concentration of polymer, HLB of surfactant blend, temperature of
emulsification and stirring speed on the physical characteristics of the gellan
gum microspheres produced. Other factors such as concentration of cross-
linking agent, volume of organic phase and stirring time were set at fixed
values. In addition, the optimal condition for the production of microspheres
with desirable physical characteristics was determined using model equations
and desirability function. Before the application of the experimental design,
several preliminary trials were conducted to determine the range of conditions
at which the process was capable of forming microspheres with acceptable
98
physical characteristics. Based on the results, low, intermediate and high
levels of the variables were used in the factorial design. The mathematical
modelling of the relation between the independent variables and the responses
resulted in the response surface plots. Table 13 shows the different sets of the
experiments carried out, which include the different factors and their levels
along with the values of their respective responses
Table 13. Factorial design matrix employed in the optimisation study for
the microencapsulation process
Run A
B C D Mean size
(microns)
Span Aggregation
index (%)
1 1 10 40 800 38.3 2.0 4.5
2 2 10 60 400 154.5 1.9 7.6
3 2 8 60 800 45.5 1.8 5.2
4 1 8 40 800 33.6 1.9 7.8
5 1 10 60 800 25.5 1.5 13.3
6 2 8 40 400 127.7 1.4 5.9
7 2 8 40 400 128.7 1.3 4.8
8 2 8 60 800 35.1 1.7 4.4
9 1 10 40 400 103.4 1.8 11.1
10 1 8 60 400 140.3 1.7 11.6
11 2 10 60 800 60.7 1.6 9.8
12 1 10 60 400 132.2 1.9 9.3
13 1 10 60 400 179.8 1.8 10.3
14 1 8 40 400 128.7 1.7 13.6
99
15 1 10 40 400 82.5 1.8 11.1
16 1 10 40 800 29.2 1.9 5.7
17 1.5 9 50 600 58.0 1.6 2.9
18 1 8 40 800 35.7 1.8 8.3
19 2 10 60 800 51.4 1.5 9.8
20 1 8 40 400 106.9 1.6 13
21 2 8 60 400 132.8 1.6 8.3
22 1.5 9 50 600 78.0 1.8 3.1
23 1 8 60 800 38.1 1.7 13.6
24 1 8 60 800 19.8 1.7 15.6
25 2 10 60 400 127.8 1.8 6.9
26 1 8 60 400 138.5 1.4 10.2
27 2 10 40 800 60.6 1.5 12.1
28 2 8 60 400 159.0 1.6 7.9
29 1.5 9 50 600 66.0 1.6 3.4
30 2 10 40 800 60.8 1.5 10.3
31 2 8 40 800 88.7 1.8 16.0
32 2 8 40 800 78.7 1.7 15.2
33 2 10 40 400 122.2 1.8 10.6
34 2 10 40 400 159.4 1.8 9.2
35 1 10 60 800 34.5 1.6 10.4
*A: Concentration of gellan gum (%, w/v), B: HLB of surfactant blend,
C: Temperature of emulsification (°C), D: Stirring speed (rpm)
100
B.1 Influence of the variables on size
The size of the microspheres produced using various combinations of the
process and formulation variables ranged from 25.5 to 179.8 microns. The
extent of size reduction that was attained during emulsification was dependent
on a range of formulation factors such as the viscosity of the dispersed and
continuous phases, stirring speed and the interfacial tension between the two
phases (Heiskanen et al., 2012). Since the viscosity of isooctane was
negligible compared to that of gellan gum, the major effect on size of the
microspheres was due to the gellan gum solution viscosity. Increasing the
polymer concentration increases the viscosity of the dispersed phase. This
yields larger microspheres at the same mixing intensity because higher shear
forces are necessary to break up the more viscous droplets (Yang et al., 2000;
Yang et al., 2001). This implies that as the concentration of gellan gum and
subsequently its viscosity increased, a higher energy level of emulsification
was required to disperse this high viscosity solution. This was evident from
the response surface plot (Figure 17a). As the concentration of gellan gum
increased, there was a corresponding increase in the size of the microspheres.
Similar results were obtained in another study wherein increasing the polymer
concentration led to increase in microsphere size (Silva et al., 2006). Stirring
speed also had a significant main effect on the size of gellan gum
microspheres produced. Increasing the stirring speed from 400 to 800 rpm led
to approximately four-fold decrease in the microsphere size (Figure 17b). This
can be attributed to higher shear forces applied at higher rotational speeds,
101
resulting in a more finely dispersed emulsion leading to the formation of
smaller microspheres upon gelation.
B.2 Influence of the variables on span
Emulsification method for microsphere production usually yields small-sized
microspheres with a broad size distribution (Rabanel et al., 2009). The span of
the microspheres obtained in this study ranged from 1.3 to 2.0. Concentration
of gellan gum was found to have a significant effect on span of the
microspheres. It was observed that the span value was lower when higher
concentration of polymer was used. It may be hypothesised that the emulsion
droplets harden into microspheres relatively faster at lower polymer
concentration compared to higher polymer concentration. This probably
caused the spheres to have a higher size distribution because shearing forces
induced by the stirrer have limited effect on them once the microspheres were
formed.
In addition, there was a significant interaction effect of HLB and stirring speed
on span. At lower HLB, increase in stirring speed led to increase in span value
whereas at higher HLB, increase in stirring speed reduced the span value.
(Figure 17c). The size distribution of the microspheres obtained by
emulsification method is correlated with the size distribution of the emulsion
droplets which in turn are influenced by various parameters such as apparatus
design, viscosity of the two immiscible phases and stirring speed (Silva et al.,
2006). Increase in stirring speed lead to decrease in droplet size as was
observed in the previous study. The effect of higher stirring speed on span
102
value may be caused by the distribution of turbulent throughout the emulsion
(Lim et al., 1997). Since, shear stress is higher at the tip of the propeller than
at the centre, a faster stirring speed increased this difference, thereby
providing a less uniform distribution of energy and giving rise to microspheres
of a wider size distribution. On the other hand, at higher HLB value, the
stability of the emulsion was maintained by the surfactants and changes in
stirring speed. Stirring further enhanced the energy distribution thus yielding
microspheres with low size distribution.
B.3 Influence of the variables on aggregation index
Aggregation was found to be mainly affected by concentration of gellan gum
and stirring speed. At low polymer concentration, increase in stirring speed led
to decrease in aggregation index (Figure 17d). The opposite was observed at
high polymer concentration with increase in stirring speed. This can be
justified by increase in number of collisions phase during emulsification at
higher stirring speed, enhancing the adhesion of partially hardened
microspheres to each other thereby promoting formation of aggregates (Lim et
al., 1997).
B.4 Model equations and model adequacy
Regression equations were developed for each of the responses taking into
account the significant main and two factor interaction effects. Exceptions
were made only for terms which were essential to maintain the hierarchical
model; i.e. terms A, B, C, and D.
103
By substituting the coefficients Xi in Equation (1) (section B.3.1) by their
values (in terms of coded values of the factors) following equations for each of
the responses were generated, representing the best fitting model for each of
the responses. Positive signs indicate a synergistic effect while negative signs
indicate an antagonistic effect in the response. For example, concentration of
gellan gum had positive effect on size while stirring speed had a significant
negative effect on size.
Size = 89.07 + 9.89A - 0.15B + 3.14C – 43.7D– 6.27AC – 9.74CD
Span = 1.68 – 0.049A + 0.036B – 0.017C + 0.016D + 0.062AC – 0.029BC –
0.099BD – 0.046CD
Aggregation index = 9.8 – 0.8A – 0.29B – 0.16C + 0.34D + 0.82AB –
1.36AC + 1.01AD + 0.34BC–0.34BD
where A is the concentration of gellan gum (%, w/v), B is the HLB of
surfactant blend, C is the temperature of emulsification (°C) and D is the
stirring speed (rpm).
The model for each response variable was checked for goodness of fit using
various statistical parameters such as ANOVA, regression coefficient estimate
and sum of squares. The regression coefficient estimate for each response
value along with their respective sum of squares and p-values is given in Table
14.
104
Figure 17. Response surface plots of the effects of (a) temperature and
concentration of gellan gum on size and (b) concentration of gellan gum
and stirring speed on size
(a)
(b)
105
Figure 17. Response surface plots of the effects of (c) stirring speed and
HLB on span and (d) concentration of gellan gum and stirring speed on
aggregation index
(d)
(c)
106
Table 14. Coefficient estimate, sum of squares and their respective p-values for the three responses
Mean Size Span Aggregation index
Factor Coefficient
estimate
Sum of
squares
p-value Coefficient
estimate
Sum of
squares
p-value Coefficient
estimate
Sum of
squares
p-value
A 17.70 10026.2 < 0.0001 -0.049 0.076 0.0007 -0.80 20.29 < 0.0001
B 0.16 0.82 0.664 0.036 0.042 0.0074 -0.29 2.66 0.0704
C -3.42 375.2 < 0.0001 -1.7 X 10-2 8.95 X 10-3 0.1781 -0.16 0.78 0.3107
D -60.26 1.16 X 105 < 0.0001 1.6 X 10-2 8.10 X 10-3 0.199 0.34 3.64 0.037
AB -1.34 57.69 0.0016 -1.2 X 10-2 4.26 X 10-3 0.3465 0.82 21.70 < 0.0001
AC -6.58 1387.25 < 0.0001 0.062 0.12 < 0.0001 -1.36 58.86 < 0.0001
AD 2.09 139.64 < 0.0001 -1.3 X 10-2 5.13 X 10-3 0.3028 1.00 32.80 < 0.0001
BC 4.67 699.32 < 0.0001 -0.029 0.028 0.0242 0.34 3.63 0.0373
BD -0.42 5.63 0.2614 -0.099 0.31 < 0.0001 -0.34 3.71 0.0354
CD -8.14 2120.86 < 0.0001 -0.046 0.067 0.0012 0.30 2.83 0.0625
Residuals 196.23 0.0047 10.43
Lack of
fit
0.74 0.53 0.63
*A: Concentration of gellan gum (%, w/v), B: HLB of surfactant blend, C: Temperature of emulsification (°C), D: Stirring
speed (rpm)
107
The smaller the value of p, the more significant was the corresponding
coefficient. Sum of squares (SS) of each factor quantifies its importance in the
process and as the value of the SS increases, the significance of the
corresponding factor in the undergoing process also increases. It was found
that the estimated models of size had R2
value approximately one (0.9325)
while those of span and aggregation index were 0.9023 and 0.9605
respectively. This meant that more than 90 % of the variation in the response
variables could be explained by the independent variables while the remaining
10 % can be explained by unknown or inherent variability. Using the model
equations, the predicted and experimental values were compared with each
other using the linear correlation model. The experimental and predicted
values showed a good linear relation with predictive R2 values of 0.8435,
0.6308 and 0.8424 for size, span and aggregation index respectively. Thus it
could be concluded that the model describing size was found to be statistically
significant, explaining more than 90 % of the response while the models for
the span and aggregation index were acceptable with a predictive ability of 63
% and 84 % respectively. Thus, the models for all the three responses proved
to be adequate in describing the observed data.
B.5 Optimisation of formulation and process parameters in the
production of microspheres with the desired properties
In developing microspheres it is particularly important to produce
microspheres with reproducible mean size and span with low degree of
aggregation, especially for industrial scale-up purposes. Though small-sized
108
microspheres are preferred due to reduced mass transfer limitations, very
small microspheres may pose a problem when recovering them from the
fermentation medium (Pilkington et al., 1998). Thus, a balance must be
achieved so as to fulfil the above requirements. Preliminary studies with blank
microspheres showed that microspheres in the size range of 60-100 microns
could be recovered satisfactorily using Whatman filter paper no. 54 through in
vacuo filtration. The average size of Cl. acetobutylicum ATCC 824 cells is 1
to 4 microns (Long et al., 1984). Microspheres in the selected size range could
encapsulate sufficient number of cells to carry out the fermentation process.
Thus, microspheres with an average size of 60-100 microns were selected as
suitable size range.
Optimisation of the microencapsulation process was carried out by a multiple
response method which employed a desirability (D) function to optimise the
different combinations of formulation and process parameters such as
concentration of gellan gum, HLB of surfactant blend, temperature and stirring
speed as well as the respective polynomial equations obtained The goal of
optimisation was to produce microspheres in the desired size range with
minimum span and aggregation index. Based on the desired criteria for the
three response variables and the mathematical models, the optimal condition
for the production of microspheres was derived. The optimal formulation and
process parameters and the properties of the resultant microspheres consisted
of gellan gum concentration: 1.5 %, w/v, HLB: 8.05, Temperature of
emulsification: 60 °C and stirring speed: 600 rpm. These conditions produced
109
microspheres with mean size of 88.6 microns, mean span value of 1.7 and
mean aggregation index value of 9.6 %.
In order to validate the optimised production condition, production of
microspheres based on this condition and three additional random runs based
on conditions covering the entire range of experimental domain were
performed. For each of these test runs, responses were estimated by use of the
generated mathematical model and by the experimental procedures. Figure
18a-c shows linear correlation plots between the observed and predicted
response variables. The graphs demonstrate high values of correlation
coefficient (R2 > 0.9), indicating excellent goodness of fit. Therefore, it can be
concluded that the model functions for size, span and aggregation index were
well interpreted. Furthermore, the low magnitude of error (less than 10 %) for
all the three responses indicates the robustness of the respective models.
110
Figure 18. Correlation between observed and predicted values for : (a)
size, (b) span and (c) aggregation index of microspheres
y = 1.964x - 1.6311
R² = 0.939
1.58
1.63
1.68
1.73
1.78
1.83
1.64 1.66 1.68 1.7 1.72 1.74 1.76
Pre
dic
ted
sp
an
Actual span
y = 0.947x + 6.388
R² = 0.968
0
20
40
60
80
100
120
140
160
0 50 100 150
Pre
dic
ted
siz
e
Actual size
y = 1.1174x - 1.0684
R² = 0.9823
0
2
4
6
8
10
12
14
16
18
0 5 10 15 20Pre
dic
ted
aggre
gati
on
in
dex
Actual aggregation index
(c)
(b)
(a)
111
C. Effect of emulsification process on viability of Cl. acetobutylicum
ATCC 824 vegetative cells and spores
As mentioned previously, the purpose of cell encapsulation is to protect the
cells from harsh conditions during fermentation such as toxic metabolites and
shear forces prevailing in the fermentation medium. The preparation of
microspheres by the emulsification method involved chemicals and dispersive
forces (Silva et al., 2006). Not much information is available regarding
encapsulation of anaerobic bacteria using the emulsification technique.
However, similar techniques have been used for the encapsulation of other
aerobic or facultative anaerobes. For example, the aerobe, Lactobacillus
acidophilus, was encapsulated in poly-lactic-co-glycolic acid by supercritical
emulsion extraction method (Porta et al., 2012). The procedure resulted in
drastic reduction in the viability of the encapsulated cells. Similarly, the
viability of the facultative anaerobe, yeast, was reduced by approximately 10
times after emulsification to produce chitosan coated carrageenan
microspheres (Raymond et al., 2004).
In order to encapsulate Cl. acetobutylicum ATCC 824 cells using the
emulsification method, it was first necessary to determine whether the
bacterial cells could survive the emulsification process used for
microencapsulation. Cl. acetobutylicum ATCC 824 exists in different
morphological structures during the course of its growth. The cells in the
exponential stage are rod-shaped while those in the stationary phase are
usually swollen cigar-shaped sporulating cells. In the final stage of the growth
112
cycle, the spore is released from the mother cell. It was hypothesised that the
more robust spores were suitable than the vegetative cells for
microencapsulation by the emulsification process. To test this hypothesis, both
vegetative cells and spores were subjected to the conditions encountered
during the emulsification process and their viability before and after the
process determined. In this study, the suspension of vegetative cells or spores
was added directly (without gellan gum) into the continuous phase of
emulsion. Gellan gum was not used because it was difficult to disintegrate the
gellan gum microspheres to liberate the cells for viable count determination
without causing harm to the cells. It was observed from the viable count
results that the log reduction value of the vegetative cells was markedly higher
than that of the spores (Table 15).
Table 15. Effect of emulsification process on the viability of vegetative
cells and spores of Cl. acetobutylicum ATCC 824
Before
emulsification
After
emulsification
Log
reduction
Vegetative cells 1.07 (±0.22) × 107 2.20 (±0.81) × 10
2 4.69
Spores 1.2 (±0.40) × 107 2.56 (±1.25) ×10
6 0.67
The results clearly showed that the conditions encountered during
emulsification had a dramatic effect on the viability of vegetative cells and
most of the cells could not survive. Although Cl. acetobutylicum ATCC 824 is
classified as strict anaerobe in the literature, it was found to be able to survive
several hours of oxygen exposure (Nasuno and Asai, 1960). The vegetative
113
cells of Cl. acetobutylicum ATCC 824 were also successfully encapsulated by
the extrusion method (Häggström and Molin, 1980; Prüße et al., 1998).
Quantitative survival measurements using various laboratory strains belonging
to clostridia species showed that most of the cells lost viability slowly when
exposed to normal atmospheric conditions (Labbé, 1997). In the case of
microencapsulation by the emulsification technique, the total time required for
the emulsification process was one and half hour. Based on the findings of
(Nasuno and Asai 1960), the vegetative cells of Cl. acetobutylicum ATCC 824
should be able to withstand exposure to oxygen for the total time of
emulsification used in this study. Other factors involved in the emulsification
process such as chemicals and high shear may also have a negative effect on
the survival of the vegetative cells. Collectively, all the factors resulted in low
viability of the vegetative cells. On the other hand, the spores of Cl.
acetobutylicum ATCC 824 could withstand the conditions encountered during
emulsification owing to their robust nature. Further investigation to test the
fermentation ability of the revived spores, obtained after subjecting to the
emulsification process, and showed that they could produce equivalent
quantity of butanol when compared to the control. This meant that the spores
retained their viability and the fermentative ability of the regenerated
vegetative cells was not compromised. Hence the spores were suitable for
microencapsulation by the emulsification method. In addition, it is possible to
utilise a wider range of materials and conditions to encapsulate the spores
without drastic adverse effects on their viability.
114
D. Microencapsulation of Cl. acetobutylicum ATCC 824 spores by
emulsification method
The spores of Cl. acetobutylicum ATCC 824 were encapsulated in gellan gum
microspheres by the optimised emulsification method as described in section
B.5. The microspheres produced were discrete and spherical (Figure 19a).
Unencapsulated spores were not seen in the samples of freshly harvested
microspheres and washings. The size of microspheres was in the range of 60
to 100 microns, with low aggregation index of 2.3 to 3.7 %. The size of the
microspheres was large enough to accommodate the clostridial spores (1-3
microns). The translucent gellan gum microspheres allowed the entrapped
spores to be seen under the microscope (Figures 19b). The spores appeared
uniformly dispersed within the gellan gum matrix. It was clearly seen that the
spores were successfully encapsulated in the gellan gum microspheres by the
emulsification method using the pre-determined optimal processing
conditions. Thus this method could successfully encapsulate Cl.
acetobutylicum spores via emulsification in gellan gum. Encapsulation of both
vegetative cells and spores of Cl. acetobutylicum in calcium alginate beads
was carried out by Haggstrom and Molin, (1980). Since the method used for
the same was mild, the viability and fermentative activity of vegetative cells
were retained. It was concluded that the spore form of the bacterium gave
better results with respect to butanol yield. The spore form of the bacterium
could be revived into vegetative cells within the gel and the immobilised
preparation could be used for continuous operation.
115
Figure 19. Photographs of (a) blank gellan gum microspheres and (b)
gellan gum microspheres loaded with Cl. acetobutylicum ATCC 824
spores prepared using the optimised microencapsulation method
E. Optimisation of gas chromatography-mass spectrometry conditions for
the assay of butanol
Optimisation of the GC-MS analytical parameters was carried out by varying
the column oven temperature program, gas flow rate, gas flow pressure and
split ratio. The selection of the best conditions for analysis was based on the
shape and resolution of the analyte peak. A solution containing 5 g/L butanol
in 2-ethyl 1-hexanol (after suitable dilution) was injected into the GC column.
A narrow peak for butanol was obtained when the following condition was
employed: Initial oven temperature of 70 °C for 3 min, followed by increase to
200 °C at a rate of 50 °C per minute with pressure of 60.6 kPa, total flow of
103.1 mL/min and column flow of 0.99 mL/min.
E.1 Assay of butanol
Butanol was extracted from the fermentation medium using 2-ethyl 1-hexanol
as the extractant by liquid-liquid extraction. This solvent has been reported to
116
be a suitable solvent for the extraction of butanol (Heitmann et al., 2012). It
has a good distribution coefficient for butanol and is relatively cheap (Shukla
et al., 1989). Based on the results, it was found that the extraction efficiency
of 2-ethyl 1-hexanol was 70 %. In order to fully recover the butanol from the
fermentation medium, more extraction cycles were performed. Full extraction
of butanol from the fermentation medium using 2-ethyl 1-hexanol was
observed after three extraction cycles. Thus, it could not extract butanol
completely from the fermentation medium in a single extraction step. A
calibration plot for estimation of butanol produced in the fermentation medium
was therefore required.
Known amount of butanol was added to the fermentation medium. The
extracting solvent was then added to an equivalent volume of fermentation
medium and agitated for 5 min using a vortex mixture. The mixture was
centrifuged for 5 min at 6000 rpm resulting in the separation of upper phase
composed of 2-ethyl 1- hexanol with extracted butanol. The upper phase was
then removed for assay of butanol. An equivalent amount of 2-ethyl-1-hexanol
was added to the lower phase and the above procedure repeated up to three
times. The samples were then assayed for butanol by GC-MS. A calibration
plot of peak areas (area under curve) against the concentration of butanol in
g/L added to the fermentation medium was constructed (Figure 20). The peak
areas increased with butanol concentration in accordance with a linear
relationship expressed by y = 477267x- 58383. Butanol concentration in the
117
fermentation medium was estimated by extrapolating the peak areas of the test
samples in the equation.
Figure 20. Calibration plot for estimation of butanol concentration in
fermentation medium
F. Fermentation using free (non-encapsulated) cells of Cl. acetobutylicum
ATCC 824
Despite the common characteristics of various strains of solventogenic
clostridia, the nature of metabolic shift and kinetic pattern of solvent formation
are markedly strain-dependent because each strain exhibits its own intrinsic
genetic and metabolic characteristics (Blaschek, 1999). These factors result in
different solvent yields by the different strains.
Preliminary optimisation studies were carried out to determine the suitable
conditions for fermentation by non-encapsulated vegetative cells of Cl.
y = 477267x - 58383
R² = 0.999
0
1000000
2000000
3000000
4000000
5000000
6000000
7000000
8000000
0 5 10 15 20
Are
a u
nd
er c
urv
e (
a.u
.)
Butanol concentration (g/L)
118
acetobutylicum ATCC 824 using glucose as the fermentation substrate.
Different fermentation runs with various levels of inocula size, glucose
concentration and inocula age were carried out. The time required to obtain
maximum butanol yield from different fermentation runs varied from 48 to 72
h. Similar observations were reported by other workers (Jones and Woods,
1986; Keis et al., 2001). The time required to attain maximal butanol yield was
considered as the fermentation time. The optimisation studies for free (non-
encapsulated) vegetative cells of Cl. acetobutylicum ATCC 824 were carried
out as a series of 8 fermentation runs with various combinations of inoculum
size, inoculum age and glucose concentration. The results of are summarised
in Table 16.
F.1 Influence of glucose on fermentation efficiency
Different concentrations of glucose were investigated as substrate for butanol
fermentation by Cl. acetobutylicum ATCC 824 cells. A minimum
concentration of 6 %, w/v is usually preferred when utilising glucose as
fermentation substrate for biobutanol fermentation (Jones et al., 1982).
Increasing the glucose concentration was postulated to improve product yield
and productivity as more substrate would be available for fermentation. In the
preliminary study, three different concentrations of glucose viz. 6, 8 and 9 %,
w/v were investigated. No butanol could be detected when 9 %, w/v glucose
was used. This could be attributed to excessive acidification of the medium by
the acid produced from the high concentration of glucose, leading to
premature cell death (Qureshi et al., 2008). Thus for the later part of the
119
experiments, only two concentrations i.e. 6 and 8 %, w/v of glucose were
studied.
The pH of the fermentation runs was found to reduce from 5.7 - 5.9, at the
start of fermentation, to 4.2 - 4.9, at the end of fermentation. The drop in pH
was attributed to the formation of acids in the medium. As a general trend,
during ABE fermentation, vegetative cells of Cl. acetobutylicum ATCC 824
consume glucose for growth and simultaneously produce organic acids
including butyric and acetic acid (Monot et al., 1982) which lower the pH of
the fermentation medium. This increase in acidity stimulates the formation of
solvents (Dürre, 2011). The drop in pH of the fermentation medium was
greater when a higher glucose concentration (8 %, w/v) was employed. High
substrate concentration has been reported to stimulate cell growth, thus
enhancing fermentation of glucose to acetic and butyric acids (Ezeji et al.,
2003). The greater reduction in pH after fermentation with a higher
concentration of glucose was due to the formation of a larger amount of acids
during the fermentation process. The cell densities achieved when higher
glucose concentration was used were lower than corresponding cell densities
at lower glucose concentration. Furthermore, the butanol yields and
productivities were also on the lower side at a glucose concentration of 8 %,
w/v. As mentioned previously, excessive acidification of the media is harmful
to cell viability. Besides, at higher substrate concentration, the rate of acid
production could exceed the rate of induction of the solventogenic pathway
enzymes so that the cells could not switch to solventogenesis (Stim-Herndon
120
et al., 1996). This explains the low cell count and subsequent low butanol
yield obtained when higher concentration of glucose was used.
On the other hand, the lower glucose concentration of 6 %, w/v resulted in
significantly higher butanol yields (p-value <0.05) and higher productivities
(p-value <0.05), suggesting that the lower glucose concentration was more
suitable for fermentation by Cl. acetobutylicum ATCC 824 cells. The results
obtained are in agreement with the findings of other studies, which reported
that glucose concentration above 6 %, w/v adversely affected the viability of
Cl. acetobutylicum cells and consequently reduced their fermentation
efficiency (Häggström and Molin, 1980; Lin and Blaschek, 1983; Papoutsakis,
2008). Thus, glucose concentration of 6 %, w/v was used in subsequent
studies.
F.2 Influence of inocula age
Solventogenesis has been widely related with the older sporulating stages in
the stationary phase of growth of Cl. acetobutylicum (McNeil and Kristiansen,
1985). Hence, in order to reduce the overall lag phase during fermentation,
the cells in the late stationary phase were used as inoculum for fermentation.
However, it was found that the 48 h inocula took longer time to attain
maximum butanol yield compared to 24 h inocula (Table 16). This resulted in
corresponding lower butanol yields and productivities. Studies have reported
that initiation of solventogenesis requires a certain minimum concentration of
acids, produced exclusively by cells in exponential phase of growth (Qureshi
121
et al., 2008). Butanol is formed only when the acid concentration in the
medium reaches the threshold value (Schoutens et al., 1985). Since most of the
cells in the 48 h inocula were in the stationary/solventogenic, there was little
acid produced. As a result, sufficient acid concentration may not have been
achieved in these cultures to promote solvent production by the cells.
Moreover, the cell density achieved when a 48 h inocula was used was
significantly lower than that achieved with 24 h inocula (p < 0.05). Most of
the cells in the 48 h inocula existed as stationary phase cells which were
devoid of division and proliferation capabilities resulting in low cells densities
achieved. The above reasons accounted for the poor butanol yields and
productivities when 48 h inocula were utilised for fermentation. Based on the
results, it was concluded that the age of the inocula used for fermentation
should be 24 h to obtain higher butanol yields and productivities.
F.3 Influence of inocula size
Fermentation yields and productivities are usually higher when high bacterial
cell densities are used (Riesenberg and Guthke, 1999). An initial inocula size
of 10 %, v/v is normally utilised for ABE fermentation (Qureshi and Blaschek,
2001). In order to investigate the effect of increasing the initial inocula size to
15 %, v/v on the fermentation efficiency, fermentation was carried out with
inocula size of 10 %, v/v and 15 %, v/v.
It was found that increasing the inoculum size from 10 to 15 %, v/v did not
improve the fermentation efficiency of the clostridial cells (Table 16). The cell
densities obtained with 10 %, v/v initial inocula size were higher than 15 %,
122
v/v inocula size when 24 h inoculum was used. Thus age of the inoculum had
more significant effect on the cell density than size of the inoculum. This
implied that even though the initial cell count was high, most of the cells could
not survive in the presence of toxic metabolites formed during the
fermentation process.
Based on the results from this study, the conditions used in fermentation run
number F-10-6-24 produced the highest butanol concentration and
productivity of 11.2 g/L and 0.23 g/L/h respectively. Thus, these conditions
were used for the subsequent studies that involved fermentation by free Cl.
acetobutylicum ATCC 824 cells.
123
Table 16. Results from the fermentation optmisation studies of free (non-encapsulated) cells of Cl. acetobutylicum ATCC
824
Code F-10-6-24 F-10-8-24 F-10-6-48 F-10-8-48 F-15-6-24 F-15-8-24 F-15-6-48 F-15-8-48
Fermentation
time (h) 48 48 72 72 48 48 72 72
Initial pH 5.87 ± 0.12 5.79 ± 0.05 5.9 ± 0.07 5.73 ± 0.04 5.84 ± 0.03 5.87 ± 0.05 5.92 ± 0.05 5.73 ± 0.02
Final pH 4.85 ± 0.03 4.2 ± 0.11 4.7± 0.03 4.38 ± 0.06 4.89 ± 0.04 4.26 ± 0.08 4.55 ± 0.01 4.32 ± 0.07
Cell density 1.9 (±0.24)
×1010
2.6 (±0.32)
×108
1.9 (±0.14)
×106
4.42 (±0.59)
×105
3.97 (±0.17)
×107
7.03 (±0.42)
×106
6.37 (±0.19)
×106
4.49 (±0.15)
×105
Butanol yield
(g/L)
11.2 (±0.07) 7.2 (±0.09) 6.5 (±0.1) 5.7± (0.05) 8.3 (±0.12) 4.6 (±0.03) 5.32 (±0.05) 3.9 (±0.3)
Butanol
productivity
(g/L/h)
0.23 0.15 0.09 0.08 0.17 0.06 0.09 0.05
124
G. Fermentation using encapsulated spores of Cl. acetobutylicum ATCC
824
Studies were carried out to investigate the effect of various factors viz. glucose
concentration, inocula size and heat shock treatment on the fermentation
efficiency of encapsulated spores of Cl. acetobutylicum ATCC 824. In the
preliminary studies, the HST condition found suitable for maximum revival of
free (non-encapsulated) spores was 80 °C for 10 min. However, this condition
could not revive the encapsulated spores and no butanol was detected in the
fermentation medium. This could be attributed to the hindrance of heat
transfer to the encapsulated spores due to the polymeric barrier of gellan gum
microspheres. Hence, harsher HST conditions were investigated in the
optimisation studies. The optimisation studies were carried out as a series of 8
fermentation runs with various combinations of the investigated factors. The
results of all experiments are summarised in Table 17.
It was found that glucose concentration as well as inocula size did not have
any significant effect on the fermentation efficiency. The latter improved only
when the heat shock treatment (HST) at 90 °C for 10 min was employed.
Thus, HST had a major effect on performance of encapsulated cells. The HST
condition of 90 °C for 10 min consistently gave better results than HST
condition of 90 °C for 15 min with the highest butanol yield of 7.23 g /L
(Table 17). On the other, HST at 90 °C for 15 min consistently gave poor
butanol yields. It is important to mention that though exposure to high
temperature is important to revive clostridial spores, too harsh conditions may
125
be detrimental to their viability. Previous studies with non-encapsulated spores
also showed that when the HST condition was too harsh, the viable count of
revived spores was lower (section A.5). Similarly, in the case of encapsulated
spores, harsher HST conditions might have resulted in loss of cell viability and
thus lower butanol yields. The fermentation time was longer for encapsulated
spores compared to free cells resulting in comparatively lower productivities.
This could be mainly attributed to longer time required by the encapsulated
spores to revive into vegetative form to initiate butanol production. Moreover,
the polymeric barrier of the gellan gum microsphere may have resulted in
diffusion limitation which was absent in the case of free cells. For subsequent
studies, the following conditions were used for comparison of fermentation
efficiency between encapsulated spores and free cells: Glucose concentration
of 6 %, w/v, inocula size of 10 %, v/ v and HST at 90 °C for 10 min. Detailed
fermentation profiles of the encapsulated cells will be discussed in the
following sections.
126
Table 17. Results from the fermentation optmisation studies of encapsulated spores of Cl. acetobutylicum ATCC 824
Code S-10-6-
9015
S-15-6-
9015
S-10-8-
9010
S-15-6-
9010
S-15-8-
9010
S-10-6-
9010
S-15-8-
9015
S-10-8-
9015
Fermentation
time (h) 72 96 72 96 96 72 120 144
Initial pH 5.87
(±0.17)
5.79
(±0.23)
5.9
(±0.07)
5.73
(±0.15)
5.84
(±0.22)
5.87
(±0.13)
5.92
(±0.03)
5.73
(±0.24)
Final pH 4.85
(±0.24)
4.7
(±0.28)
4.65
(±0.17)
4.62
(±0.12)
4.89
(±0.26)
4.69
(±0.11)
4.45
(±0.29)
4.42
(±0.08)
Butanol yield
(g/L)
2.43
(±0.29)
1.97
(±0.19)
6.99
(±0.78)
7.09
(±1.08)
7.23
(±0.97)
7.17
(±0.85)
2.45
(±0.14)
2.34
(±0.25)
Butanol
productivity
(g/L/h)
0.03 0.02 0.097 0.098 0.075 0.099 0.02 0.016
127
H. Cell leakage from gellan gum microspheres
The spores encapsulated in the gellan gum microspheres were revived by HST
and expected to develop into vegetative cells. Thus, at a later stage of
fermentation, the encapsulated spores were referred to as encapsulated cells.
At the end of the fermentation process, a considerable number of free Cl.
acetobutylicum ATCC 824 cells were found in the fermentation medium
(Figure 21). These cells were liberated from the microspheres. This indicates
that the gellan gum matrix was porous and encapsulated cells (revived from
spores) were able to free themselves from the matrix. The resulting cell
growth inside the beads is known to depend on diffusional limitations
imparted by the polymer matrix and subsequently the porosity of the gel
matrix and later by the effect of accumulating biomass (Freeman and Lilly,
1998). Further studies were carried out to assess the extent of cell leakage,
contribution of encapsulated and liberated cells to butanol production as well
as physical stability of the microspheres.
Figure 21. Photograph of liberated Cl. acetobutylicum ATCC 824 cells
from gellan gum microspheres
Gellan gum microspheres
containing
Cl. acetobutylicum ATCC
824 cells
Liberated
Cl. acetobutylicum
ATCC 824 cells
128
H.1 Contribution by liberated cells to butanol production
The viable counts of the Cl. acetobutylicum ATCC 824 cells liberated from
the gellan gum microspheres during the course of fermentation were
determined. The viability profile of liberated cells was compared to the
fermentation profile of a combination encapsulated and free cells in Figure 22.
A significant number of viable cells liberated from the microspheres were
observed after 48 h of fermentation. The cell count increased exponentially till
72 h of fermentation followed by a plateau like stage where the growth halted.
Thereafter, the cell count dipped significantly. The viability profile of
liberated cells follows a typical growth curve of Cl. acetobutylicum ATCC 824
in RCM medium. It was seen that the butanol yield was negligible at 24 h of
fermentation. Many factors could have contributed to this observation. Firstly,
the encapsulated spores required time to revive into vegetative cells to carry
out the fermentation. Secondly, the polymeric barrier of the microsphere
slowed down the mass transfer of nutrients into the microspheres. Thirdly, due
to the aforementioned reasons, there was insignificant number of liberated
cells to contribute to butanol fermentation. After 24 h, the butanol
concentration increased gradually. Thus, as the cell number increased in the
fermentation medium, the butanol yield also increased simultaneously. The
maximum butanol yield of 8.2 g/L was obtained at 120 h of fermentation.
Based on the viability profile of liberated cells, it can be seen that at 120 h
most of the liberated cells were in the late stationary phase. As discussed
previously, butanol production occurs majorly during the stationary phase of
129
growth of the solventogenic bacterium. This explained the highest butanol
yield recorded at 120 h of fermentation. With the cell count declining after 120
h, no further increase in butanol yield was seen. This indicates conditions in
the medium were no longer amenable for cell proliferation and metabolism as
the concentration of toxic butanol reached its maximum concentration. Based
on the results from this study, it could be concluded that once the spores
revived into vegetative cells inside the microspheres, they liberated and
proliferated in the fermentation medium. The liberated cells then followed a
typical growth profile of Cl. acetobutylicum ATCC 824 cells in the
fermentation medium and contributed to butanol production.
Figure 22. Viability profiles of Cl. acetobutylicum ATCC 824 cells
liberated from gellan gum microspheres and fermentation profile of
encapsulated and liberated cells
0
1
2
3
4
5
6
7
8
9
10
0
20
40
60
80
100
120
140
0 20 40 60 80 100 120 140 160
Bu
tan
ol
yie
ld (
g/L
)
Via
ble
cou
nt
(mil
lion
s/m
L)
Fermentation time (h)
Viable count Butanol yield
130
In order to determine whether butanol production was carried out solely by the
liberated cells, fermentation study by free cells corresponding to the number of
liberated cells (at a time point where maximum butanol was attained) was
carried out. Figure 23 shows the fermentation and viability profiles of
equivalent number of free cells. The viable count of free cells increased
exponentially from 0 to 24 h with maximum cell count of 6 X 108
cells/mL.
There was a very short stationary phase at 48 h followed by a steep decline in
cell count. After 120 h of fermentation, there were no viable cells present in
the fermentation medium. Maximum butanol yield of 4.44 g/L was obtained at
48 h of fermentation corresponding to stationary phase of bacterial growth
curve. No significant improvement in butanol yield was seen thereafter. Thus
even though higher cell densities were obtained at the initial stage of
fermentation, the conditions formed in the medium later were harsh for the
cells. Butanol concentration as low as 4.44 g/L was found to be toxic to the
free cells. This value is significantly lower than that achieved with
encapsulated cells. The difference in butanol yields between encapsulated and
liberated cells could be attributed to the former. Thus butanol production
seemed to be a combined effect of both encapsulated and liberated cells.
The gellan gum microspheres were recovered at various stages of fermentation
for examination under the microscope (Figure 25). No free cells were
observed after 24 h of fermentation. However, increasing number of cells was
liberated from the microspheres thereafter. At the beginning, the cells
appeared to be uniformly dispersed throughout the microspheres. As the
131
fermentation proceeded, a number of cells were seen at the periphery of the
microspheres (Figure 24). It could be postulated that due to mass transfer
limitations presented by the polymeric barrier of the microspheres, cells
concentrated at the boundary of the microspheres. As the cell density
increased due to cell proliferation, the cells were eventually liberated from the
microspheres. It should be mentioned that Cl. acetobutylicum is a motile
bacterium. Thus it could have led to an enhancement in the liberation of the
cells from the microspheres. The increase in the liberation of cells from the
microspheres also showed a corresponding increase in butanol concentration
in the fermentation medium. The liberated cells were no longer facing mass
transfer limitation by gellan gum matrix. Thus, the cells could carry out
butanol production at a faster rate now. At the same time, the free cells were
exposed to the toxicity of the accumulated metabolites and products in the
fermentation medium. This led to drop in viable count and consequent drop in
butanol concentration. Nevertheless, the gellan gum microspheres remained
intact and spherical at the end of the fermentation process withstanding the
harsh conditions including drop in pH value of the fermentation medium.
Thus, the gellan gum matrix was strong and stable and could be used for
butanol fermentation.
Collectively, the results clearly showed that the both liberated and
encapsulated Cl. acetobutylicum ATCC 824 cells played a role in the
production. The gellan gum microspheres provided suitable microenvironment
for the encapsulated cells to grow and multiply. As the cell density increased,
132
the cells migrated towards the periphery and were later liberated at
approximately after 48 h of fermentation, implying that gellan gum matrix was
sufficiently porous and allowed liberation of cells during the fermentation
process. The liberated cells were no longer faced with diffusion limitations of
polymeric membrane of the microspheres and therefore proliferated rapidly in
the fermentation medium. This led to increase in butanol yield. The viable cell
count in the fermentation medium was consistently high till 120 h of
fermentation as new cells were liberated from the microspheres (Figure 22).
Thus, besides contributing to the butanol production, the microspheres served
as nurseries to generate free Cl. acetobutylicum ATCC 824 cells into the
medium to carry out fermentation. The fermentation activity of the liberated
cells was later inhibited by the high butanol concentration in the fermentation
medium.
133
Figure 23. Viability and fermentation profiles of free Cl. acetobutylicum
ATCC 824 cells (equivalent to the number of liberated cells from the
microspheres during the course of fermentation)
Figure 24. Photographs of microspheres with Cl. acetobutylicum ATCC
824 cells at the periphery of gellan gum microspheres
0
0.5
1
1.5
2
2.5
3
3.5
4
4.5
5
0
100
200
300
400
500
600
700
0 50 100 150 200
Bu
tan
ol
yie
ld (
g/L
)
Via
ble
co
un
t (m
illi
on
s/m
L)
Fermentation time (h)
Viable count Butanol yield
Cl. acetobutylicum ATCC 824 cells
at the periphery of gellan gum
microspheres
134
Figure 25. Photographs of gellan gum microspheres recovered from fermentation after (a) 24 h, (b) 48 h, (c) 72 h, (d) 96 h,
(e) 120 h and (f) 144 h of fermentation
(b)
(e) (d) (f)
(c) (a)
135
I. Reusability of free (non-encapsulated) vegetative cells/spores and
encapsulated spores of Cl. acetobutylicum ATCC 824
In this study, free (non-encapsulated) and encapsulated cells of Cl.
acetobutylicum ATCC 824 were recovered at the end of fermentation cycle
and reused in a fresh fermentation medium. Comparison of the fermentation
efficiency between free and encapsulated cells of Cl. acetobutylicum ATCC
824 in successive fermentation cycles was made.
The fermentation efficiency of free (non-encapsulated) cells was high during
the first fermentation cycle (Figure 26a). This could be due to the presence of
highly active culture as well as sufficient nutrients in the fermentation
medium. Within the fermentation cycle, the butanol yield steadily increased
with time. The maximum butanol yield of 9.79 g/L was achieved after 72 h of
fermentation. Thereafter, there was no significant increase in butanol yield.
This implied that the conditions in the fermentation medium could no longer
support cell growth resulting in loss of activity of the cells. Compared to free
cells, the maximum butanol concentration produced by encapsulated cells
during the first fermentation cycle was 7.66 g/L after 120 h of fermentation.
The fermentation curve of the encapsulated cells followed a trend of an initial
lag phase, exponential phase and stationary phase. As discussed before, the lag
phase was due to the time required for the spores to revive to vegetative cells
as well as acclimatisation of the cells to the microenvironment of the gellan
gum microspheres. This also explained the longer fermentation time required
by the encapsulated cells to achieve maximum butanol concentration. In
136
addition, impairment of mass transfer by gellan gum matrix might have also
slowed down the growth and proliferation of the encapsulated cells.
When the free (non-encapsulated) Cl. acetobutylicum ATCC 824 cells were
recovered and reused in the second fermentation cycle, a significant drop in
butanol concentration was observed from 9.79 g/L in first cycle to 2.9 g/L in
second cycle (p- value <0.05) (Figure 26b). This could be due to exposure of
the cells to toxic metabolites such as acids and solvents formed during the first
fermentation cycle, thereby affecting their viability and fermentation
efficiency. On the other hand, the maximum butanol concentration produced
by encapsulated cells (6.89 g/L) was markedly higher than that produced by
free cells (2.9 g/L) in the second fermentation cycle (p- value <0.05) .
The free cells could not be used after second fermentation cycle. On the other
hand, the encapsulated cells consistently gave high butanol yields in third
cycle (Figure 26c). Maximum butanol yield of 6.99 g/L was observed after 72
h of fermentation in the third cycle. A slight reduction in butanol production
by encapsulated cells was observed in the fourth cycle (Figure 26d). The
decrease might be associated with the weakening of the gellan gum matrix,
which would compromise its protective function. In addition, conversion of
the viable cells to spores might have further reduced the fermentation
efficiency. These effects were more pronounced when the cells were reused in
the fifth fermentation cycle (Figure 26e). The cells could no longer be used for
further fermentation cycles.
137
Despite the decrease, the reusability of the encapsulated cells and fermentation
efficiency was significantly higher compared to free cells (p-value <0.05).
These observations clearly showed that the encapsulated cells exhibited higher
fermentation efficiency and tolerance to ethanol than the free cells. This
suggests that the gellan gum matrix improved the fermentation efficiency of
the free cells.
From the results, it could be inferred that the encapsulated cells would exhibit
better survival rate than the free cells when used in repeated batch
fermentation. Apart from the lag phase in the first fermentation cycle, the
performance of encapsulated cells was markedly better than non-encapsulated
free cells. The microspheres could provide suitable conditions for the cells to
grow. The cells from the microspheres were later liberated and contributed
significantly to butanol production. However, a significant drop in
fermentation efficiency in the fifth fermentation cycle was observed. As Cl.
acetobutylicum cells grow and metabolise fermentation substrate, the cells are
converted into various morphological forms such as vegetative cells,
forespores, spores and dead cells during the course of fermentation. There is a
wide variation in the number of each type of morphological structure formed.
Viable count after fifth cycle revealed dead cells non-viable spores. Based on
the results from the above studies, it could be assumed that initially the
encapsulated cells were able to grow and proliferate well. Overtime, the
number of non-viable cells in the form of dead cells and spores might have
increased. Thus, when the microspheres were introduced into fresh
138
fermentation medium with more substrate, fewer viable cells present in the
microspheres could not utilise the substrate efficiently resulting in low butanol
yield.
The gellan gum microspheres were recovered at the end of each fermentation
cycle to investigate the physical stability of the microspheres (Figure 27).
When observed under the light microscope, the microspheres were found to be
intact throughout the five fermentation cycles. Thus, the gellan gum
microspheres could withstand the low pH as well as presence of conditions in
the fermentation medium.
In this study, significant cell leakage from gellan gum microspheres was
observed. It was found that these cells also contributed to butanol production
besides the encapsulated cells. Thus, the microspheres served as cell nurseries
supplying free vegetative cells into the fermentation medium for butanol
production. These microspheres could be reused in subsequent five batch
fermentation cycles unlike two cycles for free cells. Thus, overall, cell leakage
was deemed to be a desirable trait. A similar study was done by Tan et al.,
2012, wherein leakage of yeast cells was observed from gellan gum
microspheres. The microspheres could be reused fifteen times for successive
batch fermentation with good fermentation efficiency.
139
Figure 26. Plots of fermentation profile of free cells vs. encapsulated cells
in (a) first fermentation cycle
0
2
4
6
8
10
12
0 20 40 60 80 100 120 140 160
Bu
tan
ol y
ield
(g
/L)
Time (h)
Free cells Encapsulated cells
e
(a)
140
Figure 26. Plots of fermentation efficiency of free cells vs. encapsulated cells
in (b) second and (c) third fermentation cycles
0
2
4
6
8
10
12
0 50 100 150 200
Bu
tan
ol y
ield
(g
/L)
Time (h)
Free cells Encapsulated cells
0
2
4
6
8
10
12
0 50 100 150 200
Bu
tan
oly
ield
(g
/L)
Time (h)
Encapsulated cells
(b)
(c)
141
Figure 26. Plots of fermentation efficiency of free cells vs. encapsulated cells
in (d) fourth and (e) fifth fermentation cycles
0
2
4
6
8
10
12
0 50 100 150 200
Bu
tan
ol
yie
ld (
g/L
)
Time (h)
Encapsulated cells
0
2
4
6
8
10
12
0 50 100 150 200
Bu
tan
ol
yie
ld (
g/L
)
Time (h)
Encapsulated cells(e)
(d)
e
142
Figure 27. Photographs of gellan gum microspheres recovered from
fermentation medium after (a) 1 cycle, (b) 2 cycles, (c) 3 cycles, (d) 4
cycles and (e) 5 cycles of fermentation
(e)
(a) (b)
(c) (d)
143
PART TWO
A. Potential of Samanea saman leaf litter as a source of fermentable
sugars for biobutanol production
As the feedstock or substrate for biobutanol fermentation amounts to more
than 60 % of the production costs, efforts have been made to replace the
conventional food crop-based substrates such as corn starch and molasses with
cheaper alternatives, preferably waste lignocellulosic material (Nigam and
Singh, 2011; Qureshi et al., 2007). Samanea saman tree, also known as rain
tree, commonly planted along local roadsides for the purpose of shade,
generates huge quantities of leaf litter daily. This constitutes a type of
municipal waste which needs removal and disposal by the local authorities.
However, the common approach of incinerating the leaf litter often results in
adverse environmental impacts (Ghimire et al., 2012). While the potential
contribution of other biomass sources, such as agricultural residues and
forestry wastes have been increasingly studied (Champagne, 2007; Karimi et
al., 2006; Qureshi et al., 2008; Zhu et al., 2010), the biomass potential of S.
saman leaf litter has yet to be investigated.
In this part of the project, the possibility of using the leaf litter from S. saman
tree as a potential lignocellulosic substrate for biobutanol production was
explored. The total amount of fermentable sugars present in S. saman leaf
litter was first determined. This was followed by the investigation of various
pretreatment methods and their effects on the sugar recovery before and after
144
enzymatic hydrolysis. Once a suitable pretreatment method was selected,
optimisation studies were carried out using response surface methodology.
Subsequently, different detoxification strategies were studied to make the
pretreated leaf litter amenable for bacterial fermentation. Finally, a
comparative study between the fermentation efficiency of the pretreated leaf
litter by free and encapsulated cells of Cl. acetobutylicum ATCC 824 was
carried out.
A.1 Recovery of total fermentable sugars from S. saman leaves
Fermentable sugars can be obtained from structural carbohydrates present in
the lignocellulosic biomass. Structural carbohydrates are composed of
polymeric carbohydrates, namely cellulose and hemicellulose, which are
bound in the matrix of the biomass (Fatih Demirbas et al., 2011; Hendriks and
Zeeman, 2009). In order to determine the total fermentable sugars in S. saman
leaf litter, a two-step acid hydrolysis procedure was used as per the guidelines
given in National renewable energy laboratory (NREL) (Sluiter et al., 2008).
During hydrolysis, the polymeric structural carbohydrates were hydrolysed
into the monomeric sugars, which were then measured by the DNS method.
The leaf litter was found to contain 48.66 g of fermentable sugars per 100 g of
substrate. This corresponds to 48.66 %, w/w on a dry matter basis, which is
comparable to the value (48.50 %, w/w), reported in literature (Chanda et al.,
1993; Datt et al., 2008). Based on the availability of this high fermentable
sugar content, it was concluded that the S. saman leaf litter is a promising
source of lignocellulosic substrate for biofuel production.
145
It is possible to completely recover the available fermentable sugars from the
leaf litter using the procedure described by the NREL. However, the procedure
was not recommended because of the use of very strong acid (72 %, w/w).
Besides being hazardous, it may cause degradation of the sugars into harmful
products which can be detrimental to the microbial cells employed in the
biofuel production process. Thus, this study adopted a less harsh method for
the recovery of fermentable sugars from the leaf litter.
A.2 Determination of filter paper activity of Accellerase® 1500
The cellulolytic enzyme used in this study was Accellerase® 1500. In order to
determine its cellulolytic activity, filter paper assay was carried out. Assay of
filter paper activity (FPA) is recommended by the International Union of Pure
and Applied Chemistry (IUPAC) for the assessment of cellulase activity
because of its simplicity and readily duplicated conditions (Ghose, 1987). This
assay is based on the degree of conversion of a fixed amount of filter paper to
glucose by the test enzyme. The cellulase activity is described in terms of
“filter-paper units” (FPU) where one FPU is defined as the amount of enzyme
that releases 1 mole of glucose per minute during the hydrolysis reaction. In
this study, 50 mg of filter paper were subjected to known concentrations of
Accellerase® 1500 for a fixed reaction time of 60 min. The amount of glucose
released by each of the enzyme concentration was determined by the DNS
assay and a calibration curve. The latter was first constructed by plotting
known concentrations of glucose subjected to DNS assay against their
absorbance at 540 nm (Figure 28). Based on the plot, a linear equation was
146
derived to compute the glucose concentrations released by the aforementioned
enzyme concentrations. A plot of concentration of glucose liberated against
enzyme concentration was then constructed (Figure 29)
The enzyme concentration required to release 2.0 mg/mL of glucose was
extrapolated from the graph and found to be the 0.0081. Substituting the latter
value into equation 7 (section B.10), the activity of Accellerase®1500 was
found to be 45.68 FPU.
Figure 28. Calibration curve of glucose
y = 0.182x + 0.093
R² = 0.998
0
0.5
1
1.5
2
2.5
0 2 4 6 8 10 12
Ab
sorb
an
ce a
t 5
40
nm
Glucose concentration (mg/mL)
147
Figure 29. Plot of enzyme concentration vs. glucose concentration
A.3 Pretreatment of S. saman leaf litter
The recovery of fermentable sugars from any lignocellulosic biomass is
initiated by the selection of a suitable pretreatment of the biomass followed by
enzymatic hydrolysis. Only a small number of pretreatment methods have
been reported as being potentially cost-effective thus far. These include
milling, steam explosion, liquid hot water and dilute acid coupled with heat
treatment (Cadoche and López, 1989; Cao et al., 2012; De Bari et al., 2013;
El-Zawawy et al., 2011; Fox et al., 1989; Lu et al., 2012; Rohowsky et al.,
2013; Zhang et al., 2013; Zhang et al., 2012) An ideal pretreatment method
should be able to obtain high sugar recovery, minimize sugar loss, produce no
or very little toxic by-products and should be cheap and easy to scale-up.
y = 0.013x - 0.018
R² = 0.974
0
0.01
0.02
0.03
0.04
0.05
0.06
0 1 2 3 4 5 6
En
zym
e co
nce
ntr
ati
on
(m
g/m
L)
Glucose concentration (mg/mL)
148
A.3.1 Milling
Milling of S. saman leaf litter was investigated as a mechanical pretreatment
method. Apart from disrupting the lignocellulosic structure and increasing the
specific surface area for enzymatic action, milling also make handling of
biomass easier. The extent of size reduction required depends on the type of
biomass (Schell and Harwood, 1994). Particle size of 1 - 2 mm is usually
preferred for herbaceous biomass feedstock such as leaves (Belovsevic, 2010).
The hammer mill, with the cutter blade, was initially employed in this study to
shred the leaf litter into smaller fragments (Figure 30a). However, despite
allowing prolonged milling, the particle size remained rather coarse in the
range of 3-10 mm (Figure 30b). Hence, a second mill, the disintegrator mill
was employed. The leaf litter was further reduced to particles in the size range
of 0.3 - 1 mm (Figure 30c). It was possible to use the disintegrator mill alone,
albeit over a longer milling period, to achieve the desired size reduction of the
leaves. Hence, the leaf litter was milled using a combination of the two mills.
Following milling, the leaves were subjected to enzymatic hydrolysis to
evaluate the efficiency of the pretreatment in improving the accessibility of
enzyme.
A.3.1.1 Enzymatic hydrolysis of milled leaves
A preliminary enzymatic hydrolysis of the milled leaves was carried out to test
the efficiency of milling as a pretreatment method. From the literature, low to
high doses of Accellerase® 1500 were identified. These doses, which ranged
from 7 to 35 FPU/g substrate, were utilised for hydrolysis of the milled leaves
149
over 48 hours. A positive linear relationship was observed between the
enzyme dose and percent recovery of fermentable sugars (Figure 31). The
level of sugar recovered was approximately 7 %, w/w after 48 h of incubation
with a high dose of enzyme (35 FPU/g of substrate). This implied that
although the milling process had successfully reduced the particle size, it was
ineffective in breaking down the lignocellulosic structure of the leaves to
facilitate penetration of the enzyme to the holocellulosic components.
Extrapolation of the relationship between enzyme dose and sugar recovery
(Figure 31) suggests that more than 800 FPU/g of Accellerase® 1500 would
be needed for complete recovery of the fermentable sugars. However, use of
such a high dose of enzyme is not recommended as it will increase the cost of
the process considerably. Clearly, milling of S. saman leaf litter alone was
inadequate as a pretreatment process. Hence, further pretreatment of the milled
leaves was necessary prior to the enzymatic hydrolysis step.
150
Figure 30. Leaf litter of S.saman: (a) before milling, (b) after using
hammer mill, (c) followed by disintegrator mill
(a)
(b)
(c)
151
Figure 31. Relationship between sugar recovery and Accellerase® 1500
dose
A.3.2 Hydrothermal pretreatment of S. saman leaf litter
In this method, the lignocellulosic substrate was subjected to water in hot,
compressed state throughout the procedure unlike in steam explosion wherein
the pressure is suddenly released to cause decompression of the lignocellulosic
substrate. This resulted in an increase in mean pore size of the substrate as
well as other structural alterations thereby enhancing the enzymatic
accessibility. It is advantageous over other methods as it is non-toxic and
inexpensive.
It was seen that hydrothermal treatment could not recover significant amount
of sugars (~2 %) from the milled leaf litter (Figure 32). Further, treatment of
y = 0.12x + 2.37
R² = 0.988
0
1
2
3
4
5
6
7
8
0 10 20 30 40
Su
ga
r re
cov
ery
(%
)
Dose of enzyme
(FPU/g of substrate)
152
the hydrothermal pretreated leaf litter with the cellulolytic enzyme at a
moderate dose of 20 FPU/g substrate led to the recovery of significantly
higher sugar content (~12 %) than the control (~6 %) (p-value < 0.05). The
control consisted of milled leaf litter without hydrothermal treatment. This
implied that milling-hydrothermal treatment was a better pretreatment method
than milling alone. Nonetheless, the highest sugar recovery obtained with
hydrothermal treatment was still on the lower side (~12 %). It was thus
concluded that other pretreatment methods need to be investigated as an
alternative to hydrothermal pretreatment.
Figure 32. Effect of hydrothermal pretreatment on sugar recovery from
S. saman leaf litter before and after enzyme addition
A.3.3 Dilute acid coupled with heat treatment
A preliminary dilute acid couple with heat treatment of the milled S. saman
leaf litter was carried out without the use of Accellerase® 1500. It has been
0
2
4
6
8
10
12
14
30 60 90
Su
gar
reco
ver
y (
%)
Treatment time (min)
Before enzyme addition After enzyme addition
153
reported that moderate temperatures (100-150 °C) were adequate for
hemicellulose hydrolysis, with little sugar decomposition while temperatures
above 160 °C caused considerable sugar decomposition despite being more
favourable for hydrolysis (Lenihan et al., 2010). Hence, a treatment
temperature of 121 °C was selected for this study. An acid concentration of
1.0 %, w/w was used with treatment times of 30, 60 or 90 min. An appreciable
amount of fermentable sugars was obtained after the dilute acid coupled with
heat treatment, indicating significant disintegration of lignin and hydrolysis of
hemicellulose/cellulose to produce fermentable sugars (Figure 33). Compared
to enzyme hydrolysis of the milled leaves with a high dose of Accellerase®
1500 (~6 % sugar recovery) and hydrothermal pretreatment of the milled
leaves (~12 % sugar recovery), dilute acid coupled with heat treatment of the
milled leaves for 30 min produced markedly higher (~60 %) sugar recovery.
In general, leaves have a high hemicellulose content (80 - 85 % on dry basis)
and low lignin content compared to other lignocellulosic substrates (Sun and
Cheng, 2002). The acid increases porosity of the substrate by solubilization of
hemicellulose, making the substrate more accessible to cellulolytic enzymes
(Esteghlalian et al., 1997; Zheng et al., 2009a). This could have explained the
high sugar recoveries using dilute acid coupled with heat treatment of S.
saman leaf litter.
The operating condition for the acid treatment must be tailored to the specific
chemical and structural composition of the various sources of biomass so as to
maximize the sugar yield and recovery (Hendriks and Zeeman, 2009). The
154
results indicated that the recovery of fermentable sugars increased as the
treatment time increased. Thus, treatment time played a crucial role in the
hydrolysis process and needs to be further investigated. Besides the treatment
time, the type and concentration of acid also affect the process (Castro et al.,
2011; Gil et al., 2010; Xu and Hanna, 2010). Hence, further optimisation study
for dilute acid coupled with heat treatment of S. saman leaf litter was carried
out with respect to acid concentration and treatment time.
Figure 33. Sugar recovery from milled S. saman leaf litter subjected to
1.0 %, w/w acid for different treatment times
0
10
20
30
40
50
60
70
80
30 60 90
Su
gar
reco
ver
y (
%)
Treatment time (min)
155
A.3.3.1 Optimisation of dilute acid coupled with heat treatment of S.
saman leaf litter
The optmisation study to assess the effect of acid concentration and treatment
time on sugar yield and sugar recovery was performed using response surface
methodology (RSM). It is a statistical technique useful for the modelling and
analysis of experiments in which one or more responses of interest are
influenced by several variables and the aim is to optimize this response
(Koyamada et al., 2004). Response surface plots were created by assessing the
effect of the two variable factors on the response. Table 18 shows the
experimental design along with the values of the respective responses.
Table 18. Experimental design used for the optimisation of dilute acid
coupled with heat treatment along with the values of the response
variables
Run A
B
Sugar yield
(g/L)
Sugar recovery
(%)
1 2.75 60 18.15 85.85
2 2.75 30 15.55 81.79
3 5 90 12.99 50.36
4 0.5 60 11.23 47.5
5 0.5 30 8.87 34.4
6 5 30 16.34 67.22
7 2.75 90 17.21 83.32
8 5 60 15.12 65.02
9 0.5 90 12.87 49.12
A: Acid concentration (%, w/w) B: Treatment time (min)
156
The following quadratic models were obtained to represent the relationships
between treatment conditions and yield or recovery of fermentable sugars
from the milled leaf litter based on the results shown in Tables 18 and 19.
Only the significant terms were included in each model.
Sugar yield = 17.54 + 1.91A + 0.39B – 4.07A2 – 0.86B
2 – 1.84AB
Sugar recovery = 87.05 + 8.61A – 0.12B – 7.87AB – 31.41A2 – 5.1B
2
where A: acid concentration (%, w/w) and B: treatment time (min)
The yield of fermentable sugars from the acid hydrolysis of the milled S.
saman leaf litter was found to depend on acid concentration (p-value <0.05)
while treatment time had insignificant effect (p-value >0.05). The above
effects can also be observed in the surface plots (Figure 34a). It was observed
within the experimental range that the sugar yield increased with increase in
acid concentration till approximately 4.0 %, w/w. Any further increase in acid
concentration led to drop in sugar yield. The drop in sugar yield could be
attributed to the degradation of sugars due to the harsher conditions produced
by a higher concentration of acid. It was also found that at low acid
concentration, increase in treatment time led to subsequent increase in sugar
yield whereas at higher acid concentration, increase in treatment time led to
drop in sugar yield. This implied that a combination of high acid and treatment
time was too harsh and could have led to degradation of the sugars. In
addition, it also indicated a greater impact of acid concentration on sugar yield
over treatment time.
157
Table 19. ANOVA table for yield and recovery of fermentable sugars in dilute acid coupled with heat treatment
Sugar yield Sugar recovery
Source Sum of
Squares
Mean
Square
F value p value
Sum of
Squares
Mean
Square
F value p value
Model 70.92 14.18 29.92 0.0092 2717.47 543.49 287.13 0.0003
A 21.97 21.97 46.33 0.0065 445.14 445.14 235.17 0.0006
B 0.89 0.89 1.88 0.2643 0.089 0.089 0.047 0.8424
AB 13.51 13.51 28.49 0.0129 247.43 247.43 130.72 0.0014
A2 33.08 33.08 69.77 0.0036 1972.76 1972.76 1042.20 < 0.0001
B2 1.48 1.48 3.13 0.1749 52.05 52.05 27.50 0.0135
Residual 1.42 0.47 5.68 1.89
Lack of fit 0.36 0.775 1.44 0.72
158
The sugar recovery from the leaf litter was found to be significantly affected
by acid concentration (p-value <0.05). The sugar recovery increased with
increase in acid concentration till approximately 3 %, w/w after which there
was a steady decline observed (Figure 34b). The trends for the two responses
were found to be similar owing to the fact that they were correlated. Thus
increase in sugar yield led to corresponding increase in sugar recovery and
vice versa.
Further statistical analysis of the results is shown in Table 19. The two models
for sugar yield and sugar recovery showed insignificant lack of fit. The F
value for model of the sugar yield was 29.92 and the p value was 0.0092,
further indicating that the model was significant and accurate. The F and p
values of the model for sugar recovery were 287.13 and 0.0003 respectively,
also indicating that the model was significant and accurate. In addition, the
correlation coefficients (R2) of the models (0.9803 and 0.9979, respectively)
indicate that the models explained the responses well.
159
Figure 34. Surface plots for effects of treatment time and acid
concentration on (a) sugar yield and (b) sugar recovery
(a)
(b)
160
Figure 35. Contour plot for optimisation of the dilute acid coupled with
heat treatment of milled S. saman leaf litter
Clearly, the optimal choice of treatment time and acid concentration is
important in order to obtain maximal sugar yield and sugar recovery. The
optimisation was performed using graphical approach in which equal
importance was given to the yield and recovery of fermentable sugars (Figure
35), the optimal treatment parameters were 2.80 %, w/w of acid at a treatment
time of 58.92 min. Using this treatment condition, the predicted sugar yield
was 17.77 g/l (corresponding to approximately 41 g of reducing sugars per
100 g of leaf litter) and the sugar recovery was 87.35 % with a desirability
value of 0.979.
161
A.3.3.2 Enzymatic hydrolysis of milled leaves pretreated with dilute acid
coupled with heat
In the earlier study, less than 7 % of the available fermentable sugars were
recovered from the milled leaf litter despite using a high enzyme dose of 35
FPU/g substrate over 48 h. Further hydrothermal pretreatment increased sugar
recovery but it was still unsatisfactory. Hence, dilute acid coupled with heat
treatment was explored and found to enable approximately 86 % sugar
recovery.
Further studies were carried out to evaluate the yield and recovery of sugars
after pretreatment of the milled leaf litter with different concentrations of acid
and treatment time prior to enzymatic hydrolysis. A comparison of the total
sugar recovery obtained is given in Figure 36a. The recovery of sugars
obtained from the different dilute acid coupled with heat treatment conditions
ranged from 34 % to 86 %, indicating that substantial quantities of the sugars
were recovered by dilute acid coupled with heat treatment. It is important to
note that high concentration of acid should be avoided as it resulted in lower
sugar recovery. Moreover, complete sugar recovery could not be achieved
with dilute acid coupled with heat treatment alone and further treatment with
enzyme was needed. The latter could recover an additional 8 % to 14 %, w/w
of the sugar content of the leaves (Figure 36b). Most of the recoverable sugar
content of the leaves could be liberated by pretreatment with 2.75 %, w/w
acid, followed by enzymatic hydrolysis. It could thus be inferred that the
enzymatic susceptibility of the substrate was greatly enhanced after dilute acid
162
coupled with heat treatment. This improvement in enzymatic susceptibility
may be attributed to the degradation of lignin and/or hemicellulose by the acid
treatment. As seen from Figure 36b, higher acid concentration with treatment
time of 30 min resulted in higher sugar recovery by the corresponding
enzymatic hydrolysis. However, when the treatment time was increased to 60
and 90 min, it was observed that the increase in acid concentration may not
necessarily lead to increase in sugar recovery. For instance, at a treatment time
of 60 min, increasing acid concentration form 0.5 to 2.75 %, w/w led to
improvement in sugar recovery. But further increase in acid concentration led
to significant drop in the amount of recovered sugar. Similar trend was
observed at longer treatment times of 90 min. This could be attributed to the
harmful effects of harsher conditions, affecting amount of useful substrate
available for enzymatic hydrolysis. The degraded compounds might also have
inhibited the action of the cellulolytic enzyme. Highest total sugar recovery
was achieved when the leaves were treated with 2.75 %, w/w acid for 60 min.
In the earlier part of this study using response surface analysis, similar
conditions were found to be optimum for dilute acid coupled with heat
treatment.
163
Figure 36. Comparison of (a) total sugar recovery from S. saman leaf
litter after both dilute acid coupled with heat treatment and enzyme
hydrolysis (b) total sugar recovery due to enzymatic hydrolysis alone after
different dilute acid coupled with heat treatment conditions
0
20
40
60
80
100
120
30 mins 60 mins 90 mins
Su
ga
r re
cov
ery
(%
)
Treatment time
0.5 %, w/w 2.75 %, w/w
(a)
0
2
4
6
8
10
12
14
16
30 min 60 mins 90 mins
Su
gar
reco
ver
y (
%)
Treatment time
0.5 % w/w 2.75 % w/w 5.0 % w/w
Enzyme hydrolysis 5.0 %, w/w
164
A.4 Detoxification of acid hydrolysate of S. saman leaf litter
The processing conditions of high temperature and pressure used for dilute
acid coupled with heat treatment of lignocellulosic biomass often results in the
formation of many by-products. Examples of acid hydrolysis by-products
include furfural, hydroxymethylfurfural, acetic acid and succinic acid
(Purwadi et al., 2004; Talebnia and Taherzadeh, 2006). These by-products are
known to have severe inhibitory effects on fermenting bacterium in the
subsequent fermentation stage resulting in low yields and productivities
(Mussatto and Roberto, 2004; Palmqvist and Hahn-Hägerdal, 2000). Factors
affecting the type and concentration of the toxic compounds formed include
type of lignocellulosic substrate, acid concentration, treatment time and
temperature (Mussatto and Roberto, 2004). In addition, the hydrolysate was
highly acidic due to the use of acid (pH range 1-2). Therefore, neutralisation
of the hydrolysate is an unavoidable step before its use for fermentation.
Alkalis, such as calcium hydroxide or sodium hydroxide, are usually used for
neutralisation of the hydrolysates to pH 6.0-7.0, resulting in precipitation of
toxic compounds such as furfurals and phenols (Alriksson et al., 2006;
Martinez et al., 2001). Both sodium hydroxide and calcium hydroxide were
used in this study for neutralisation as well as chemical detoxification of the
acid hydrolysate of S. saman.
The acid hydrolysate of S. saman leaf litter was separated from the solids by
filtration in vacuo. The clear filtrate was then subjected to different
neutralisation-detoxification methods: treatment of the acid hydrolysate with
165
sodium hydroxide (A), treatment of the acid hydrolysate with sodium
hydroxide and high temperature (B), treatment of the acid hydrolysate with
calcium hydroxide (overliming) (C), treatment of the acid hydrolysate with
calcium hydroxide (overliming) and high temperature (D). The pH of the
resultant liquid was in the range of 6.6 to 7.0. It was then sterilised by
filtration before it was employed for fermentation by Cl. acetobutylicum
ATCC 824 cells over a period of 144 h. It should be mentioned that the sugar
content obtained after the hydrolysate was subjected to different detoxification
procedures was constant implying that the sugars were not degraded in the
presence of extraneous chemicals or heat. The amounts of butanol produced
are shown in Figure 37.
Figure 37. Butanol yield achieved from the acid hydrolysate of S. saman
leaf litter subjected to different detoxification methods
0
1
2
3
4
5
6
7
8
24 48 72 96 120 144
Bu
tan
ol yie
ld (
g/L
)
Fermentation time (h)
Treatment A Treatment B Treatment C Treatment D
166
Highest butanol concentration was achieved with the hydrolysate subjected to
treatment A i.e. neutralization with sodium hydroxide. This indicates that
simple neutralization of the acid hydrolysate with sodium hydroxide was the
most effective method in improving the fermentability of the hydrolysate.
Highest butanol yield was achieved after 48 h of fermentation with no
significant increase thereafter.
Further heating of the hydrolysate obtained from treatment A did not improve
the fermentability of the hydrolysate. In fact, there was a significant drop in
butanol concentration as can be seen from Figure 37. Heating was employed
as a physical detoxification method for reducing the contents of volatile toxic
compounds, such as acetic acid, furfural and vanillin, found in the hydrolysate
(Converti et al., 2000). However, heating also moderately increased the
concentration of non-volatile toxic compounds due to loss of water through
evaporation, thus affecting the fermenting cells and the degree of fermentation
adversely (Parajó et al., 1998). Reduction in microbial conversion of xylose in
present in rice straw hydrolysate to xylitol was reported when the hydrolysate
was subjected to vacuum-evaporation (Silva and Roberto, 1999). The same
authors pointed out that the production of xylitol was drastically hindered by
the increase in concentration of non-volatile compounds, which were toxic to
the microorganisms and strongly interfered with fermentation. The above
findings apparently accounted for the drop in butanol produced from the acid
hydrolysate subjected to heat.
167
The other chemical detoxification method used to enhance fermentability of
the acid hydrolysate of S. saman leaf litter was overliming. It involved the use
of calcium hydroxide to increase the pH of the hydrolysate to 10, followed by
neutralisation with sulphuric acid. The overliming method is known to
precipitate toxic compounds in the acid hydrolysate, thereby reducing its
degree of toxicity. Compared to treatment A, fermentability of the hydrolysate
subjected to overliming was poorer (Figure 37). Heating the hydrolysate
further reduced the fermentability of the hydrolysate. Since there was no
significant improvement in fermentability of the hydrolysate subjected to
overliming, it could be inferred that the hydrolysate did not contain
compounds which could be neutralized/ precipitated with calcium hydroxide.
Heating further increased the concentration of non-volatile components,
resulting in a significant drop in concentration of butanol produced.
A.5 Fermentation of detoxified leaf hydrolysate by free (non-
encapsulated) and encapsulated cells of Cl. acetobutylicum ATCC 824
The major drawback of the utilisation of S. saman leaf litter is the relatively
low content of fermentable sugars, which results, even after successful
pretreatment and hydrolysis, a rather low butanol concentration in the
fermentation medium. The maximum butanol concentration (6.8 g/L) achieved
in the fermentation studies of acid hydrolysate of S. saman leaf litter by Cl.
acetobutylicum ATCC 824 cells was on the lower side in spite of the
detoxification of the hydrolysate. It may have been possible that the
detoxification procedure could not completely remove the toxic compounds
168
from the acid hydrolysate of the leaf litter. Moreover, the fermentation
metabolites formed during the fermentation might have further enhanced the
adverse conditions presented to the bacterial cells.
Previous studies concluded that encapsulation of Cl. acetobutylicum ATCC
824 cells in gellan gum microspheres enhanced the tolerance of the cells to
adverse conditions in the fermentation medium. Moreover, the microspheres
acted as nurseries for continuous generation of free cells, thereby increasing
the cell density in the fermentation medium.
It was postulated that encapsulated cells could produce higher butanol yield
and productivity compared to free (non-encapsulated cells) owing to the
aforementioned reasons. In order to compare the efficiency of non-
encapsulated and encapsulated Cl. acetobutylicum ATCC 824 cells to utilise
fermentable sugars from S. saman leaf litter, fermentation studies were carried
out.
The fermentation profiles of non-encapsulated cells and encapsulated of Cl.
acetobutylicum ATCC 824 is shown in Figure 38. In the case of free cell
fermentation of detoxified acid hydrolysate of S. saman leaf litter, maximum
butanol concentration of 6.5 g/L was obtained after 48 h of fermentation. The
butanol production dropped continuously thereafter. It was assumed that the
free cells could not perform well due to the adverse conditions present in the
fermentation medium. A viable count of the cells at various stages of
fermentation was carried out to ascertain the effect of fermentation metabolites
on cell viability. As can be seen from Figure 39, the viable count of cells
169
dropped significantly from an initial high count to zero at the end of
fermentation. This clearly implied that the conditions in the fermentation
medium were very toxic for the cells. The combination of adverse effects of
solvents and inhibitors in the hydrolysate may have led to dramatic loss of cell
viability and consequent lower yield.
On the other hand, the fermentation efficiency of encapsulated Cl.
acetobutylicum ATCC 824 cells was found to be significantly higher than free
cells (p-value < 0.05). A lag phase of 24 h was observed in the case of
encapsulated Cl. acetobutylicum ATCC 824 cells. This was also observed in
Figure 38. Fermentation of detoxified acid hydrolysate by free and
encapsulated Cl. acetobutylicum ATCC 824 cells
0
1
2
3
4
5
6
7
8
9
10
0 24 48 72 96 120 144 168
Bu
tan
ol
yie
ld (
g/L
)
Fermentation time (h)
Free cells Encapsulated cells
170
Figure 39. Viability profiles of free and encapsulated cells of Cl.
acetobutylicum ATCC 824 during fermentation of detoxified acid
hydrolysate of S. saman leaf litter
previous studies with glucose as substrate. It reinstated the fact that the
encapsulated cells required longer time than the free cells to adjust to the
microenvironment inside the gellan gum microspheres. Moreover, since the
encapsulated cells were in spore form, they required time to germinate and
outgrow into vegetative cells to carry out fermentation. After the initial lag
phase, the butanol concentration increased steadily till 72 h with the maximum
concentration of 8.3 g/L butanol. This was significantly higher than that
achieved with free cells (p-value < 0.05). However, after 72 h, there was a
drop in butanol production, albeit at a slower rate than free cells. The viability
profile of cells liberated from the microspheres is shown in Figure 39. Cell
leakage was apparent after 24 h of fermentation and maximum viable cell
0
100
200
300
400
500
600
0 24 48 72 96 120 144 168
Via
ble
co
un
t (m
illi
on
/mL
)
Fermentation time (h)
Free cells Encapsulated cells
171
count was observed at 72 h. This is in correlation with the butanol
concentration observed in the fermentation medium. Thus, as the viable count
in the medium increased, there was a corresponding increase in butanol
concentration. However, due to the toxic conditions in the microspheres, the
viable cell count slowly diminished, resulting in drop in butanol concentration
as well. Interestingly, when the viable cell count reduced to approximately 10
X 108 cells /mL which were equivalent to that observed for free cells, the
butanol concentration was still 6.2 g/L. Again, this observation confirms the
earlier conclusion that the encapsulated cells also contribute to butanol
production apart from liberated cells.
The results of the current study showed that encapsulation of Cl.
acetobutylicum ATCC 824 cells in gellan gum microspheres is a viable option
to enhance the fermentability of acid hydrolysate of S. saman leaf litter.
Besides providing protection to the cells against both solvents and acid
hydrolysate inhibitors, the microspheres served as cell nurseries and
continuously generated free cells, thereby increasing cell density in the
fermentation medium.
172
CONCLUSIONS
173
V. CONCLUSIONS
The feasibility of microencapsulation technique as a cell immobilisation of Cl.
acetobutylicum ATCC 824 was investigated in this project. Spore form of the
bacterium was found to be suitable for microencapsulation by emulsification
technique. The gellan gum microspheres could be easily recovered and reused
for five fermentation cycles. The butanol yield and productivity of
encapsulated spores were lower than that achieved with free cells during first
fermentation cycle. This was mainly due to the lag time required by the
encapsulated spores to revive and acclimatise to the microenvironment of the
microspheres. Moreover, presence of the matrix barrier impaired mass transfer
of nutrients and metabolites, affecting the growth and fermentation
performances of the encapsulated cells. Additional time was also required for
liberation of encapsulated cells into the fermentation medium. The
fermentation efficiency of the microspheres improved over the number of
cycles, before it reduced by the fourth fermentation cycle. Gellan gum
microspheres were stable throughout the fermentation process. Significant cell
leakage was observed during the fermentation process. Thus, unlike the free
cells which could be reused only twice, Cl. acetobutylicum ATCC 824 cells
encapsulated in gellan gum microspheres could be reused for five consecutive
batch fermentations with acceptable fermentation efficiency. The superior
performance of the encapsulated cells can be explained by the protective effect
174
of gellan gum against the toxicity of high butanol concentrations as well as the
ability of the microspheres to function as nurseries.
In the second part of the project, the feasibility of utilisation of a readily
available municipal waste, in the form of S. saman leaf litter, as a cheap and
environmentally friendly biomass feedstock to generate fermentable sugars for
biofuel production was investigated. The leaf litter of S. saman consisted of
significant reducing sugars that could be recovered and utilised for biobutanol
production. Milling alone or in combination with hydrothermal treatment was
inadequate as a pretreatment method and necessitated additional dilute acid
coupled with heat treatment. Treatment of the milled leaf litter with dilute
sulphuric acid at 121 °C for 30 - 90 min was able to disrupt the lignocellulosic
matrix extensively and hydrolyse a major portion of the available
holocellulosic component of the leaves. Predictive pretreatment models built
using response surface methodology were useful for determining the optimal
acid concentration and treatment time. Dilute acid coupled with heat treatment
was able to release a large portion of fermentable sugars and subsequently a
low dose of enzyme was sufficient for the recovery of residual fermentable
sugars from the pretreated leaf litter. Further neutralisation with sodium
hydrolysate detoxified the acid hydrolysate and made it amenable for
biobutanol fermentation by free cells of Cl. acetobutylicum ATCC 824.
Fermentation studies by both free and encapsulated cells of Cl. acetobutylicum
ATCC 824 revealed that the latter could produce more butanol owing to the
protective effect of gellan gum microspheres on the encapsulated cells.
175
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LIST OF PUBLICATIONS/PRESENTATIONS
200
PUBLICATIONS/PAPERS PRESENTED AT SCIENTIFIC
MEETINGS
Journal publications
Rathore, S., Desai, P.M., Liew, C.V., Chan, L.W., Heng, P.W.S. (2012),
Microencapsulation of microbial cells. Journal of Food Engineering, 116,
369-381.
Manuscripts in preparation
Rathore, S., Chan, L.W., Heng, P.W.S. (2013), Optimisation of pretreatment
of Samanea saman leaf litter to obtain fermentable sugars for biofuel
production
Rathore, S., Chan, L.W., Heng, P.W.S. (2013), Feasibility study of
microencapsulation of Clostridium acetobutylicum cells by emulsification
method
Rathore, S., Chan, L.W., Heng, P.W.S. (2013), Investigation of sporulation
triggers in Clostridium acetobutylicum ATCC 824.
Oral Presentations
Rathore, S., Chan, L.W., Heng, P.W.S. (2010), Understanding growth
characteristics of Clostridium acetobutylicum for the production of bio-
products. International Pharmatech Conference on Drug Delivery 2010, Kuala
Lumpur, Malaysia.
201
Poster presentations
Rathore, S., Chan, L.W., Heng, P.W.S. (2012), Evaluation of
microencapsulation technique to immobilize Clostridium acetobutylicum cells
for the bio production of butanol. 26th AAPS Annual meeting and exposition,
Chicago, United States.
Rathore, S., Chan, L.W., Heng, P.W.S. (2011), Optimisation of acid
hydrolysis of an alternative lignocellulosic substrate using response surface
methodology. 25th AAPS Annual meeting and exposition, Washington DC,
United States.
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