1
PRODUCTION OF BIOSURFACTANT BY LOCALLY ISOLATED
BACTERIA FROM PETROCHEMICAL WASTE
RUZNIZA BINTI MOHD ZAWAWI
UNIVERSITI TEKNOLOGI MALAYSIA
2
3
4
PRODUCTION OF BIOSURFACTANT BY LOCALLY ISOLATED BACTERIA
FROM PETROCHEMICAL WASTE
RUZNIZA BINTI MOHD ZAWAWI
A thesis submitted in fulfilment of the
requirements for the award of the degree of
Master of Science (Chemistry)
Faculty of Science
Universiti Teknologi Malaysia
DECEMBER 2005
5
iii
Mohamad Najib Abdullah,
Mak & Ayah
Having you is all the happiness I have in this world…
iv
ACKNOWLEDGEMENT
I would like to give my special thank to my supervisor, Dr. Adibah Yahya for
her continuous guidance, attention, advices and inspiration that really help me to
finish up this project.
I am grateful to all family members of Research Laboratory 1 and
Microbiology and Molecular Biology Laboratory, Department of Biology, Faculty of
Science, especially to Mrs. Fatimah, Mrs. Radiah and all other postgraduate students
for their help and guidance in doing this research. Not forgotten to technicians of
Bioprocess Laboratory and Reservoir Laboratory, Faculty of Chemical Engineering
and Natural Resource for allowing me to use the research facilities in their
laboratory.
I also would like to thank all the lecturers and postgraduate students of
Microbiology and Enzyme Technology Laboratory, Faculty of Biotechnology and
Molecular Science, Universiti Putra Malaysia for their guidance and help during my
6 months of research attachment there. I am also indebted to MARA and IRPA under
vot 74048, for the funding of doing this research.
Last but not least, I would like to express my appreciation to my husband, my
family and my friends for the support and inspirations given to me to accomplish
these challenging years of study.
Thank you very much.
v
ABSTRACT
Ten bacterial strains previously isolated from petrochemical wastes were
selected for the screening of biosurfactant producer(s), via four different methods; (i)
surface tension measurements, (ii) blood hemolysis test, (iii) drop-collapsing test, and
(iv) bacterial adherence to hydrocarbon (BATH) test. Two isolates coded AB-Cr1
and ETL-Cr1 identified as Actinobacillus sp. and Aeromonas sp., respectively were
chosen to be the best candidates for biosurfactant production. Biosurfactant
productions by both isolates were found to be growth-associated in all conditions
tested. Biosurfactant production in glucose/crude oil medium (7.18-8.26 g/L) was
found similar to that observed in crude oil-free medium (6.33-8.76 g/L). The
production of biosurfactant was also studied in a fermentor using isolate AB-Cr1, as
a factor of temperature, initial glucose concentration, pH and initial nitrogen
concentration. The highest production of 12.45 g/L was obtained with AB-Cr1 grown
in medium (pH 7) supplemented with 25mM NH4NO3 as nitrogen source and 3mM
glucose as carbon source, incubated at 37°C under non-pH controlled strategy. TLC
and FTIR characterization of crude biosurfactant produced by both isolates in
medium supplemented or not with crude oil indicated the presence of lipoprotein and
non-aromatic glycolipid types of biosurfactant. GC-MS analysis of fatty acid metyl
esters indicated the presence of pentadecanoic acid in crude biosurfactant from both
isolates as well as octadecanoic and heptadecanoic acid in the biosurfactant produced
by AB-Cr1 and ETL-Cr1, respectively. The CMC of the biosurfactant produced in
the presence and absence of crude oil were approximately (g/L) 1.0 and 0.1 for ABCr1, and 1.2 and 0.2 for ETL-Cr1, respectively. The biosurfactants were found
capable of producing a relatively stable emulsion with hydrocarbon at pH 10. It was
also found stable at various pHs (3.0-13.0 and 5.0-9.0) for AB-Cr1 and ETL-Cr1,
respectively and thermostable for 1 hour at 100°C, based on the value of surface
tension.
vi
ABSTRAK
Sepuluh strain bakteria yang telah dipencilkan dari sisa petrokimia telah
dipilih untuk penyaringan bakteria penghasil-biosurfaktan, melalui empat kaedah; (i)
pengukuran ketegangan permukaan, (ii) ujian hemolisis darah, (iii) ujian keruntuhan
titisan, dan (iv) ujian pelekatan bakteria kepada hidrokarbon. Dua strain, AB-Cr1 dan
ETL-Cr1 dikenalpasti masing-masing sebagai Actinobacillus sp. dan Aeromonas sp.,
telah dipilih sebagai bakteria yang paling berpotensi menghasilkan biosurfaktan.
Penghasilan biosurfaktan oleh kedua-dua strain didapati bergantung kepada
pertumbuhan sel dalam semua keadaan ujian. Penghasilan biosurfaktan di dalam
medium glukosa/minyak mentah (7.18-8.26 g/L) didapati serupa dengan medium
tanpa minyak mentah (6.33-8.76 g/L). Penghasilan biosurfaktan oleh strain AB-Cr1
juga telah dijalankan di dalam fermenter terhadap faktor suhu, kepekatan awal
glukosa, pH dan kepekatan awal nitrogen. Penghasilan maksimum sebanyak 12.45
g/L didapati oleh AB-Cr1 di dalam media (pH 7) yang mengandungi 25mM NH4NO3
sebagai sumber nitrogen dan 3mM glukosa sebagai sumber karbon, pada suhu 37ºC
tanpa kawalan pH. Pencirian biosurfaktan mentah bagi kedua-dua strain melalui
kaedah TLC dan FTIR menunjukkan kehadiran biosurfaktan jenis lipoprotein dan
glikolipid bukan aromatic. Analisis GC-MS terhadap metil ester asid lemak
menunjukkan kehadiran asid pentadekanoik di dalam ekstrak biosurfaktan mentah
bagi kedua-dua strain dan juga asid oktadekanoik dan heptadekanoik di dalam
biosurfaktan yang masing-masing dihasilkan oleh AB-Cr1 dan ETL-Cr1. Nilai CMC
bagi biosurfaktan yang dihasilkan dengan dan tanpa minyak mentah adalah masingmasing (g/L) 1.0 dan 0.1 bagi AB-Cr1, dan 1.2 dan 0.2 bagi ETL-Cr1. Biosurfaktan
ini berupaya menghasilkan emulsi yang stabil terhadap hidrokarbon pada pH10. Ia
juga didapati stabil pada pelbagai pH (3.0-13.0 dan 5.0-9.0) bagi AB-Cr1 and ETLCr1, masing-masing dan stabil terhadap haba selama 1 jam pada 100ºC, berdasarkan
nilai ketegangan permukaan.
vii
CONTENTS
CHAPTER
TITLE
PAGE
SUPERVISOR’S APPROVAL
THESIS TITLE
i
DECLARATION
ii
DEDICATION
iii
ACKNOWLEDGEMENT
iv
ABSTRACT
v
ABSTRAK
vi
CONTENTS
vii
LIST OF ABBREVIATION
xiv
LIST OF TABLES
xv
LIST OF FIGURES
xvii
LIST OF APPENDICES
xxii
1
INTRODUCTION
1.1
General Overview: Surfactant and Biosurfactant
1
1.2
Scope and Objectives of the Current Project
4
2
LITERATURE REVIEW
2.1
Introduction to Biosurfactant
6
2.1.1
6
Definition and Classification
viii
2.1.2
2.2
2.3
2.4
Structure and Properties of Biosurfactant
7
2.1.2.1 Glycolipids
8
2.1.2.2 Lipoproteins and Lipopeptides
9
2.1.2.3 Fatty acids, Phospholipids and Neutral Lipids
10
2.1.2.4 Polymeric Biosurfactants
10
2.1.2.5 Particulate Biosurfactants
11
Screening of Biosurfactant-producing Bacteria
12
2.2.1
Cell Hydrophobicity Test
13
2.2.2
Drop-collapsing Technique
13
2.2.3
Hemolytic Activity
14
2.2.4
Surface Tension Reduction
15
Biosynthesis of Biosurfactant
16
2.3.1
General Features of Biosynthesis
16
2.3.2
Biosynthetic Pathway of Biosurfactant Synthesis
18
2.3.3
Regulation of Biosurfactant Synthesis
20
Production of Biosurfactant
22
2.4.1
Factors Affecting Biosurfactant Production
22
2.4.1.1 Effect of Carbon Source
22
2.4.1.2 Effect of Nitrogen Source
23
2.4.1.3 Effect of Environmental Factors
23
Kinetics of Biosurfactant
24
2.4.2.1 Growth-associated Biosurfactant
25
2.4.2
Production
2.4.2.2 Biosurfactant Production Under Growth-
26
limiting Conditions
2.4.2.3 Biosurfactant Production by Resting or
27
Immobilized Cells
2.4.2.4 Biosurfactant Production in Addition to
28
Precursors
2.5
Extraction of Biosurfactant
28
2.6
Applications and Roles of Biosurfactant
30
2.7
Characteristics of Chemical Surfactant and Biosurfactant
33
2.7.1
34
Advantages and Disadvantages of Biosurfactants
in Commercial Application
ix
3
GENERAL MATERIALS AND METHODS
3.1
Microorganisms
35
3.1.1
Bacterial Isolates: Origin and Route of Isolation
35
3.1.2
Crude Oil
36
3.2
Media Preparation
38
3.2.1
Liquid Medium
38
3.2.1.1 Ramsay Liquid Medium
38
Solid Media
38
3.2.2.1 Nutrient Agar
38
3.2.2.2 Ramsay Agar
38
3.2.2.3 Blood Agar
39
3.2.2
3.3
3.4
3.5
4
Growth and Maintenance of Bacterial Isolates
39
3.3.1
Inoculum Preparation
39
3.3.2
Culture Maintenance and Storage
39
Analytical Methods
40
3.4.1
Determination of Bacterial Biomass
40
3.4.1.1 Optical Density
40
3.4.1.2 Cell Dry Weight
40
3.4.2
Determination of Glucose Concentrations
40
3.4.3
Surface Activity Measurements
41
3.4.3.1 Surface Tension Measurement
41
3.4.3.2 Interfacial Tension Measurement
41
3.4.3.3 Spreading Tension Measurement
42
Production of Biosurfactant
42
3.5.1
Biosurfactant Extraction
42
3.5.2
Determination of Biosurfactant Dry Weight
43
SCREENING AND CHARACTERIZATION OF
BIOSURFACTANT-PRODUCING BACTERIA
4.1
Introduction
44
4.2
Methodology
45
4.2.1
45
Screening of Biosurfactant-producing Bacteria
x
4.2.1.1 Bacterial Adherence To Hydrocarbon
45
(BATH) Test
4.2.2
4.2.1.2 Drop-collapsing Test
45
4.2.1.3 Blood Hemolysis Test
46
4.2.1.4 Surface Tension Measurement
46
Characterization of Biosurfactant-producing
47
Isolates
4.3
4.2.2.1 Morphological Analysis
47
4.2.2.2 Biochemical Analysis
47
Results and Discussion
48
4.3.1
Screening of Biosurfactant-producing Bacteria
48
4.3.1.1 Bacterial Adherence To Hydrocarbon
48
(BATH) Test
4.3.2
4.3.1.2 Drop-collapsing Test
50
4.3.1.3 Blood Hemolysis Test
52
4.3.1.4 Surface Tension Measurement
54
Characterization of the Selected Biosurfactant-
55
producing Isolates
4.3.2.1 Colony and Cellular Morphological
55
Characterizations
4.3.2.2 Biochemical Characterization
5
57
PRODUCTION OF BIOSURFACTANT BY PURE AND
MIX BACTERIAL CULTURES IN SHAKE FLASKS
5.1
Introduction
5.2
Methodology
5.2.1
58
Optimization of Growth
59
5.2.1.1 Effect of Initial Glucose Concentrations
59
on Growth
5.2.2
5.2.1.2 Effect of Initial pH on Growth
59
5.2.1.3 Effect of Temperature on Growth
60
Biosurfactant Production under the Optimized
60
Growth Condition
xi
5.2.3
Effect of Glucose and Crude Oil on
61
Biosurfactant Production
5.2.4
Production of Biosurfactant by Bacterial
61
Mix Cultures
5.3
Results and Discussion
62
5.3.1
Optimization of Growth
62
5.3.1.1 Effect of Initial Glucose Concentrations
62
on Growth
5.3.2
5.3.1.2 Effect of Initial pH on Growth
65
5.3.1.3 Effect of Temperature on Growth
67
Biosurfactant Production under the Optimized
69
Growth Condition
5.3.3
Production of Biosurfactant in Crude Oil-
71
containing Medium
5.3.4
Production of Biosurfactant by Bacterial
79
Mix Cultures
6
PRODUCTION OF BIOSURFACTANT BY STRAIN
AB-Cr1 IN BIOREACTOR
6.1
Introduction
84
6.2
Methodology
85
6.2.1
Batch Fermentation
85
6.2.1.1 Effect of Temperature on Biosurfactant
85
Production
6.2.1.2 Effect of Initial Glucose Concentrations on
86
Biosurfactant Production
6.2.1.3 Effect of Controlled pH Condition on
86
Biosurfactant Production
6.2.1.4 Effect of Initial NH4NO3 Concentrations on
87
Biosurfactant Production
6.3
Results and Discussion
87
6.3.1
Effect of Temperature on Biosurfactant Production
87
6.3.2
Effect of Initial Glucose Concentrations on
93
xii
Biosurfactant Production
6.3.3
Effect of Controlled pH Condition on
97
Biosurfactant Production
6.3.4
Effect of Initial NH4NO3 Concentrations on
102
Biosurfactant Production
7
CHARACTERIZATION OF CRUDE BIOSURFACTANT
7.1
Introduction
108
7.2
Methodology
109
7.2.1
Emulsification Activity Tests
109
7.2.1.1 Assay of Emulsification
109
7.2.1.2 Assay of Emulsion Stability
109
Critical Micelle Concentration (CMC)
109
7.2.2
Determination
7.3
7.2.3
Stability Studies
110
7.2.4
Thin Layer Chromatography (TLC)
110
7.2.5
Fourier Transform Infrared (FTIR)
111
7.2.6
Fatty Acid Analysis
111
Results and Discussion
112
7.3.1
Emulsification Activities
112
7.3.2
Critical Micelle Concentration (CMC)
114
7.3.3
Stability Studies
116
7.3.4
Thin Layer Chromatography (TLC)
118
7.3.5
Fourier Transform Infrared (FTIR)
120
7.3.6
Fatty Acid Analysis
124
8
GENERAL DISCUSSION AND CONCLUSION
8.1
Conclusion
128
8.2
Suggestion
131
REFERENCES
133
xiii
APPENDICES A-G
147
xiv
LIST OF ABBREVIATIONS
mN/m
-
Milinewton per meter
g/L
-
Gram per litre
mL
-
Mililitre
ºC
-
Degree Celsius
rpm
-
Rotation per minute
nm
-
Nanometer
w/v
-
Weight per volume
v/v
-
Volume per volume
CMC
-
Critical Micelle Concentration
µ
-
Specific growth rate
Pmax
-
Maximum product concentration
Xmax
-
Maximum biomass concentration
Yp/s
-
Product yield coefficient (g product formed per g
substrate utilized)
Yp/x
-
Product yield coefficient (g product formed per g
biomass formed)
Yx/s
-
Biomass yield coefficient (g biomass formed per g
substrate utilized)
et al.
-
And friends
sp.
-
Species
h
-
Hour
NH4NO3
-
Ammonium nitrate
HCl
-
Hydrochloric acid
Kd
-
Decay constant
xv
LIST OF TABLES
TABLES
TITLE
PAGE
2.1
Various biosurfactants produced by different microbes.
6
2.2
Common methods employed for the recovery of
29
biosurfactants.
2.3
Some properties of biosurfactant commonly used in
32
several applications.
2.4
Differences between biosurfactant and synthetic
33
surfactant.
3.1
Origin of bacteria isolated from petroleum-related
37
industries.
4.1
Screening of biosurfactant-producing bacteria using
48
four different methods.
4.2
Results for biochemical tests of the selected isolates.
57
5.1
Specific growth rates and maximum biomass of
65
AB-Cr1 and ETL-Cr1 grown at 37ºC, pH 6.5-6.8 in
medium supplemented with various initial glucose
concentrations.
5.2
Specific growth rates and maximum biomass of
67
AB-Cr1 and ETL-Cr1 grown in Ramsay medium
supplemented with 3mM glucose adjusted to various
initial pH.
5.3
Specific growth rates and maximum cell biomass of
68
AB-Cr1 and ETL-Cr1 grown in medium
supplemented with 3mM glucose at pH 7.0,
incubated at various temperatures.
5.4
Kinetic analysis of growth and biosurfactant
77
xvi
production for isolates AB-Cr1 and ETL-Cr1
grown at 37ºC, in medium supplemented with either
glucose or crude oil or both glucose and crude oil.
5.5
Kinetic analysis of growth and biosurfactant
82
production for bacterial mix culture system 1:1
(AB-Cr1:ETL-Cr1) grown at 37ºC, in medium
supplemented with either glucose or both glucose
and crude oil.
6.1
Kinetic analysis for growth and biosurfactant
95
production by AB-Cr1 grown at 37ºC, in medium
supplemented with various initial glucose
concentrations.
6.2
Kinetic analysis for growth and biosurfactant
101
production by AB-Cr1 grown in medium controlled
at various pH values, supplemented with 3mM glucose
and incubated at 37ºC.
7.1
Emulsification activity and stabilization of
113
bioemulsifiers by isolated biosurfactants.
7.2
TLC analysis of biosurfactant produced by AB-Cr1
118
and ETL-Cr1 isolates based on the Rf values.
7.3
Relative positions of peaks from GC-MS for methyl
esters of fatty acids.
125
xvii
LIST OF FIGURES
FIGURE
2.1
TITLE
PAGE
Structure of rhamnolipid produced by Pseudomonas
9
aeruginosa.
2.2
Structure of surfactin produced by Bacillus subtilis.
10
2.3
The amphiphilic structure of a surfactant.
12
2.4
Metabolic pathway of glucose utilization during
19
biosurfactant production
2.5
Schematic illustration showing different types of
25
fermentation kinetics of biosurfactant production.
4.1
-hemolysis on blood agar indicated to the presence
53
of biosurfactant in the culture of AB-Cr1 and
ETL-Cr1.
4.2
Colony of AB-Cr1 observed under stereo scan
56
microscope using magnification 50x.
4.3
Colony of ETL-Cr1 observed under stereo scan
56
microscope using magnification 50x.
4.4
Digital photos of bacterial isolates AB-Cr1 and ETL-
56
Cr1 under phase-contrast microscope using
magnification 100x with oil immersion.
5.1
Growth curve of AB-Cr1 grown in Ramsay medium
63
pH 6.5-6.8 at 370C as a factor of initial glucose
concentrations.
5.2
Growth curve of ETL-Cr1 grown in Ramsay medium
63
0
pH 6.5-6.8 at 37 C as a factor of initial glucose
concentrations.
5.3
The specific growth rates of AB-Cr1 and ETL-Cr1
64
xviii
grown in Ramsay medium pH 6.5-6.8 at 370C,
as a factor of initial glucose concentrations.
5.4
Growth optimization of isolates AB-Cr1 and ETL-Cr1
66
grown at 370C in medium supplemented with 3mM
glucose, as a factor of pH.
5.5
Growth optimization of isolates AB-Cr1 and ETL-Cr1
67
grown in medium supplemented with 3mM glucose
at pH 7.0, as a factor of temperature.
5.6
Relationship of growth, glucose consumption and
69
biosurfactant production by AB-Cr1 isolate grown in
Ramsay medium supplemented with 3mM glucose,
adjusted to initial pH 7.0 and incubated at 370C.
5.7
Relationship of growth, glucose consumption and
70
biosurfactant production by ETL-Cr1 isolate grown
in Ramsay medium supplemented with 3mM glucose,
adjusted to initial ph 7.0 and incubated at 370C.
5.8
Relationship of growth, glucose consumption, pH,
72
surface tension and biosurfactant production for
isolates AB-Cr1 and ETL-Cr1 grown in Ramsay
medium supplemented with glucose and crude oil,
respectively.
5.9
Relationship of growth and biosurfactant production
73
by isolates AB-Cr1 and ETL-Cr1 grown in Ramsay
medium supplemented with 5% (v/v) crude oil.
5.10
Relationship between specific growth rates (µ) of
79
isolates AB-Cr1 and ETL-Cr1 with the specific rates
of product formation (q p) in medium supplemented
with either (i) crude oil, or (ii) both glucose and
crude oil, or (iii) glucose.
5.11
Relationship of growth and biosurfactant production
by bacterial mix culture system 1:1, grown
in Ramsay medium supplemented with glucose and
glucose + crude oil.
80
xix
6.1
Maximum cell biomass and biosurfactant production
88
by AB-Cr1 grown in medium supplemented with
3mM glucose, as a factor of temperature.
6.2
Relationship between biosurfactant production, growth
89
and oxygen consumption (A), glucose consumption
and pH (B), surface, interfacial and spreading tension
(C) by AB-Cr1, grown in medium supplemented with
3mM glucose adjusted to initial pH 7.0 and incubated
at 37ºC.
6.3
Surface and interfacial tension reduction of the cell-free
92
culture of AB-Cr1 grown in medium supplemented
with 3mM glucose, as a factor of temperature.
6.4
The yield coefficients for biosurfactant and biomass
93
production by AB-Cr1, grown in medium supplemented
with 3mM glucose, as a factor of temperature.
6.5
Maximum cell biomass and biosurfactant production
94
by AB-Cr1 grown at 37ºC, as a factor of various initial
glucose concentrations.
6.6
Maximum cell biomass and biosurfactant production
97
by AB-Cr1 grown in medium supplemented with 3mM
glucose at 37ºC, as a factor of pH.
6.7
Surface tension and interfacial tension reduction of the
99
cell-free culture of AB-Cr1, grown in medium
supplemented with 3mM glucose at 37ºC, as a factor
of pH.
6.8
The relationship between biosurfactant production,
100
growth and oxygen consumption (A), surface and
interfacial tension (B) by AB-Cr1 grown in medium at
controlled pH 7.0 and incubated at 37ºC.
6.9
Maximum biomass and biosurfactant production
103
by AB-Cr1 grown in medium supplemented with
3mM glucose at 37ºC, as a factor of various initial
NH4NO3 concentrations.
6.10
The relationship between biosurfactant production,
104
xx
growth and oxygen consumption (A), surface and
interfacial tension (B), by AB-Cr1 grown in medium
supplemented with 15mM NH4NO3 and incubated
at 37ºC.
6.11
The yield coefficients for biosurfactant and biomass
106
production by AB-Cr1 grown in medium supplemented
with 3mM glucose and incubated at 37ºC, as a factor of
various initial NH4NO3 concentrations.
7.1
Effect of pH on the activity of the emulsifier
112
produced by AB-Cr1 and ETL-Cr1 isolates.
7.2
Schematic diagram of the variation of surface tension,
114
interfacial tension and the CMC point with surfactant
concentration.
7.3
Surface tension of a solution against the concentration
115
of the biosurfactant produced by AB-Cr1 and ETLCr1, grown in medium supplemented with glucose
as sole source of carbon.
7.4
Surface tension of a solution against the concentration
116
of the biosurfactant produced by AB-Cr1 and ETLCr1, grown in medium supplemented with glucose
and crude oil.
7.5
The pH stability test of biosurfactant produced by AB-
117
Cr1 and ETL-Cr1 grown in medium supplemented
with glucose, based on the change of surface tension
values.
7.6
Thermal stability test of biosurfactant produced by
117
AB-Cr1 and ETL-Cr1 grown in medium supplemented
with glucose, based on the change of surface tension
values.
7.7
Infrared spectrum of the surface-active fraction
120
extracted from culture of AB-Cr1 grown in medium
supplemented with glucose as the sole source of carbon.
7.8
Infrared spectrum of the surface-active fraction
extracted from culture of ETL-Cr1 grown in
121
xxi
medium supplemented with glucose as the sole
source of carbon.
7.9
Infrared spectrum of the surface-active fraction
122
extracted from culture of AB-Cr1 grown in
medium supplemented with both glucose and
crude oil as carbon sources.
7.10
Infrared spectrum of the surface-active fraction
123
extracted from culture of ETL-Cr1 grown in
medium supplemented with both glucose and
crude oil as carbon sources.
7.11
GC-MS chromatogram of fatty acid methyl ester
124
from a culture medium of AB-Cr1.
7.12
GC-MS chromatogram of fatty acid methyl ester
125
from a culture medium of ETL-Cr1.
7.13
Structure of pentadecanoic acid.
126
7.14
Structure of octadecanoic acid.
126
7.15
Structure of heptadecanoic acid.
126
7.16
Mass spectrum of pentadecanoic acid from a
127
culture of AB-Cr1.
7.17
Mass spectrum of octadecanoic acid from a
127
culture of AB-Cr1.
7.18
Mass spectrum of heptadecanoic acid from a
culture of ETL-Cr1.
127
xxii
LIST OF APPENDICES
APPENDIX
TITLE
PAGE
A
Graft OD600 versus cell biomass
147
B
Glucose standard curve
148
C
Biochemical characterization methods
149
D
Production of biosurfactant and surface tension
161
reduction in the medium grown with AB-Cr1 isolate
E
Relationship of growth, glucose consumption and
162
biosurfactant production by bacterial mix culture
system 1:1, grown in Ramsay medium supplemented
with glucose and glucose + crude oil
F
Determination of decay constant
163
G
Mass spectrums of fatty acid methyl esters from
164
the culture of AB-Cr1 and ETL-Cr1 isolates.
CHAPTER 1
INTRODUCTION
1.1
General Overview: Surfactant and Biosurfactant
Surfactants are amphiphilic compounds that reduce the free energy of the
system by replacing the bulk molecules of higher energy at an interface [Mulligan,
2004]. They contain a hydrophobic moiety with little affinity for the bulk medium
and a hydrophilic portion that is attracted to the bulk medium. Surfactants have been
used industrially as adhesives, deemulsifiers, flocculating, wetting and forming
agents, lubricants and penetrants [Mulligan and Gibbs, 1993].
Because of their amphiphilic nature, surfactants tend to accumulate at
interfaces (air-water and oil-water) and surfaces. As a result, surfactants reduce the
forces of repulsion between unlike phases at interfaces or surfaces and allow the two
phases to mix more easily [Bodour and Miller-Maier, 2002]. Due to the presence of
surfactant, less work is required to bring a molecule to the surface and the surface
tension is reduced. The ability to reduce surface tension is a major characteristic of
surfactant. It is obvious that their surface and membrane-active properties play an
important role in the expression of their activities.
Surfactants are key ingredients used in detergents, shampoos, toothpaste, oil
additives, and a number of other consumer and industrial products. They constitute
an important class of industrial chemicals widely used in almost every sector of
modern industry. The total surfactant production has exceeded 2.5 million tones in
2002 [Deleu and Paquot, 2004] for many purposes such as polymers, lubricants and
2
solvents. The growth rate is related to the world demand in detergents since this
sector uses over 50% of surfactant production [Deleu and Paquot, 2004].
From the total surfactants output, about 54% of them is consumed as
household or laundry detergents, with only 32% destined for industrial use
[Cameotra and Makkar, 1998]. Almost all surfactants currently in use are chemically
derived from petroleum. The choice of surfactant is based on product cost. Generally,
surfactants has been extensively used to save energy and consequently energy cost.
For example, the new generation of detergents wash effectively at much lower
temperatures, resulting in significant energy saving. Physicochemical behavior,
charge-type, solubility and adsorption behavior are some of the most important
selection criteria for surfactants [Mulligan, 2004].
However, as many industry and research organizations concern to the
environmental approach, they are currently attempted to find new ways of producing
surfactants. There are two new strategic approaches that are taken into account in
developing new surfactant, which are i) the impact of the surfactant to the
environment and ii) the functionalities of the surface-active molecules. Synthetic
surfactants exhibit a low rate of biodegradation and a high potential to aquatic
toxicity. For these reasons, biosurfactants are seen to be the promising alternative for
many purposes even though their performance could be slightly inferior or their
prices are more expensive.
Biosurfactant is a structurally diverse group of surface-active molecule
synthesized by microorganisms. Their capability of reducing surface and interfacial
tension with low toxicity and high specificity and biodegradability, lead to an
increasing interest on these microbial products as alternatives to chemical surfactants
[Banat et al., 2000]. Hester (2001) from the Technical Insights estimated that
biosurfactants could capture 10% of the surfactant market by the year 2010 with sales
of $US200 million. However, up to now, biosurfactants is still unable to compete
with the chemically synthesized surfactants in the surfactant market. This could be
due to their high production costs in relation to inefficient bioprocessing method
available, poor strain productivity and the need to use expensive substrates
[Cameotra and Makkar, 1998; Deleu and Paquot, 2004].
The interest in biosurfactant has been steadily increasing in recent years due
to the possibility of their production through fermentation and their potential
3
applications in such areas as the environmental protection. The uniqueness with
unusual structural diversity, the possibility of cost-effective ex-situ production and
their biodegrability are some of the properties that make biosurfactant a promising
choice for use in environmental application [Hua et al., 2003].
Initial focus of industrial interest towards biosurfactants concentrates on the
microbial production of surfactants, cosurfactants and so on for the application on
microbial-enhanced oil recovery (MEOR) [Kosaric et al., 1987]. The applications of
biosurfactants however, are still currently remained at the developmental stage of
industrial level. The development of biosurfactant application in industries has
focused mainly on high biosurfactant production yield and the production of highly
active biosurfactants with specific properties for specific applications.
Majority of surfactants produced today is of petrochemical origin beside of
the renewable resources like fats and oils [Deleu and Paquot, 2004]. Amongst the
renewable raw materials, oleochemical products represent half of the total surfactant
production. The petrochemical industry is one of the important sector in Malaysia,
with investments totaling RM28 billion as at the end of 2002 [Mida Malaysia].
Exxon Mobil is one of the multinational petrochemical companies that work in
collaboration with Malaysia’s national petroleum company, Petronas. This
collaboration clearly make Malaysia as a potential country as an investment location
for petrochemical industries.
Unfortunately, industrial wastewater from petroleum-related industries has
been identified as one of the major source of pollution in Malaysia. The
biodegradation of petroleum pollutant and its related compound is limited by poor
availability to the microorganisms, due to their hydrophobicity and low aqueous
solubility. This suggested that by applying biosurfactants to influence the
bioavailability of the contaminant, can possibly enhancing the solubility of these
compounds. Due to their biodegradability and low toxicity, they are in demand to be
use in remediation technologies [Mulligan, 2004].
At present, biosurfactants plays an important application in petroleum-related
industries which is use in enhanced oil recovery, cleaning oil spills, oil-contaminated
tanker cleanup, viscosity control, oil emulsification and removal of crude oil from
sludges [Daziel et al., 1996, Bertrand et al., 1994]. These industries are known to be
the potential target for the application of these compounds. This is due to the ability
4
of biosurfactant-producing microorganisms to use petroleum or its’ products as
substrates as well as the properties of the biosurfactant which required less rigorous
testing than chemical surfactant [Cooper, 1986].
To date, there are numbers of reports on the synthesis of various types of
biosurfactants by microorganisms using water-soluble compounds such as glucose,
sucrose, ethanol or glycerol as substrates [Desai and Banat, 1997]. Petroleum-related
industry was found to be one of the industries that have a great potential in producing
a microorganism that may produced biosurfactants. Hence, there could probably be a
potential chance of producing biosurfactants using locally isolated bacteria originated
from petrochemical wastes or other wastewater available in this country. It has been
focused here that improving the method of biosurfactant production and
characterizing the major properties of the biosurfactant are highly important in the
commercial application of biosurfactant.
1.2
Scope and Objectives of the Current Project
The present study focused on studying the production of biosurfactant by
bacteria isolated from petrochemical wastes. Ten bacterial isolates were screened for
potential biosurfactant producer(s) and two of them were found able to produce
biosurfactant by various screening methods. It was therefore of interest to
characterize these bacteria and study their ability to produce biosurfactant.
The major part of this thesis describes research into the production of
biosurfactant by these bacteria in various conditions tested. The study was initiated
with basic identification based on cellular and colony morphologies, followed
biochemical characteristics of these bacteria. The study on production of
biosurfactant by these isolates was initiated by optimizing the growth of the potential
biosurfactant producers as the factor of several parameters such as initial glucose
concentration, initial pH and incubating temperature. The ability of these bacteria to
produce biosurfactant as single and mix bacterial cultures, in medium supplemented
with glucose and/or crude oil were then studied using the optimum growth
conditions. Optimization of biosurfactant production by the best biosurfactant
producer was further studied in bioreactor as a factor of temperature, initial glucose
5
concentration, pH and initial nitrogen concentration. This study was also sought to
the preliminary characterization of the crude biosurfactant produced by means of
their physicochemical properties. Characterization studies included emulsification
activity, critical micelle concentration (CMC), stability test, thin layer
chromatography (TLC), fourier transform infrared (FTIR) and gas chromatographymass spectrometry (GC-MS) analyses.
In general, the objective of this research is to study the biosurfactant
production by microbial fermentation process and characterized the crude
biosurfactant in order to determine their physicochemical properties. Therefore, this
study is conducted with the specific objectives:
§
To screen and characterize the potential biosurfactant-producing microbes
from petrochemical waste samples.
§
To optimize the biosurfactant production in terms of productivity and the
yield of biosurfactants from the substrates.
§
To characterize the crude biosurfactant produced by the bacterial isolates.
CHAPTER 2
LITERATURE REVIEW
2.1
Introduction to Biosurfactant
2.1.1
Definition and Classification
A biosurfactant is defined as a surface-active agent produced by living cells,
mostly by microorganisms [Fiechter, 1992]. The molecule of biosurfactant, which
has both water-soluble and water insoluble parts in the same molecule, balance the
hydrophilic and hydrophobic moieties, imparts unusual properties including an
ability to lower the surface tension of water [Cooper, 1986].
Basically, there are six major types of biosurfactant; hydroxylated and
crosslinked fatty acids (mycolic acids), glycolipids, polysaccharide-lipid complexes,
lipoproteins-lipopeptides, phospholipids and the complete cell surface itself [Kosaric
et al., 1987]. Table 2.1 showed some of the various biosurfactants produced from
different microbes.
Table 2.1: Various biosurfactants produced by different microbes.
Biosurfactants
Microrganisms
References
Rhamnolipid
Pseudomonas aeruginosa
Sophorolipid
Candida bombicola
Lang and Wagner (1987); Haba et
al. (2000)
Schippers et al. (2000)
7
Surfactin
Bacillus subtilis
Fatty acid
Corynebacterium lepus,
Neutral lipid
Lipopeptide
Nocardia erythropolis
Arthrobacter MIS38, Bacillus
subtilis, Pseudomonas
fluorescens
Acinetobacter sp.,
Corynebacterium insidiosum
Phospholipid
Arima et al. (1968); Moran et al.
(2002)
Cooper et al. (1979); Rosenberg
and Ron, 1999
MacDonald et al. (1981)
Morikawa et al. (1993); Akpa et
al. (2001); Braun et al. (2001)
Kaeppeli and Finnerty (1980);
Akit et al. (1981)
Microbial surfactants can also be divided into two major classes according to
their molecular-mass. The low molecular-mass biosurfactants include glycolipids
such as rhamnolipids and sophorolipids, or lipopeptides like surfactin and
polymyxin, has a function in lowering the surface and interfacial tensions. Whereas
the high molecular-mass biosurfactants such as lipoproteins, lipopolisaccharides and
amphipathic polysaccharides are more effective at stabilizing oil-in-water emulsions
[Rosenberg and Ron, 1999].
According to Rosenberg and Ron (1999), there are three possible roles of
biosurfactant in applications; increasing the surface area of hydrophobic substances,
increasing the bioavailability of hydrophobic water-insoluble substrates, and finally
regulating the attachment-detachment of microorganisms to and from surfaces.
2.1.2
Structure and Properties of Biosurfactant
Biosurfactants are classified mainly by their chemical composition and
microbial origin. The major classes of biosurfactant include glycolipids, lipopeptides
and lipoproteins, phospholipids and fatty acids, polymeric surfactants and particulate
surfactants [Desai and Banat, 1997].
8
2.1.2.1 Glycolipids
Glycolipids are carbohydrates in combination with long-chain aliphatic acids
or hydroxyaliphatic acids. These include the rhamnolipids, sophorolipids,
trehalolipids and fructose-lipids.
Rhamnolipid is a glycolipid that contains one or two molecules of rhamnose
that are linked to one or two molecules of β-hydroxydecanoic acid. Up to 7
homologues have now been identified [Abalos et al., 2001]. L-Rhamnosyl-Lrhamnosyl-β-hydroxydecanoyl-β-hydroxydecanoate and L-rhamnosyl-βhydroxydecanoyl-β-hydroxydecanoate, referred to as rhamnolipid 1 and 2,
repectively, are the principal glycolipids produced by P. aeruginosa [Desai and
Banat, 1997]. Rhamnolipids from P. aeruginosa (Figure 2.1) have been demonstrated
to lower the surface tension to 25 to 30mN/m and the interfacial tension against nhexadecane to 1mN/m [Lang and Wagner, 1987; Mulligan, 2004].
Different species of the yeast Torulopsis produce extracellular sophorolipids,
which consist of dimeric carbohydrate sophorose linked to -1,2 long chain
hydrocarboxylic acids. The lipid portion is connected to the reducing end through a
glycosidic linkage [Rosenberg and Ron, 1999]. These biosurfactants are a mixture of
at least 6-9 different hydrophobic sophorosides.
Sophorolipids has been reported capable of lowering both surface and
interfacial tension, though they are not effective emulsifying agents [Cooper and
Paddock, 1984; Kitamoto et al., 2002]. Both lactonic and acidic sophorolipids
lowered the surface tension to 33mN/m and the interfacial tension against nhexadecane and water from 40 to 5mN/m with 10mg/L of pure sophorolipid. It is
also showed a remarkable stability toward pH and temperature changes. The surfaceactive properties were consistent between pH values of 6-9 and temperature ranging
from 20-90ºC.
9
O
CH 3
CH
(CH 2) 6
CH 3
CH 2
O
C=O
HO
O
OH
H3C
O
O
CH
(CH 2 )6
CH 3
CH 2
HO
OH OH
COOH
Fig. 2.1: Structure of rhamnolipid produced by Pseudomonas aeruginosa.
2.1.2.2 Lipoproteins and Lipopeptides
Lipopeptides usually appear as mixtures of closely related compounds which
show slight variations in their amino acid composition and/or lipid portion which is
mostly a hydroxy fatty acid. A family of cyclic lipopeptides consists of 8 to 17 amino
acids and a lipid portion which is composed of 8 to 9 methylene groups and a
mixture of linear and branched tails [Desai and Banat, 1997]. These include surfactin
produced by B. subtilis [Lang, 2002] (Figure 2.2) and viscosin from P. fluorescence
[Hommel, 1990; Braun et al., 2001].
Surfactin, a cyclic acidic lipopeptide produced by B. subtilis, is one of the
most effective biosurfactants known so far [Arima et al., 1968; Mulligan, 2004]. It
contains seven amino residues and is closed by lactone formation [Carillo et al.,
2003]. Surfactin is known to be capable of lowering the surface tension from 72 to
27.9mN/m at a concentration of 0.005% (w/v). An important characteristic of this
compound is its ability to lyse red blood cells and may act as an antibiotic, antiviral
and hemolytic agent [Arima et al., 1968; Carillo et al., 2003; Cameotra and Makkar,
2004]. This property has been used to detect surfactin production through blood agar.
10
Fig 2.2: Structure of surfactin produced by Bacillus subtilis [Hommel, 1990;
Mulligan, 2004].
2.1.2.3 Fatty Acids, Phospholipids and Neutral Lipids
Fatty acid and phospholipid produced during growth on n-alkanes by several
bacteria and yeast, has received considerable attention as surfactants [Rosenberg et
al., 1999]. These biosurfactants are able to produce optically clear microemulsions of
alkanes in water [Desai and Desai, 1993]. The hydrophilic lipophilic balance (HLB)
of fatty acids is found clearly related to the length of the hydrocarbon chain. Example
of microorganisms that produced these types of biosurfactant are sulphur-reducing
bacteria, Thiobacillus thiooxidans [Beeba and Umbreit, 1971; Desai and Banat,
1997] and Corynebacterium lepus [Cooper et al., 1979; Rosenberg and Ron, 1999].
Extracellular free fatty acids produced by microorganisms grown on alkanes
also showed surfactant activity. They include saturated fatty acids in the range of C12 to C-14 and the complex fatty acids containing hydroxyl groups and alkyl
branches. Phosphatidylethanolamine produced by Rhodococcus erythropolis caused a
lowering of interfacial tension against hexadecane to less than 1mN/m and a CMC of
30mg/L [Kretschmer et al., 1982; Rosenberg and Ron, 1999].
2.1.2.4 Polymeric Biosurfactants
Many bacterial species from different genera produced exocellular polymeric
surfactants composed of proteins, polysaccharides, lipopolysaccharides or complex
mixture of these biopolymers [Rosenberg et al., 1999]. Fatty acids are covalently
linked to the polysaccharide through o-ester linkages [Zukerberg et al., 1979; Desai
11
and Banat, 1997]. The best-studied polymeric biosurfactants are emulsan, liposan and
mannoprotein [Desai and Banat, 1997].
Emulsan has been characterized as a polyanionic amphipatic
heteropolysaccharide [Rosenberg et al., 1979; Rosenberg and Ron 1999]. It is a very
effective emulsifying agent for hydrocarbons in water even at a concentration of
0.001%. Emulsan also is one of the most powerful emulsion stabilizer known today
and resists inversion even at a water-oil ratio of 1:4 [Desai and Banat, 1997].
2.1.2.5 Particulate Biosurfactants
This type of biosurfactant includes vesicles and fimbriae produced by
Acinetobacter sp. The purified vesicles are composed of protein, phospholipid and
lipopolisaccharide. This extracellular membrane vesicles partition hydrocarbons form
a microemulsion, which plays an important role in alkane uptake by microbial cells
[Kappeli and Finnerty, 1979; Desai and Banat, 1997].
Generally, biosurfactant molecules consisted of both hydrophilic and
hydrophobic moieties (Figure 2.3), which enable them to accumulate at the interfaces
and mediated between phases of different polarity such as oil-in-water or water-in-oil
interfaces [Fiechter, 1992]. The polar, water soluble part of a biosurfactant maybe as
simple as a carboxylate or hydroxyl function or a complex mixture of phosphate,
amino acids or peptides, anions or cations, or mono-, di- or polysaccharides. Whereas
the lipophilic portions are the hydrocarbon tail that usually made of long chain,
saturated, unsaturated, hydroxyl or α-alkyl-β-hydroxy fatty acids and may contain
cyclic structures [Banat, 1995]. This fatty acid is linked to the hydrophilic group by a
glycosidic, ester or amide bond [Hommel, 1990; Rosenberg and Ron 1999].
Therefore, most of the biosurfactants are lipids, which have the typical amphiphilic
structure of a surfactant.
12
Hydrophobic portion
Hydrophilic portion
Fig 2.3: The amphipathic structure of a surfactant [Cooper, 1986].
Most biosurfactants are either neutral or negatively charged. The negatively
charged is an anionic biosurfactants which due to a carboxylate, phosphate or
sulphate group. Least number of cationic biosurfactants contains amine functions
[Cooper, 1986].
A special property of a biosurfactant is their ability to reduce the surface
tension of water from 72 mNm-1 to below 40 mNm-1. Furthermore, a good
biosurfactant will reduce the surface tension of water to below 35 mNm-1 [Cooper,
1986; Desai and Banat, 1997]. Effective physicochemical properties, which are low
interfacial tensions and critical micelle concentration (CMC) and also temperature
stability, are the characteristics of these compounds. Their heterogeneous group of
surface-active molecules also reduces the CMC and interfacial tension in both
aqueous solutions and hydrocarbon mixtures. These properties create microemulsions
in which micelle formation occurs where hydrocarbons can solubilize in water or
water-in-hydrocarbons [Banat, 1995].
A biosurfactant must have the ability to improve water loss, which can wet
the solid surfaces [Cooper, 1986]. Some of the biosurfactants also has the ability to
act as an emulsifier. Unfortunately, many of the emulsifiers that were characterized
were found to be polymeric, with minimal ability to lower surface tension [Cooper,
1986].
2.2
Screening of Biosurfactant-producing Bacteria
The diverse applications of biosurfactant necessitate an easy, rapid, and
reliable method to screen the bisurfactant-producing bacteria with a minimum
number of false positive and/or negative. Biosurfactant production is always detected
by measuring cell surface hydrophobicity [Pruthi and Cameotra, 1997], drop-
13
collapsing ability [Bodour and Miller-Maier, 1998], hemolytic activity [Yonebayashi
et al., 2000] and their surface activity [Desai and Banat, 1997].
2.2.1
Cell Hydrophobicity Test
Hydrophobicity of the cell surface is an important factor in predicting
bacterial cell adhesion to surfaces. The hydrophobic nature of the outermost surface
of the microbial cells could be used to measure the potential cell affinity to the
hydrophobic substrates. Correlations have been found between the adherence of
bacteria to hydrocarbons and their attachment to other surfaces including nonwettable solid surfaces, epithelial cells, teeth [Rosenberg, 1984] and partitioning of
bacteria at liquid-liquid and liquid-air interfaces [Rosenberg et al., 1980].
Pruthi and Cameotra (1997) found a direct correlation between cell
hydrophobicity and biosurfactant production. Neu and Poralla (1990) used this
property to screen for biosurfactant production based on the fact that hydrophobic
surfaces are usually associated with molecules with low surface energy [Youssef et
al., 2004; Mozes and Rouxhet, 1987]. However, it is not clear which is an
appropriate method in measuring cell surface hydrophobicity for general screening
[Youssef et al., 2004].
2.2.2
Drop-collapsing Technique
Biosurfactant is a microbially produced surface-active agent that contains
both hydrophobic and hydrophilic groups. Due to their amphipathic nature,
surfactants are not uniformly distributed in the solvent but congregate at the solvent
surface [Jain et al., 1991]. Thus, availability of hydrocarbons and slightly soluble
organic compounds can be enhanced by biosurfactants, which can increase aqueous
dispersion by many orders of magnitude and reduce the surface and interfacial
tensions of aqueous medium.
There are two types of intermolecular attractive forces occurred to molecules
liquid [Rao, 1972]. Cohesive forces are referred when those forces occur between
14
like molecules. When this cohesive forces at the surface are strong enough, the
molecules of a water droplet are held together constitute surface tension. When the
attractive forces are between unlike molecules, they are called adhesive forces. Both
of the attractive forces between molecules in a liquid can be viewed as residual
electrostatic forces and this is called Van der Waals forces [Garret, 1972].
A drop-collapsing technique has been defined as a qualitative assay to screen
biosurfactant-producing bacteria. Solutions containing potent biosurfactant will be
unable to form stable drops and spread completely over the oily surface, where as
solutions without surfactant will retain the drop configuration on the oily surface
[Jain et al., 1991]. This method is simple, sensitive, easy to perform, reproducible
and requires little specialized equipment [Bodour and Miller-Maier, 1998]. However,
this technique is not correlated to surface tension reduction to confirm its reliability
[Youssef et al., 2004].
2.2.3
Hemolytic Activity
Hemolysis on blood agar has been widely used to screen biosurfactantproducing bacteria and for preliminary identification of many types of clinically
important bacteria [Mulligan et al., 1984]. Blood agar is purposely used as an
enriched medium for growing of fastidious bacteria and as a differential medium.
This technique was first discovered by Bernheimer and Avigad (1970) that
reported the production of biosurfactant (surfactin) by B. subtilis may cause the red
blood cells to lysis. It has been used previously to quantify surfactin [Moran et al.,
2002] and rhamnolipids [Johnson and Boese-Marrazzo, 1980]. Nowadays, many
researchers have used this technique to screen for biosurfactant production by new
isolates [Carrillo et al., 1996; Yonebayashi et al., 2000].
Hemolytic reactions are generally classified as alpha, beta or gamma
according to the appearance of zones around the isolated colonies growing on blood
agar [Pape et al., 1987]. The hemolytic reaction is called alpha hemolysis when the
colony is surrounded by a zone of intact caused by the decolorisation of which
appear as a greenish zone. This appearance is generally due to the action of peroxide
15
(peroxidases) produced by the bacteria capable of decolorisation [Folman et al.,
2003].
Beta hemolysis indicates a zone of clearing in the blood agar in the area
surrounding a bacterial colony. Few or no intact of erythrocytes are found. One or
more erythrocytes-lysing enzymes (hemolysins) caused this type of hemolysis, which
completely lyse the red blood cells and hemoglobin resulting a complete clear and
white zone around colonies.
If there is no change in the medium around the colony, which is no hemolysis
occurred on the blood agar, the reaction is called gamma hemolysis.
2.2.4
Surface Tension Reduction
Surface tension is a phenomenon involving the cohesive forces between
liquid molecules. The molecules at the surface have no neighboring atoms and adhere
more strongly to those directly associated with them on the surface [Garrett, 1972].
This would enhance the intermolecular attractive forces at the surface makes it more
difficult to move an object through the surface than to move it when it is completely
submersed [Attwood and Florence, 1983].
The phenomenon of surface tension also can be explained in terms of energy.
Surface tension is a measurement of the surface free energy per unit area required to
bring a molecule from the bulk phase to the surface [Rosen, 1978]. The larger the
surface, the more energy there is. Thus to minimize the energy, most fluids assume
the shape with the smallest surface area. This is why, small drops of water are sphere
in shape with minimum surface area for a given volume.
Surface tension can be defined in terms of work, W as follows [Garret, 1972]:
Surface tension, = W/ A
where A is the change in surface area.
It can also be defined as the force, F per unit length, L tending to pull the
surface back [Garret, 1972]:
Surface tension, = F/L
16
Thus, surface tension is a measurement of the intermolecular attractive forces,
which is Van der Waals force in a given liquid. The molecules on the surface of the
liquid experience these forces differently to the air than to the liquid. By introducing
a substrate into the surface, one with a zero degree contact angle with the liquid, all
of the intermolecular forces will pull down on the substrate, thus making the surface
tension directly proportional to the balance force of a balance connected to the
substrate [Tantec Inc., 2002].
The association between surfactants and phases of different polarity like oilwater and air-water, cause reduction in surface tension. One of the factors that can
cause the reduction of surface tension is the presence of microbial surfactants.
Biosurfactant is defined as the one that can reduce the surface and interfacial tension
of aqueous medium. A good biosurfactant producer was defined as one being able to
reduce the surface tension of the growth medium by
20 mN/m compared with
distilled water [Willumsen and Karlson, 1997].
The measurement of surface tension has traditionally been used to detect
biosurfactant production. The du Nouy ring method is the most widely used method
for the measurement of surface and interfacial tension. This method measures the
force required to pull a platinum wire ring through the liquid-air or liquid-liquid
interface. It is widely use because of its accuracy, ease to use and it provides a fairly
rapid measurement of surface and interfacial tension.
2.3
Biosynthesis of Biosurfactant
2.3.1
General Features of Biosynthesis
Generally, biosurfactants are amphiphilic compounds with hydrophilic
moiety consisting of carbohydrate, amino acid, cyclic peptide, alcohol or phosphate,
and hydrophobic moiety may be a long chain fatty acid, a hydroxy fatty acid or αalkyl-β-hydroxy fatty acid. They are synthesized by two primary metabolic
pathways, namely hydrocarbon and carbohydrate pathways [Desai and Banat, 1997].
The metabolic pathways involved in the synthesis of these two groups of precursors
17
are diverse and utilize a specific set of enzymes. Usually, regulatory enzymes are the
first enzymes for the synthesis of these precursors in the biosynthetic pathways.
Syldatk and Wagner (1987) documented the following possibilities could
occur for the synthesis of biosurfactant and their linkage: (1) the hydrophilic and
hydrophobic moieties are synthesized de novo by two independent pathways
followed by their linkage to form a complete biosurfactant molecule, (2) the
hydrophilic moiety is synthesized de novo while the hydrophobic moiety is substratedependent synthesis followed by its linkage, (3) the hydrophobic portion is
synthesized de novo while the synthesis of hydrophilic portion is induced by
substrate, (4) both hydrophilic and hydrophobic moieties have substrate-dependent
synthesis.
Concerning the biosynthesis of glycolipids, the pathway of the sugar-lipid
biosurfactant formation depends on the microorganism. An example for glycolipid
synthesis is the biosynthesis of anionic rhamnolipids by Pseudomonas species.
Rhamnolipid synthesis using enzymology and different radioactively labeled
precursors and the proposed biosynthetic pathway has been studied extensively by
Hauser and Karnovsky (1958) [Banat et al., 2000]. The use of different carbon
source in the medium and the cultivation conditions influence the productivity and
crude product composition, but the chain length of several hydrocarbon substrates
has no effect on the chain length of the fatty acids and sugar moiety of the
glycolipids formed [Syldatk et al., 1985]. Here, both of the hydrophilic and
hydrophobic portions of the rhamnolipid are formed by de novo synthesis. Other
similar observation for the de novo synthesis of biosurfactant is the production of
sophorolipids by resting and growing cells of Torulopsis bombicola from different
lipophilic substrates [Gobbert et al., 1984; Kitamoto et al., 2002].
An example of the second group of biosynthetic pathway is the synthesis of
nonionic trehalose mono- and dicorynomycolates produced by Rhodococcus
erythropolis. While the chain lengths of the hydrophobic portion are dependent on
the hydrocarbon substrates, the sugar moiety is formed by de novo synthesis [Rapp et
al., 1977; Lang and Philp, 1998]. Therefore, the biosynthesis of corynomicolates
does not proceed by de novo synthesis from C-2 units but by chain elongation.
Suzuki et al. (1974) reported the influence of substrate on the sugar moiety of
the glycolipid synthesized by Arthrobacter paraffineus [Kitamoto et al., 2002]. The
18
nonionic trehalose lipid is formed when this bacteria grown on n-alkanes, but
produced fructose lipids when used fructose as the sole carbon source [Itoh and
Suzuki, 1974; Kitamoto et al., 2002]. A similar result has been noted in the resting
cells of Arthrobacter sp. [Li et al., 1984].
The pathways involved in biosynthesis are dependent on the carbon source
and the type of biosurfactant produced [Mulligan and Gibbs, 1993]. A glycolipid
synthesized from a carbohydrate will be regulated by both the lipogenic pathway and
glycolytic metabolism. By these mechanisms, the addition of lipophilic compounds
to the carbohydrate will enhance the production of biosurfactant [Boulton and
Ratledge, 1987].
The biosynthesis of surfactin by B. subtilis has been studied extensively by
Kluge et al. (1988) [Desai and Desai, 1993]. The formation of surfactin occur
nonribosomally, which involved two mechanisms for amino acid activation.
Surfactin synthesizing enzyme, gramicidin S synthetase activate the substrate that
involved in the formation of aminoacyl adenylate and thioester. Ullrich et al. (1991)
reported that enzymatic synthesis of surfactin also requires ATP, Mg2+, sucrose and
precursors. The fatty acid component is incorporated only as an acetyl-CoA
derivative and L-isomer of amino acids are incorporated in the peptide chain. The
enzyme involved also catalyzed the ATP-Pi-exchange reactions, which are mediated
by the amino acid components of surfactin. This pattern was consistent with a
peptide-synthesizing system that activates its substrate simultaneously as aminoacyl
phosphates [Kluge et al., 1988; Desai and Desai, 1993].
2.3.2
Biosynthetic Pathway of Biosurfactant Synthesis
The metabolic pathways involved in the synthesis of the precursors of
biosurfactants are diverse and to some extent dependent on the nature of the principal
carbon source. Sugars are all required either for the synthesis of structural entities of
the cell or for the biosynthesis of amino acids, proteins and nucleic acids. Glucose
may be regarded as the starting point for most microbial fermentations because it is
the most universally used and cheapest carbons source available so far.
19
Formation of dissacharides and polysaccharides found in many biosurfactants
follows the glycolytic pathway prior to sugar modification and transformation [Lang
and Wagner, 1993]. A specific synthetase acts to ensure that the correct reactions
occur. The phosphorylated dissacharide will then be used as the activated sugar for
the formation of the glycolipid biosurfactant.
The biosynthesis of biosurfactant in glucose-grown cells will involve
glycolytic (glucose is degraded) and lipogenic (accumulation of synthesized fatty
acids) metabolisms. In all cases, fatty acid biosynthesis begins with acetyl-CoA as
the key intermediate (Figure 2.4). The differences of the synthesized unsaturated
fatty acids occur mainly because of the organization of the enzymes making up the
individual fatty acid synthetase complexes [Hommel and Ratledge, 1993].
GLUCOSE
75
Trehalose
Glucose-6-P
Surface tension, mN/m
Mannose
Pentose
Pentose phosphate
pathway
Sophorose
Polysaccharides
Fructose-6-P
Glyceraldehyde-3-P
Dihydroxy acetone P
Pyruvate
Malate
Oxaloacetate
Glycerol
Acetyl-CoA
Citrate
Succinate
Malonyl-CoA
Isocitrate
2, Oxoglutarate
Isocitrate
dehydrogenase
LIPIDS
Fatty acids
Fig. 2.4: Metabolic pathway of glucose utilization during biosurfactant production
[Boulton and Ratledge, 1987; Mulligan and Gibbs, 1993].
As shown in Figure 2.4, pyruvate is the end product of glucose metabolism in
glycolysis. If the energy level of the cell is high due to the glucose catabolism, the
flow of carbon into the tricarbocylic acid cycle will be slowed, and acetyl-CoA
accumulated can be diverted into the biosynthesis of fatty acids (lipid). In this case,
20
acetyl-CoA is formed directly in the cytoplasm from pyruvate by pyruvate
dehydrogenase [Hommel and Ratledge, 1993].
(Glucose
) pyruvate + NAD+ + CoA
acetyl-CoA + CO2 + NADH
Acetyl-CoA is carboxylated by acetyl-CoA carboxylase (ACC) to produce
malonyl-CoA, which then becomes the activated C2 donor for the biosynthesis of
fatty acids.
Acetyl-CoA + HCO3 + ATP
malonyl-CoA + ADP + Pi
The elongation of acetyl-CoA to long chain fatty acids is a combination of
various enzymatic reactions. The essential steps are the condensation of an acetyl
group with a malonyl group to yield a C4 unit by -ketoacyl synthase, followed by
reduction, dehydration and further reduction of the C4 unit until a saturated C4
(butyryl) group is formed. The cycle is then repeated by condensation of the butyryl
group with a further malonyl group leading to a C6 moiety. The reaction cycle
continues until a long-chain fatty acyl group is formed.
The overall reaction may therefore be written as [Hommel and Ratledge,
1993]:
7Malonyl-CoA + acetyl-CoA + 14NADPH
Fatty acid synthetases
Palmitoyl-CoA + 7CO2 + 7CoA + 14NADP+ + 7H2O
At this point, the palmitoyl group is then transferred from the protein back to
coenzyme A.
2.3.3 Regulation of Biosurfactant Synthesis
A large variety of biosurfactants are influenced by the nature of the carbon
source, the concentration of nitrogen, phosphorus, magnesium, iron and manganese
ions in the medium, and culture conditions including pH, temperature, agitation and
dilution rate [Abu-Ruwaida et al., 1991; Desai and Banat, 1997]. Biosurfactant-
21
producing bacteria react to changes in their environment by modifying their surface
composition and structure [Angelova and Schmauder, 1999].
The carbon source also important in influencing the biosurfactant synthesis
either by induction or repression [Cameotra and Makkar, 1998]. These have been
reported for the repression of biosurfactant production by Arthrobacter calcoaceticus
[Gobbert et al., 1984] and A. paraffineus [Duvnjak et al., 1982] using organic acids
and D-glucose as carbon source, respectively. In some cases, addition of waterimmiscible substrates results in induction of biosurfactant production. Tulloch et al.
(1962) have found that the induction of sophorolipid synthesis by addition of longchain fatty acids, hydrocarbons or glycerides to the growth medium of Torulopsis
magnoliae [Desai et al., 1994].
Nitrogen also plays an important part to the regulation of biosurfactant
synthesis. Duvnjak et al. (1983) found that urea led to a satisfactory biosurfactant
production [Cameotra and Makkar, 1998]. Moreover, nitrogen limitation also
changed the composition of the biosurfactant production [Syldatk et al., 1985; Desai
and Desai, 1993]. Phosphate limitation also influences the metabolism of
biosurfactant. The change in activity of several intracellular enzymes like alkaline
phosphatase, glucose-6-phosphate dehydrogenase and transhydrogenase dependent
on phosphate levels indicated a shift in biosurfactant metabolism [Mulligan and
Gibbs, 1993].
The limitation of multivalent cations also causes overproduction of
biosurfactants [Guerra-Santos et al., 1984; Cameotra and Makkar, 1998]. Higher
yield of rhamnolipid could be achieved in P. aeruginosa DSM 2659 by limiting the
concentrations of magnesium, calcium, potassium, sodium and trace element salts
[Desai and Desai, 1993]. Finally, the environmental factors and growth conditions
such as temperature, agitation and oxygen availability also affect biosurfactant
production through their effect on cellular growth or activity [Cameotra and Makkar,
1998].
22
2.4
Production of Biosurfactant
2.4.1
Factors Affecting Biosurfactant Production
2.4.1.1 Effect of Carbon Source
Carbon source is very important in the production of biosurfactant. The
carbon sources that had been previously used include carbohydrates, hydrocarbons
and vegetable oils. Some organisms produce biosurfactants only in carbohydrates,
others only in hydrocarbons, and still others consume several substrates, in
combination or separately.
In general, optimal yields are obtained with hydrocarbon or carbohydrate and
lipids. For carbohydrate, most biosurfactant production has been performed using the
more expensive pure forms of the sugar. For example, glucose, fructose and sucrose
lipids are produced by several species of Corynebacterium, Nocardia and
Brevibacterium during growth on the corresponding sugar [Desai et al., 1994]. B.
subtilis also used glucose both in preliminary experiments, while P. aeruginosa used
it in pilot plant studies [Mulligan and Gibbs, 1993]. Water-soluble carbon sources,
such as mannitol, glycerol and ethanol could be used for rhamnolipid production in
Pseudomonas sp., but they are inferior to immiscible substrates like n-alkanes and
olive oil [Desai and Banat, 1997].
The chain length of the hydrocarbon substrate also has affected biosurfactant
production. Optimal production was obtained in Corynebacterium hydrocarboclastus
with linear alkanes of the chain length C-12 to C-14 [Desai et al., 1994], while
Rhodococcus erythropolis produces biosurfactants best on C-12 to C-18 n-alkanes
[Mulligan and Gibbs, 1993].
It has been concluded from a number of studies that different carbon sources
can influence the composition of the biosurfactant formed and how it is produced.
Arthrobacter produces 75% extracellular biosurfactant when grown on acetate or
ethanol but it is totally extracellular when grown on hydrocarbon [Mulligan and
Gibbs, 1993].
23
2.4.1.2 Effect of Nitrogen Source
The nitrogen source in the medium also has a great effect in the production of
biosurfactants. They may also contribute to pH control. Organic nitrogen sources
include gluten meal, yeast hydrolysates and corn germ, whereas inorganic nitrogen
sources include ammonium nitrate, ammonium sulphate, and so on. Among the
inorganic salts tested, ammonium salts and urea were preferred for biosurfactant
production by A. paraffineus, whereas nitrate supported maximum biosurfactant
production in P. aeruginosa [Desai and Banat, 1997].
For surfactin production by B. subtilis, ammonium nitrate was a superior
nitrogen source than ammonium chloride or sodium nitrate. Doubling the ammonium
nitrate from 0.4% to 0.8% increased the surfactin production by a factor of 1.6
[Mulligan and Gibbs, 1993]. Yeast extract was found required for glycolipid
production by Torulopsis bombicola, but was very poor for P. aeruginosa.
Nitrogen limitation not only causes over production of biosurfactants, but also
changes the composition of biosurfactants produced [Syldatk et al., 1985; Desai and
Desai, 1993]. According to Hommel et al. (1987), it is the absolute quantity of
nitrogen and not its relative concentration that is important to give an optimum
biomass yield, while the concentration of hydrophobic carbon source determines the
conversion of carbon available to the biosurfactant.
2.4.1.3 Effect of Environmental Factors
Growth conditions and environmental factors such as temperature, pH,
agitation and oxygen availability also affect the production of biosurfactant.
Temperature may cause alteration in the composition of the biosurfactant produced
by Pseudomonas sp. DSM-2874 [Syldatk et al., 1985]. A thermophilic Bacillus sp.
grew and produced biosurfactant at temperature above 400C [Banat, 1993]. However,
heat treatment of some biosurfactants caused no appreciable change in biosurfactant
properties such as the surface activity as well as the emulsification efficiency [Abu
Ruwaida et al., 1991].
24
The pH of the medium plays an important role in sophorolipid production by
T. bambicola [Gobbert et al., 1984]. Penta- and disaccharide lipid production by
Nocardia corynbacteroides is however unaffected in the pH range of 6.5 to 8.0
[Powalla et al., 1989]. In addition, surface tension and CMC of a biosurfactant
remained stable over a wide range of pH values, whereas emulsification had a
narrower pH range [Abu Ruwaida et al., 1991].
An increase in agitation speed due to the shear effect results in the reduction
of biosurfactant yield produced by Nocardia erythropolis [Margaritis et al., 1979;
Mulligan and Gibbs, 1993]. On the other hand, production of biosurfactant by yeast
increases when the agitation and aeration rates increased [Desai and Banat, 1997].
Depending on its effect on cellular activity, salts concentration also found to
affect the production of biosurfactant. However, some biosurfactants were not
affected by salt concentrations up to 10% (w/v), although slight reductions in the
CMC were detected [Abu Ruwaida et al., 1991].
2.4.2
Kinetics of Biosurfactant
Most of the biosurfactant are secondary metabolite, which are released into
the culture medium at the stationary phase. Some of them are also produced
throughout the exponential phase [Cameotra and Makkar, 1998].
The production of biosurfactant has been carried out in batch or continuous
fermentation at low dilution rates. The kinetics of biosurfactant production exhibit
many variations among various systems (Figure 2.5), (i) growth-associated
production, (ii) production under growth-limiting conditions, (iii) production by
resting or immobilized cells, and (iv) production with precursor supplementation
[Desai and Banat, 1997].
25
(i)
(ii)
(iii)
Growth-associated production
Production under growth-limiting conditions
Production by resting or immobilized cells
Fig. 2.5: Schematic illustration showing different types of fermentation kinetics of
biosurfactant production. (Biomass
, glucose concentration
and
biosurfactant production
).
2.4.2.1 Growth-associated Biosurfactant Production
A parallel relationship between cell growths, substrate utilization and
biosurfactant production exist in growth-associated biosurfactant production. The
production of rhamnolipid by some Pseudomonas sp., surface-active agent by
Bacillus cereus IAF 346 and biodispersan by Bacillus sp. IAF 343 are all examples
of growth-associated biosurfactant production [Desai and Banat, 1997]. The carbon
source plays important role in biosurfactant production [Itoh and Suzuki, 1974]. The
chain length of the hydrocarbon used also affects the production of biosurfactant
[Syldatk et al., 1985].
A mixed growth-associated and non growth-associated process has been
reported occurred in the production of cell-free emulsan by Acinetobacter
calcoaceticus RAG-1. Emulsan-like substance accumulates on the cell surfaces
during the exponential growth phase and is released into the medium when protein
synthesis decreases [Goldman et al., 1982]. Wang and Wang (1990) performed
extensive studies on the mechanism of biosurfactant accumulation in A.
calcoaceticus RAG-1. They revealed that the ratio of cell-bound polymer to dry cell
26
is strongly affected by shear force and as the shear stress increases, the ratio
decreases.
2.4.2.2 Biosurfactant Production Under Growth-limiting Conditions
The unique feature of this category of biosurfactant production is the sharp
increase in the biosurfactant level as a result of limitation of one or more medium
components. In some cases, an overproduction of biosurfactants was dependent on
growth-limiting conditions such as N-limitation or limitation of the multivalent
cations, so that a rhamnolipid production by P. aeruginosa occurred only after
reaching the stationary growth phase [Suzuki et al., 1974; Guerra-Santos et al.,
1986].
Powalla et al. (1989) has demonstrated that the production of pentasaccharide
lipid by Nocardia corynebacteriods SM-1 is favored by inorganic nitrogen sources,
and sodium nitrate gave maximum surfactant production. Furthermore, it has been
observed that during growth, the initial yield of glycolipid increases rapidly after the
exhaustion of the nitrogen source and after attaining the stationary phase of growth.
Hommel et al. (1987) have studied extensively the production of watersoluble biosurfactant by Torulopsis apicola. According to them, the absolute quantity
of nitrogen and not its relative concentration important to determine the optimum
concentration of biomass, whereas the concentration of the hydrophobic carbon
source determines the conversion of available carbon into biosurfactants. Moreover,
it was shown that the C:N ratio in the medium also plays an important role in
biosurfactant production, and a large amount of n-hexadecane is found to be
incorporated into the surfactant at higher C:N ratio.
Iron concentration has shown to have a dramatic effect on rhamnolipid
production by P. aeruginosa, which resulting in a threefold increase in the
production when cells were shifted from medium containing 36µM iron to medium
containing 18µM iron. However, there was no change in the biomass yield under
these conditions [Guerra-Santos et al., 1986].
27
Syldatk and Wagner (1987) stated that the effect of N-limitation or a
limitation of the multivalent cations is nonspecific and it is expressed as a change of
the physiological state of the microorganisms used for the production of
biosurfactant.
2.4.2.3 Biosurfactant Production by Resting or Immobilized Cells
In this category of biosurfactant production, cells do not multiply but
continue to utilize carbon source for the synthesis of biosurfactants. The cells used
are harvested from the surfactant-producing state of culture broth and maintained in
the same state. The wet biomass is washed and used for the production of
biosurfactant under specific conditions, so that the effect of possibly disturbing
products can be eliminated and the influence of single factors on the synthesis of the
compound, such as pH, temperature and salt concentrations can be examined.
Production of rhamnolipid by resting cells could be increased evidently in
comparison with growing cells under N-limitation [Syldatk and Wagner, 1987]. In
contrast to the rhamnolipid production by growing cells, two new rhamnolipids, R3
and R4 were synthesized by the resting cells [Syldatk et al., 1985]. The production of
these biosurfactants was dependent on the incubation temperature and the carbon
source used in the medium. By incubating resting cells in phosphate buffer, repeated
use of cells for rhamnolipid production is increased by almost 5 to 6-fold. It is
proposed that the effect may be due to relieving the product inhibition. However,
biosurfactant production rate was much lower as compared to that with growing
cells.
Production of biosurfactant by resting cells also has been observed in the
production of sophorolipid by T. bombicola [Inoue and Itoh, 1982] and trehalose
tetraester production by R. erythropolis [Syldatk et al., 1985]. In this case, the
conversion rate of substrate to product was found to be much higher than that
observed with growing cell under nitrogen limitation.
The production of biosurfactant by resting cells is important for the reduction
of cost of product recovery, as in such cases the growth phase and the product
formation phases are separated.
28
2.4.2.4 Biosurfactant Production in Addition to Precursor
Many reports have showed that the addition of biosurfactant precursors to the
growth medium causes both qualitative and quantitative changes in the product.
Addition of lipophilic compounds to the medium of T. apicola IMET 43747 [Stuwer
et al., 1987] and T. bombicola [Cooper and Paddock, 1984] resulted in the higher
production of biosurfactants. In this case, the carbon source in the medium,
particularly the carbohydrate, has great bearing on the type of glycolipid formed.
Similarly, increased production of biosurfactants containing different mono-, di-, or
trisaccharides was reported occurred in Corynebacterium sp. and Nocardia sp.
through supplementation of the corresponding sugar in the culture medium [Itoh and
Suzuki, 1974].
This method of biosurfactant production will probably be of great interest in
future because it allows the production of new surface- and interfacially active
compounds whereby the chemical and physical properties of these compounds can be
influenced by the carbon sources used for biosurfactant formation [Syldatk and
Wagner, 1987].
2.5
Extraction of Biosurfactants
The recovery and purification of biosurfactants from complex fermentation
broth is a major problem in the commercialization of biosurfactants. In many cases,
the downstream process increases the cost of biosurfactant production to as high as
60% [Desai et al., 1994; Desai and Banat, 1997]. Thus, improving product yield, low
material costs and combining the steps of recovery can reduce the recovery costs.
Economically biosurfactant recovery processes are mainly depending on its
ionic charge, water solubility and its nature location (intracellular, extracellular or
cell bound) [Desai et al., 1994]. Most biosurfactants are secreted into the medium
and they are isolated from either culture filtrate or supernatant obtained after removal
of cells. The commonly reported techniques or biosurfactant recovery are listed in
Table 2.2.
29
Table 2.2: Common methods employed for the recovery of biosurfactants
PROCESS
•
•
•
•
•
Batch Process
Solvent extraction
- Sophorolipid, trehalolipid
Acid precipitation
- Surfactin
Acetone precipitation
- Bioemulsifier
Crystallization
- Glycolipid, cellobiolipid
Ammonium sulphate precipitation
- Emulsan. biodispersan
•
•
•
•
•
Continuous Process
Centrifugation
- Glycolipid
Adsorption
- Rhamnolipid, lipopeptide
Diafiltration and precipitation
- Glycolipid
Ultrafiltration
- Glycolipid, lipopeptide
Foam separation and precipitation
- Surfactin
REFERENCES
Suzuki et al., (1974); Ristau and Wagner,
(1993).
Arima et al., (1968); Javaheri et al.,
(1985).
Chameotra and Singh, (1990).
Oberbremer and Mullar-Hurtig, (1989);
Spencer et al., (1979).
Rosenberg et al., 1979; Rosenberg et al.,
(1988).
Kitamoto et al., (1993).
Yamaguchi et al., (1976); Matsuyama et
al., (1991).
Chametra and Singh, (1990).
Mulligan and Gibbs, (1990); Lin and
Jiang, (1997).
Davis et al., (2001).
Classical recovery methods are well suited to batch fermentation. Settling,
flotation, centrifugation or rotary vacuum filtration is used in this technique. Settling
and floatation are not feasible for bacterial cells though it is the least expensive
techniques. Centrifugation is effective, but it requires a high maintenance costs and
heat generation during centrifugation, which may damage the product. Furthermore,
the complexity of recovery equipment increases as the initial product concentration
decreases. More additional steps may be required in this case [Mulligan and Gibbs,
1993].
The most widely used technique is solvent extraction with a variety of
solvents at several different ratios. The choice is dependent on cost and effectiveness.
Solvents used for this purpose include chloroform-methanol mixture,
dichloromethane, ethyl acetate, acetic acid, ether, etc. Recently, methyl tertiary-butyl
ether was able to extract crude surfactant material produced by Rhodococcus rubber
IEGM 231 with high product recovery and good functional surfactant characteristics
30
[Kuyukina et al., 2001]. However, the use of solvents is time consuming, expensive
and not very specific. Further purification must be done by column chromatography,
thin-layer chromatography or crystallization [Mulligan and Gibbs, 1993].
Stuwer et al. (1987) described an easy and cheaper purification process using
liquid chromatography on silica gel for the nonionic glycolipid produced by T.
apicola. Mannosylerythritol lipids produced by Candida sp. are settled down as
heavy oils by centrifugation [Kitamoto et al., 1993]. Bryant (1990) demonstrated an
improved method for the isolation of glycolipid from Rhodococcus sp. H13A by
using XM-50 diafiltration and isopropanol techniques. These techniques give a purer
glycolipid and removes protein impurities.
In a recent development, continuous removal of biosurfactant during
fermentation by different techniques has increased the cell density in the reactor and
eliminated product inhibition which resulting in a several fold net increase in
biosurfactant yield [Desai and Banat, 1997]. In addition, substantial reductions in the
cost of product recovery and effluent treatment were achieved.
The technique of foam fractionation has gained greater significance as it
offers an advantage of continuous in-situ removal of biosurfactant from the
fermentation broth. In the recovery of surfactin produced by B. subtilis, foam is
collected and the pH of the collapsed foam is adjusted to 2 with concentrated HCl.
Proteins and lipids are precipitated and settled down in this process. The supernatant
is decanted off and surfactin is extracted in dichloromethane from the residues [Davis
et al., 2001].
2.6
Applications and Roles of Biosurfactant
In recent years, microbial surface-active agents are required in a very large
number of diverse applications due to their broad range of functional properties.
There is no industry, which does not have some use for these compounds [Cooper,
1986; Desai and Desai, 1993]. They are potentially useful in every industry dealing
with multiphase systems, due to their basic structure that contain both hydrophilic
and lipophilic portions [Desai and Banat, 1997].
31
Petroleum industry is one of the largest markets for biosurfactant. They are
used in petroleum production and incorporation into oil formulations as to enhance
oil recovery [Van Dyke et al., 1991]. This required the solubilization of hydrophobic
pollutants found in petroleum hydrocarbons before being degraded by microbial
cells. Surface area of hydrophobic materials will increase, thus increasing their water
solubility. Hence, the presence of surfactants may increase microbial degradation of
pollutants in both soil and water [Van Dyke et al., 1991].
Various researches have studied the effect of biosurfactant on biodegradation
of organic contaminants. Biosurfactants responsible to enhance solubility of the
substrate for the microbial cells and interaction with the cell surface, which increases
the hydrophobicity of the surface allowing hydrophobic substrates to associate more
easily [Shreve et al., 1995; Mulligan, 2004].
Emulsan, the patented, commercialized biosurfactant have the potential
applications in the cleaning of oil contaminated vessels, oil spills and in microbial
enhance oil recovery (MEOR) [Desai et al., 1994]. The emulsification of heavy crude
oil and reduction in the viscosity of crude oil from 2,000,000 to 100 centipoise had
been done by emulsan.
In MEOR, biosurfactants tend to increase oil mobility by reducing the
interfacial tension at the oil-rock interface. This reduces the capillary forces
preventing oil from moving through rock pores. Biosurfactant also aid the oil
emulsification and assist in the detachment of oil films from rocks [Banat et al.,
2000].
Emulsion polymerization for paints, paper coatings and industrial coatings
was identified as the second largest market for surfactants [Van Dyke et al., 1991].,
A polymeric biosurfactant called biodispersan produced by A. calcoaceticus A2 has
potential use in paint industries [Rosenberg and Ron, 1999]. The suspension made in
the presence of this compound is easy to handle, as particles settle very slow. This is
an important characteristic for paint, because it gives better spreadibility and
improved the properties of mixing.
Biosurfactants have attracted personal care industries as emulsion stabilizer
because of their low toxicity, excellent moisturizing properties and skin
compatibility. A product containing 1 mole of sophorolipid and 12 moles of
32
propylene glycol has specific compatibility to the skin and found commercial utility
as skin moisturizer [Yamane, 1987; Banat et al., 2000].
Biosurfactants use in the food industry always acts as emulsifiers for the
processing of raw materials. Emulsification plays an important role in forming the
right consistency and texture as well as in phase dispersion [Banat et al., 2000].
Busscher et al. (1996) found that thermophilic dairy Streptococcus spp. produced a
biosurfactant that can be used for fouling control of heat exchanger plates in
pasteurizers, as they retard the colonization of Streptococcus thermophilus
responsible for fouling.
In agricultural industries, biosurfactants are always used to enhance
penetration of active compounds into plants [Kosaric et al., 1987]. Biosurfactants are
needed for the hydrophilization of heavy soils to obtain good wettability and to
achieve equal distribution of fertilizers and pesticides in the soils [Banat et al., 2000].
It also useful in formulating poorly soluble organophosphorus pesticides.
The ability of the rhamnolipid mixture to solubilize the pesticide has been
studied extensively by Mata-Sandoval et al. (2000). The biosurfactant seems to bind
pesticide tightly in the micelle and release the pesticide slowly to the aqueous phase,
which could have implications for microbial uptake [Mulligan, 2004].
Recently, the applications of biosurfactant in the field of biomedical science
have been studied extensively. Iturin A, a potent antifungal lipopeptide biosurfactant
was found to increase the electrical conductance of biomolecular lipid membranes,
which has stimulated discussion on the pore-forming activity of lipopeptides and
their action against pathogen [Singh and Cameotra, 2004].
Table 2.3: Some properties of biosurfactant commonly used in several applications.
Types of
biosurfactant
Rhamnolipid
Sophorolipid
Emulsan
Surfactin
Applications
Biodegradation of
organic contaminant
Skin moisturizer for
cosmetic
Emulsification of
heavy crude oil
Inhibition of fibrin
clot formation
Properties
High solubility and
bioavailability
Excellent skin
compatibility
Stable o/w
emulsion
Antimicrobial
activity
References
Mulligan, 2004
Banat et al.,
2000
Desai et al.,
1994
Banat et al.,
2000
33
Lipopeptide
MEOR
Rhamnolipid
Heavy metal removal
Mineral flotation in
pharmaceutical
industries
Mannosylerythritol Anti-agglomeration
agent
lipid
Trehalose lipid
2.7
Thermotolerant and
high stability over a
wide range of pH
Complexation
foaming and ability
Chemically stable
and high surface
activity
High surface
activity and low
CMC
Banat et al.,
2000
Mulligan, 2004
Mulligan and
Gibbs, 1993
Lang, 2002
Characteristics of Biosurfactant and Chemical Surfactant
Biosurfactant and chemically-synthesized surfactant were differentiated
according to their classification, type of substrate, rate of toxicity and
biodegradability, production cost and effectiveness at particular temperature, pH and
salinity. Table 2.4 summarized the differences between chemically-synthesized
surfactant and biosurfactant.
Table 2.4: Differences between biosurfactant and synthetic surfactant.
Synthetic surfactant
Categorized according to
the nature of their polar
group
Specific for certain
temperature, pH and
salinity
High toxicity
Low biodegradability and
foaming ability
No strain needed
Chemically synthesized
mostly from petroleum
resources
Specific for particular
industrial application
Low production cost
Biosurfactant
Categorized by their
chemical composition and
their microbial origin
High selectivity and
specific activity at extreme
temperature, pH and
salinity
Low toxicity
High biodegradability and
foaming ability
Poor strain productivity
Ability to synthesize from
renewable feed-stock
Broad spectrum of
industrial application
High production cost
References
Desai and Banat, 1997
Banat et al., 2000
Edwards et al., 2003
Banat, 1995
Cameotra and Makkar,
1998
Fox and Bala, 2000
Cameotra and Makkar,
1998
Banat, 1995
34
2.7.1 Advantages and Disadvantages of Biosurfactants in Commercial
Application
Biosurfactants has several advantages compared to the chemical surfactants,
such as broad range of structural and physical properties, lower toxicity, higher
biodegradability, better environmental compatibility, higher forming, high
selectivity, able to be synthesized from renewable feed-stocks and have specific
activity at extreme temperature, pH and salinity [Desai and Banat, 1997]. They can
be degraded by microorganisms to produce novel compounds, which are more
effective for specific purposes [Van Dyke et al., 1991]. They also have the ability to
be tailored by genetic engineering, physiological and biochemical techniques to meet
the specific requirements in industries.
The biosurfactant was non-toxic compounds which application would results in
the removal of oil pollutant more effectively compared to the synthetic surfactants
which was highly toxic to marine organisms and environment [Van Dyke et al.,
1991]. With their property of environmental compatibility, biosurfactants have wide
environmental applications such as bioremediation and oil recovery.
Biosurfactants can be produced using low-cost substrates. However, a low rate
product yield and purification procedures can result in higher prices than for
chemical surfactants. The high cost of toxicity testing and the time required having a
new compound approved for use further increases in the cost of biosurfactants in
market [Kosaric et al., 1987]. It is clear that improving the method of biosurfactant
production and the development of strain selection techniques are highly important in
the commercial application of biosurfactant.
Mulligan and Gibbs (1993) have proposed the strategies that can be made in
order to reduce the costs for all aspects of biosurfactant production. These would
involve the choice of inexpensive raw materials, increasing biosurfactant yield and
production rate by biosynthesis control, screening for overproducers and genetic
manipulation of biosurfactant producers, optimization of the fermentation process,
reduction of product recovery costs and finally, production of biosurfactants that
suitable for specified applications.
CHAPTER 3
GENERAL MATERIALS AND METHODS
The following sections in this chapter describe all the materials and methods
that were used routinely in the study. All chemicals used were supplied by either
Merck-BDH Laboratory Supplier or Sigma Chemicals Ltd., unless stated otherwise,
and were, where possible of AnalaR grade.
3.1
Microorganisms
The mesophilic bacteria involved in this study were isolated from petroleumrelated industries in Malaysia (Table 3.1). Those bacterial isolates include: (i)
possible non-hydrocarbon-degrading bacteria isolated from petrochemical waste
samples; and (ii) possible hydrocarbon-degrading bacteria isolated from oil samples.
Details of isolation for these mesophilic bacteria are shown in Section 3.1.1. They
were screened for the best biosurfactant-producing ability via several methods
(Section 4.2.1). The selected bacterial isolates were used for further study.
3.1.1
Bacterial Isolates: Origin and Route of Isolation
Samples of wastewater and oil were collected at various points within the
Titan Petrochemical (M) Sdn. Bhd. treatment plant, Pasir Gudang, Johor and the
Exxon Mobil Oil Refinery, Port Dickson, Negeri Sembilan, as part of research
36
exercise during March, 2002. Most of the sites sampled were within mesophilic
temperature (25ºC-40ºC); these areas were mapped and major characteristic of sites
were determined in situ. Over ten sites were sampled in all, though only five of these
are referred to in this report (Table 3.1).
Samples were subjected to enrichment in liquid medium (Section 3.2.1.1) and
then streaked onto solid medium (Section 3.2.2.1 and 3.2.2.2). For the isolation of
mesophilic bacteria, liquid enrichment culture and solid medium were incubated at
30ºC and 37ºC, respectively for up to 2 days. Plates were examined and preliminary
identification of isolates made on the basis of colony morphologies and cell
characteristics [Gerhardt et al., 1994]. Isolates were purified by repeated single
colony isolation and purity of cultures checked periodically by streaking liquid
cultures onto Ramsay agar (Section 3.2.2.2).
3.1.2
Crude oil
The crude oil samples used in this study was obtained from The Exxon Mobile
Oil Refinery, Port Dickson, Negeri Sembilan. The sample was autoclaved (121ºC,
101.3kPa for 15 minutes) separately in bottles before being added aseptically to the
growth medium (Section 3.2.1).
37
Table 3.1: Origin of bacteria isolated from petroleum-related industries.
Isolates
Site of Origin and Characteristics
MFTA-W1
FTA influent
(Titan Petrochemical (M) Sdn. Bhd., Johor)
•
Temperature: 25-30ºC
•
pH 5-6
Aeration basin wastewater
(Titan Petrochemical (M) Sdn. Bhd., Johor)
•
Temperature: 25-30ºC
•
pH 7
Aeration basin wastewater
(Titan Petrochemical (M) Sdn. Bhd., Johor)
•
Temperature: 25-30ºC
•
pH 7
Activated sludge
(Titan Petrochemical (M) Sdn. Bhd., Johor)
•
Temperature: 28-34ºC
•
pH 6.5-6.8
FTA influent
(Titan Petrochemical (M) Sdn. Bhd., Johor)
•
Temperature: 25-30ºC
•
pH 5-6
Soil-sludge farm
(Exxon Mobil Oil Refinery, Negeri Sembilan)
•
O:P:N = 100:10:1
•
pH 4 (adjusted to 5-6 with CaCO3 )
•
No lining and seeding
Biological treatment lagoon effluent
(Exxon Mobil Oil Refinery, Negeri Sembilan)
•
Temperature: 30ºC
•
pH 7.6
•
1-2 ppm oil content
•
BOD: 30-40 ppm
Biological treatment lagoon effluent
(Exxon Mobil Oil Refinery, Negeri Sembilan)
•
Temperature: 30ºC
•
pH 7.6
•
1-2 ppm oil content
•
BOD: 30-40 ppm
Biological treatment lagoon effluent
(Exxon Mobil Oil Refinery, Negeri Sembilan)
•
Temperature: 30ºC
•
pH 7.6
•
1-2 ppm oil content
•
BOD: 30-40 ppm
Biological treatment lagoon effluent
(Exxon Mobil Oil Refinery, Negeri Sembilan)
•
Temperature: 30ºC
•
pH 7.6
•
1-2 ppm oil content
•
BOD: 30-40 ppm
AB-Cr1
MAB-Cr1
RAS-Cr2
RFTA-Cr3
RSSF-Cr1
ETL-Cr1
ETL-Cr7
RETL-Cr1
RETL-Cr3
38
3.2
Media Preparation
3.2.1
Liquid Medium
3.2.1.1 Ramsay Liquid Medium
Ramsay medium [Ramsay et al., 1983] was used for growth of biosurfactantproducing isolates consisted of (per litre): 2.0g NH4NO3, 0.5g KH2PO4, 1.0g
K2HPO4, 0.5g MgSO4.7H2O, 0.01g CaCl2.2H2O, 0.1g KCl and 0.06g yeast extract.
The pH was adjusted to 6.5-6.8 before autoclaving at 121ºC and 101.3kPa for 15
minutes.
A 3 to 10mM glucose was added into the medium prior to inoculation.
Glucose stock solution (1M) was prepared by adding 49.54g glucose in 250mL
distilled water and was filter sterilized using 0.2µm nylon membrane.
3.2.1
Solid Media
3.2.2.1 Nutrient Agar
Nutrient Agar (NA) was used for growth and maintenance of isolated bacteria
from the petrochemical wastes. NA (2% w/v) was suspended in 1000mL distilled
water before autoclaving at 121ºC and 101.3kPa for 15 minutes. The medium was
then cooled to approximately 50ºC prior to pour (~20mL) into sterile Petri dishes.
The molten agar was left to cool and gel at room temperature. The medium can be
used directly following preparation or stored in room temperature for up to one week.
3.2.2.2 Ramsay Agar
The preparation of Ramsay agar was similar to that of the Ramsay liquid
medium (Section 3.2.1.1), except that agar (2% w/v) was added as gelling agent.
39
3.2.2.3 Blood Agar
Blood agar was prepared according to method described by Benson (1994)
using 4% (w/v) tripticase soy agar as gelling agent. Final pH was adjusted to 7.3
before autoclaving at 121ºC and 101.3kPa for 15 minutes. The medium was cooled to
approximately 50ºC and 5% (v/v) of fresh human blood sample obtained from Pusat
Kesihatan UTM Skudai, Johor was added to the medium. The mixture was gently
swirled to ensure thorough mixing and dispensed (~20mL) into sterile Petri plates.
The medium was left to gel at room temperature prior to incubation at 37ºC for 24
hours to check for contamination.
3.3
Growth and Maintenance of Bacterial Isolates
3.3.1
Inoculum Preparation
A fresh single pure colony of each bacterial isolates was transferred
aseptically from agar plate into Ramsay liquid medium using a sterile wire loop. The
inoculated medium was then incubated at either 37ºC or 45ºC at 200rpm in orbital
shaker (Certomat@U- B.Braun Biotech International) until the culture reached an
optical density (OD600) of between 0.5 to 0.8 prior to use as inoculum.
3.3.2
Culture Maintenance and Storage
All pure isolates were maintained in liquid (Section 3.2.1) and solid media
(Section 3.2.2). They were regularly subcultured into fresh medium for short-term
storage. Stock cultures of all pure isolates were prepared in a ‘protect mixed bacterial
preserver beads’ at –80ºC, according to manufacture instructions for a long-term
maintenance.
40
3.4
Analytical Methods
3.4.1
Determination of Bacterial Biomass
3.4.1.1 Optical Density
Bacterial biomass was determined by measuring the culture optical densities
at 600nm (OD600) using Jenway 6300 Spectrophotometer. Optical densities of the
samples removed from cultures were read against blank of distilled water. It was
also observed that the OD of the uninoculated Ramsay medium used for growth and
biosurfactant production and the OD of distilled water was similar. Therefore
distilled water was used as blank throughout this study for growth determination.
3.4.1.2 Cell Dry Weight
The bacterial cell dry weight was determined as a function of OD600,
following an assumption: 0.1 OD equivalents to 1.0mg/mL cell biomass. This
relationship was obtained from the standard growth curved plotted between OD600
versus cell biomass (g/l) as indicated in Appendix A.
3.4.2
Determination of Glucose Concentrations
Soluble glucose concentrations in the medium were determined by enzymatic
reactions, using a glucose analytical kit (Sigma, USA). Combined Enzyme Color
(CEC) reagent was prepared by adding 1 cap of PGO enzyme and 1.6mL o-ddihydrochloride into 100mL distilled water. Then, 0.2mL sample obtained by culture
supernatant after centrifugation at 5000rpm at 4ºC for 20 minutes (Hettich
Zentrifugen Universal 32) was added with 2mL CEC reagent before it was incubated
in dark at 37ºC for 30 minutes. Glucose standard solution was used to generate a
standard curve. Optical densities of the mixture were read against blank of glucose
standard solution at 450nm.
41
3.4.3
Surface Activity Measurements
3.4.3.1 Surface Tension Measurement
Surface tension of the supernatant was measured using a semi-automatic
Surface Tensiometer, model ST-Plus (Tantec Inc. Schaumburg, R) via Wilhelmy
Detach method. This method measure the surface force between a liquid and air in a
liquid medium samples. The surface tension of Ramsay medium at 25ºC was used as
control.
The Wilhelmey Detach method utilizes a Platinum-Iridium rectangular plate.
The plate was cleaned with 99% ethanol and heated with the alcohol lamp until all
residues was removed. All glass equipment used was acid washed (HCl, 0.1M) to
ensure that they are clean to get an accurate measurement.
A 20mL volume of sample obtained by culture supernatant after
centrifugation at 5000rpm at 4ºC for 20 minutes was put into a clean 50mL glass
beaker and placed onto the tensiometer platform. The rectangular plate was wetted
and the measurement was carried out by press the MEAS button to ensure that the
contact angle between the plate and the liquid is zero. The tensiometer platform was
then lowered at a constant rate until the meniscus breaks. The surface tension of each
sample will automatically detect and the measurement was repeated three times,
performed at room temperature.
3.4.3.2 Interfacial Tension Measurement
The ST-Plus semi-automatic surface tensiometer was used to measure the
interfacial tension of culture supernatant against crude oil by using the Wilhelmy
Detach method.
First, surfactant sample was put in a 50mL glass beaker to a depth of
approximately 7mm and crude oil was put on top of the sample to a depth of
approximately 13mm. The plate was then wetted in the sample (in other glass beaker)
42
before it was hanged on the tensiometer. The tensiometer platform was raised slowly
till the plate makes contact with the sample, and then lowered the platform at a
constant rate until the meniscus breaks. The value showed in tensiometer screen is
the value of interfacial tension of the sample against crude oil.
Both surface and interfacial tension values were in mN/m unit and the value
of buoyancy effect shown after the zeroing the contact angle between the plate and
the liquid must be calculated following to equation [Tantec ST-Plus]:
σactual solution = σ*measurement value – value of buoyancy effect
3.4.3.3 Spreading Tension Measurement
Spreading tension of surfactant solution on oil was calculated according to the
following equation [Ramsay et al., 1983]:
γsp = γst - γoil - γift
γsp : spreading tension
γoil : surface tension of the oil
γst : surface tension of the surfactant solution
γift : interfacial tension of the oil-surfactant solution
Since only one type of oil was used (crude oil), the surface tension of the oil was
constant. The more lower the surface tension due to the increasing concentration of
surfactant, the lower spreading tension of the surfactant solution-on-oil, and this
indicated to the increasing tendency of a surfactant to produce stable emulsion.
3.5
Production of Biosurfactant
3.5.1
Biosurfactant Extraction
Biosurfactant was extracted from the whole cell-free culture broth. The
bacterial cells were removed by centrifugation at 5000rpm, 4ºC for 30 minutes. The
43
supernatant was adjusted to pH 2 using sulphuric acid, H2SO4 (1M) prior to
biosurfactant extraction using equal volume of chloroform-methanol (2:1). The
mixture was shaken for 3 hours at 30ºC and 200rpm. The biosurfactants were then
concentrated using rotary evaporator at 60-70ºC with rotor at 40% (Buchi B-169
vacuum system).
3.5.2
Determination of Biosurfactant Dry Weight
The light yellowish product obtained after the extraction procedures (Section
3.5.1) was then dried at 60ºC to a constant weight prior to get the yield of
biosurfactant production.
CHAPTER 4
SCREENING AND CHARACTERIZATION OF BIOSURFACTANTPRODUCING BACTERIA
4.1
Introduction
This chapter describes various methods for the screening of biosurfactantproducing bacteria which include bacterial adherence to hydrocarbon test, drop
collapsing technique, hemolytic activity and surface tension reduction. Surface
tension measurement was reported as a primary method used as reference to indicate
the ability of microbes to produce biosurfactant, though other methods were also
used as comparison for a better selection of biosurfactant producer [Willumsen and
Karlson, 1997]. Youssef et al. (2004) had found that there was a strong correlation
coefficient between the surface tension reduction and the drop collapse method.
However, blood hemolysis test was found correlate only to drop collapse method but
not to surface tension. Though, this was not the drawbacks of blood hemolysis
technique for the screening of biosurfactant-producing microbes as large number of
successes have been reported using this method [Bodour and Miller-Maier, 2002].
Thus, various screening methods were used in this study in order to obtain a better
selection for the most potential bacteria that capable of producing biosurfactant
(Section 4.2.1). The selected biosurfactant-producing isolates were then characterized
morphologically and biochemically (Section 4.2.2).
45
4.2
Methodology
4.2.1
Screening of Biosurfactant-producing Bacteria
4.2.1.1 Bacterial Adherence To Hydrocarbon (BATH) Test
This technique was carried out using a method described by Rosenberg et al.
(1980), based on the degree of cell adherence to liquid hydrocarbon following a brief
period of mixing. Ten bacterial isolates were grown for 48 hours at 37ºC with
shaking in Ramsay medium (10mL) added with 5mM glucose as carbon source.
Bacteria (8mL) were then harvested, washed twice with 4mL PUM Buffer (pH 7.1)
containing 16.9g of K2HPO4, 7.3g of KH2PO4, 1.8g of urea and 0.2g of MgSO4.7H2O
dissolved in one litre distilled water. The cells were then resuspended in the same
buffer (8mL) prior to measure the initial density of the cell suspension (ODc)
spectrophotometrically at the wavelength of 400nm. The bacterial cell suspension
(8mL) were then mixed with hexadecane (2mL) in a tissue culture tubes (15 x 2.5
cm) and incubate at room temperature for 10 minutes prior to vigorous mixing by
vortex for about 2 minutes. After vortexing, the mixture was left undisturbed for 15
minutes to allow separation of hexadecane from aqueous phase. The aqueous phase
(bottom layer) was then carefully removed and the cell density remained in the
aqueous phase (ODa) was measured spectrophotometrically at 400nm.
Hydrophobicity was expressed as the percentage of cell adhered to
hydrocarbon, which was calculated as follows: 100(1-ODa/ODc).
4.2.1.2 Drop-collapsing Test
The drop-collapse technique was performed on clean glass slide following
method described by Bodour and Miller-Maier (1998). Ten isolates were grown in
Ramsay medium (10mL) with 5mM glucose as carbon source, incubated with
shaking for 48 hours at 37ºC and 200 rpm. Each of the glass slides used was rinsed
with hot water, ethanol and distilled water, and dried. The slides were then coated
46
with 1.8µL of Penzoil 10W-40 and equilibrated for 24 hours to ensure a uniform oil
coating. Penzoil 10W-40 was used in this test because it gives the best qualitative
indication of the presence of surfactant compared to other oils such as mineral oil,
kerosene, hexadecane, Castrol 10W-30 and silicone oil. Penzoil 10W-40 was
considered the most effective oil because either the water drop collapsed completely
in the presence of surfactant or it remained beaded in the absence of surfactant.
A 5µL aliquot of sample was then applied onto the center of the oil drops
using 10µL micropippetor by holding the pipet at an angle of 45º. The results were
monitored visually after 1 hour. If the drop remained beaded, the result was scored as
negative. If the drop collapsed, the result was scored as positive.
4.2.1.3 Blood Hemolysis Test
Fresh cultures from ten bacterial isolates were prepared by streaking on
Nutrient Agar and incubate at 37ºC for 24 hours. The fresh single colony of cultures
was then restreaks on Blood Agar respectively and incubates at 37ºC for 48-72 hours.
The bacterial colonies were then observed for the presence of clear zone of hemolysis
around the colonies on Blood Agar.
Result was recorded based on the type of clear zone observed i.e α-hemolysis
when the colony was surrounded by greenish zone, β-hemolysis when the colony was
surrounded by a clear white zone and γ-hemolysis when there was no change in the
medium surrounding the colony.
4.2.1.4 Surface Tension Measurement
Surface tension reduction of the culture medium was measured using a semiautomatic Surface Tensiometer, model ST-Plus (Tantec Inc. Schaumburg, R) as
described in Section 3.4.3.1.
47
4.2.2
Characterization of Biosurfactant-producing Isolates
The selected biosurfactant-producing bacteria coded AB-Cr1 and ETL-Cr1
were characterized morphologically (Section 4.2.2.1) and biochemically (Section
4.2.2.2). Cultures of less than 24 hours were used to ensure consistency and validity
of results obtained from biochemical tests.
4.2.2.1 Morphological Analysis
Morphological characterization of microbes was commonly performed to
distinguish the microbes based on colony and cellular morphologies. A stereo scan
microscope (Leica, Germany) was used to examine and characterize the formation of
bacterial colonies on solid media, using magnification of x40 to x50.
Phase-contrast microscopy allowed the visualization of colourless, small
specimens, which do not absorb enough light to be seen by bright-field microscopy.
A Leica (Germany) phase-contrast microscope, lifted with a zenike condenser and
objective (magnification x800), was used to record morphological and behavioral
characteristics of bacterial cells.
4.2.2.2 Biochemical Analysis
Biochemical tests were usually done to determine the genus of unknown
bacterial species. The selected strains that had been screened as the best biosurfactant
producers among the 10 isolates were subjected to further biochemical
characterization. The biochemical tests performed include gram stain, oxidase,
catalase, motility, citrate, urease, oxidation/fermentation, triple sugar ion, nitrate
reduction, indole and gelatin liquefaction tests, according to standard procedure
[MacFaddin, 1980] described in Appendix A. Results from the biochemical analysis
were used to find the closest match with known bacterial genus and to assign the
bacterial signature according to Bergey’s manual [Holt et al., 1994].
48
4.3
Results and Discussion
4.3.1
Screening of Biosurfactant-producing Bacteria
Results of screening for biosurfactant-producing bacteria were presented and
discussed as follow (Section 4.3.1-section 4.3.4). Conclusion was made based on the
results obtained from all four screening methods used in this study, to select for the
most potential bacteria capable of producing biosurfactant. Table 4.1 summarized the
results for the screening of biosurfactant-producing bacteria using four different
methods commonly used and described elsewhere [Desai and Banat, 1997].
Table 4.1: Screening of biosurfactant-producing bacteria using four different
methods.
Isolates
Drop
Collapse
ü
x
x
x
x
x
ü
x
x
x
ü
x
x
AB-Cr1
MFTA-W1
RSSF-Cr1
MAB-Cr1
RAS-Cr2
ETL-Cr7
ETL-Cr1
RETL-Cr1
RFTA-Cr3
RETL-Cr3
a
b
c
ü
x
a
b
c
:
:
:
:
:
Blood
Hemolysis
-
Hydrophobicity
Index (%)
5.81
0.20
-0.24
4.46
2.13
3.88
0.89
5.44
8.83
-2.44
100
-
Surface Tension
(mN/m)
35.1
47.1
46.5
54.6
46.7
44.7
35.9
52.9
52.9
43.0
70.1
55.1
Droplet collapse
Droplet remains beaded
Positive control
Negative control
Reference for the calculation of surface tension
4.3.1.1 Bacterial Adherence To Hydrocarbon (BATH) Test
The measurement of bacterial cell hydrophobicity was based on the density of
free bacterial cells remained in aqueous phase (water) of mixture (hexadecanewater), after allowing certain duration of interaction between cells and hexadecane.
49
Following mixing for 10 minutes and allowing to stand, the bacterial cells from the
bulk aqueous phase bound to hydrocarbon droplets and rose with the hydrocarbon,
forming a ‘creamy’ upper layer and a clear aqueous phase.
Rosenberg et al. (1980) found that the upper layer showed an oil-in water
emulsion that consisted of hexadecane droplets covered with the patches of bacteria.
If the cells have no affinity towards the test hydrocarbon, the hydrocarbon droplets
rose and coalesced after mixing and the cells would remained in the bulk aqueous
suspension. In this case, no significant changes would be observed in the turbidity of
the aqueous phase at the bottom layer. The ratios of ODa/ODc measured at the
wavelength of 400nm, was used to quantify (in percentage) the cell surface
hydrophobicity of the bacterial isolates. Therefore, the lower the percentage of
hydrophobicity index indicated to the higher affinity of cells towards hydrocarbon.
Results illustrating the adherence of various bacterial isolates to the
hydrocarbon were presented in Table 4.1. In general, all ten isolates showed a
significantly high ability to adhere on hydrocarbon droplets during 10 minutes of
hydrocarbon-cell interaction. This was based on the value of hydrophobicity index of
less than 9%. However, isolates ETL-Cr1 and MFTA-W1 showed the highest affinity
(0.89 and 0.20%, respectively) towards hexadecane compared to others (2-9%). More
than 99% of cell from cultures of isolates ETL-Cr1 and MFTA-Cr1 were found
removed from the aqueous phase into the hydrocarbon phase based on the value of
optical densities before and after the reaction with 1mL hexadecane. Zhang and
Miller (1994) revealed that high cell hydrophobicities enhanced the contact between
insoluble substrates and cells. This observation suggested that the cell hydrophobicity
could be the characteristic of hydrocarbon-degrading bacteria [Rosenberg et al.,
1980]. It was also reported that most hydrocarbon-degrading bacteria were also
capable of producing biosurfactant [Hommel, 1990; Bodour and Miller-Maier, 2002].
Those with lower cell hydrophobicity (AB-Cr1, MAB-Cr1, RAS-Cr2, ETLCr7, RETL-Cr1 and RFTA-Cr3) might also capable of degrading hydrocarbon and/or
producing biosurfactant though to a lesser extent or subjected to a longer period of
adaptation. Two isolates RETL-Cr3 and RSSF-Cr1 were distinct from other isolates
by their negative values of hydrophobicity index (-2.44% and –0.24%, respectively).
This was due to the higher optical density values (ODa) after the reaction with
hexadecane. This could possibly due to the emulsification of hexadecane into the
50
aqueous phase resulted from the interaction of exopolymeric substances with the
hydrocarbon, rather than the attachment of bacterial cells onto hydrocarbon
molecules [Zajic, 1987].
A simple quantitative method has been described for studying the outer cell
surface of bacteria based on the affinity of these cells to liquid hydrocarbons. Beal
and Betts (2000) showed that the cell surface hydrophobicity increased in the
biosurfactant-producing strains more than those not producing biosurfactant. In
addition, increased in cell surface hydrophobicity favors microbial adhesion and
aggregation [Liu et al., 2004]. Hua et al. (2003) found that the adhesion rate of cell to
hydrocarbon increased with the concentration of biosurfactant produced by Candida
antarctica when grown in n-undecane-containing medium. They concluded that the
biosurfactant produced had improved the hydrophobicity of the microbial cell surface
significantly.
Previous research has suggested that the ability of adhering to bulk
hydrocarbon was a characteristic feature of biosurfactant-producing bacteria [Pruthi
and Cameotra, 1997]. The variation in percentage clearance of aqueous phase in this
study suggested that the affinity for hydrocarbon may vary among biosurfactantproducing bacteria and the hydrocarbon used [Rosenberg et al., 1980]. For some
organisms, the volume of hydrocarbon would also influenced the percentage
clearance of cell from the aqueous [Dillon et al., 1986]. The viscosity of the test
hydrocarbon or size of droplets formed during mixing might affect the degree of
adherence to hydrocarbon. Besides, growth rate, substrate, temperature and pH of the
culture may also influence the hydrophobic properties of cell surface [Liu et al.,
2004]. These showed that the BATH assay was relatively insensitive partly because
of these factors. Thus, a standard set of cultural and preparation conditions could be
adapted (Section 4.2.4) to get a significantly reliable results.
4.3.1.2 Drop-collapsing Test
The drop-collapsing ability of ten bacterial isolates was tested on Penzoil
10W-40 lubricant oil and was found to give the best qualitative indication for the
presence of surfactant. In the presence of surfactant, the liquid droplet spreads over
51
the hydrophobic surface due to the reduction of interfacial tension between the liquid
droplet and the hydrophobic surface. Otherwise, the droplet remains beaded in the
absence of surfactant due to the repellence of water molecules from the hydrophobic
surface. This technique can also be applied as a quantitative method to determine the
surfactant concentrations. The diameter of the sample droplet increased with the
increasing of biosurfactant concentration [Bodour and Miller-Maier, 1998].
Results for the drop-collapsing test of ten bacterial isolates were shown in
Table 4.1. Droplet of all cultures on Penzoil coated surface remained beaded
throughout the experiment, except those of isolates AB-Cr1 and ETL-Cr1. These two
isolates were capable to destabilize the tension between oil-coated surface and
culture droplet (beaded) within an hour, followed by a total collapse of the bead. This
was similar to that observed for SDS used as the positive control in this experiment.
In contrast, drops of cell suspensions from cultures of other isolates were stable and
remained beaded similar to that showed with water as negative control of the
experiment, which indicated to the absence or lack of biosurfactant produced by
these isolates. Therefore, the drop-collapsing test found that both isolates, ETL-Cr1
and AB-Cr1 could be the potential biosurfactant-producing bacteria.
The result obtained in this study was in contrast to those obtained from the
cell hydrophobicity test which designated isolates ETL-Cr1 and MFTA-W1 as the
potential biosurfactant producers instead of ETL-Cr1 and AB-Cr1. Therefore, further
confirmation was required for an accurate selection. Two other methods used were
blood hemolysis test (Section 4.3.3) and surface tension measurement (Section
4.3.4).
The drop-collapse method was based on the ability of surfactants to
destabilize liquid droplets on an oily surface. Youssef et al. (2004) have found that
the oil spreading and drop-collapse method were correlated with the ability of the
cultures to reduce surface tension. There were strong correlation between the dropcollapse method and surface tension reduction, which cultures showing a greater
degree of collapse had low surface tension values. The correlation between the drop
collapsing with the spreading tension between aqueous and hydrocarbon phases also
has been discussed previously by Jain et al., (1991). Drops with higher spreading
tension and lower surface tension will collapse on oily surfaces. In contrast, drops
52
with lower spreading tension or higher surface tension do not have the ability to
spread on an oily surface.
However, the drop-collapse method may not be as sensitive as the oil
spreading technique in detecting low levels of biosurfactant [Youssef et al. 2004].
The amount of surfactant required to cause drop-collapse was found dependent on
the ability of the surfactant to reduce surface and interfacial tension [Bodour and
Miller-Maier, 1998]. The more potent the surfactant, the smaller the quantity and
time required to cause drop collapse.
Both isolates (AB-Cr1 and ETL-Cr1) that showed positive results in this test
tended to break the cohesive forces of the oil and increase the adhesive forces
resulting in the collapse of liquid droplets on oily surfaces. This method was
therefore, very specific since only organisms which produced significant surfaceactive compound, will cause collapse of aqueous drops on oily surfaces [Bodour and
Miller-Maier, 1998]. The test volume required in this technique is much smaller
(5µL) than the volume required for the surface tension measurement (20mL) and the
results were easy to determine visually. Furthermore, this technique was easy to
perform, more reproducible and can be used to screen large number of isolates
[Bodour et al., 2003] though it is not suitable for screening of isolates that have high
emulsifying activity which did not lower surface tension significantly [Jain et al.,
1991].
4.3.1.3 Blood Hemolysis Test
A qualitative assay to determine biosurfactant producer was also developed
based on their ability to cause hemolysis of red blood cells. Screening of
biosurfactant producers via this method was previously outlined that only those
isolates which showed -hemolysis were considered to be the potential biosurfactantproducing microbes [Bernheimer and Avigad, 1970; Carrillo et al., 1996]. The
estimation of this test was based on the fact that surfactants interact strongly with
cellular membranes and proteins [Pape and Hoppe, 1988]. Exotoxins called
hemolysins cause lysis of the red blood cells.
53
Blood agar lysis was used in this study since it is widely used to screen for
biosurfactant production and in some cases, it was used as primary method for
screening purpose [Yonebayashi et al., 2000; Youssef et al., 2004]. Mulligan et al.
(1984) had recommended this method as a preliminary screening method. In
addition, the hemolytic assay was a simple, fast and low-cost method for the
screening of biosurfactant producers on solid medium. Results of the blood
hemolysis for the bacterial isolates were presented in Table 4.1.
In this study, only two bacterial isolates tested (AB-Cr1 and ETL-Cr1)
showed -hemolysis on blood agar plates (Figure 4.1). The -hemolysis pattern was
indicated by the formation of white or clear zone around the bacterial colonies grown
on the blood agar. When all ten isolates tested were grown at 37ºC on blood agar
plates, no clearing zones was observed around colonies up to 18 hours of growth.
However, clear zones were observed after 48 hours of growth on blood agar plates
inoculated with isolates AB-Cr1 and ETL-Cr1, respectively. Further incubation for
up to 72 hours of growth, resulted to the formation of greenish zone around the
colonies of isolates MFTA-W1 and MAB-Cr1, respectively. This type of hemolysis
was known as -hemolysis. The other six bacterial isolates showed neither a greenish
nor a clear zone and were considered as -hemolysis or no hemolysis occurred.
(A)
(B)
Fig. 4.1: -hemolysis on blood agar indicated to the presence of biosurfactant in the
culture of AB-Cr1 (A) and ETL-Cr1 (B).
Carrillo et al. (1996) had proved the efficiency of this method in screening of
biosurfactant-producing bacteria. They found an association between hemolytic
activity and surfactant production. However, there were limitations of using this
method to screen the biosurfactant producer. Not all biosurfactants have a hemolytic
activity and compounds other than biosurfactant might cause hemolysis. Hemolytic
54
activity may also be associated with the presence of lytic enzymes [Jain et al., 1991]
and other microbial products such as virulence [Carrillo et al., 1996] that could
influence the results. Furthermore, this method could not be applied to screen for
microorganisms that required hydrocarbons for biosurfactant production due to the
reaction of hydrocarbons with the red blood cells [Jain et al., 1991]. In addition,
biosurfactant that was poorly diffusible may not lyse the red blood cells [Youssef et
al., 2004]. The absence of hemolytic activity could be due to diffusion restriction of
the surfactant through the blood agar [Jain et al., 1991].
Therefore, this result was further confirmed by the measurement of surface
tension (Section 4.3.4) of cell-free cultures of all isolates, respectively. The surface
tension method was not influenced by the presence of lytic enzymes.
4.3.1.4 Surface Tension Measurement
The measurement of surface tension has been used by many researchers to
measure the surface properties of the biosurfactant [Youssef et al., 2004; Neu and
Poralla, 1990]. However, the measurement of surface tension was inconvenient to
use for screening of a large number of isolates as it was time consuming and required
large volume of sample for analysis. Furthermore, parameters such as pH,
temperature, ionic strength and the composition of the medium might also affected
the surface tension measurement [Bodour and Miller-Maier, 1998].
Table 4.1 summarized the value of surface tension measured for all ten
bacterial isolates after 48 hours incubation at 37ºC in Ramsay medium supplemented
with 5mM glucose. The surface tension values showed were the average of 3
readings from the same culture. The reduction of surface tension was also calculated
with reference to the value of surface tension of sterile medium. The values of
surface tension recorded were also indirectly proportional to the amount of
biosurfactant presence in the growth medium [Robert et al., 1989; Desai and Banat,
1997]. The lowest values of surface tension recorded were 35.1 mN/m and
35.9mN/m, obtained from cell-free cultures of isolates AB-Cr1 and ETL-Cr1,
respectively. Other isolates did not showed a significant decrease of the surface
tension of the cell-free culture and thus, absence of surface-reducing ability as the
55
main characteristic of non biosurfactant-producing bacteria. The maximum reduction
of surface tension obtained with the culture of AB-Cr1 and ETL-Cr1 was
approximately 20-21.2 mN/m which was 2-10 times higher than those obtained with
the culture of other isolates (2.2-12.1 mN/m).
This result was in good agreement with those obtained from the dropcollapsing (Section 4.3.2) and blood hemolysis (Section 4.3.3) tests, which gave a
significant indication to suggest that isolates AB-Cr1 and ETL-Cr1 were the most
potential biosurfactant producer among the ten bacterial isolates used in this study.
4.3.2
Characterization of the Selected Biosurfactant-producing Isolates
4.3.2.1 Colony and Cellular Morphological Characterizations
Colony and cellular characterizations were carried out using fresh cultures of
AB-Cr1 and ETL-Cr1, grown on nutrient agar and Ramsay liquid medium,
respectively. Colonies of AB-Cr1 isolate was appeared as circular with internal
diameter ranging from 5-6mm when grown on nutrient agar. The edge of the colony
was smooth with drop-like elevation (Figure 4.2). Cultures were very sticky on
primary isolation and colonies may be difficult to remove completely from the agar
surface. Whereas, colony of ETL-Cr1 was concentric with internal diameter of 3mm,
irregular margin with flat elevation (Figure 4.3). Both of the isolates were observed
as creamy colonies with slightly slimy for AB-Cr1 isolates. They also grow well on
non-selective media such as nutrient agar with optimum temperature of 37ºC.
Phase-contrast observations revealed that cells of both isolates occurred as
non-motile rods, commonly appeared as pair or single rods. The rods were also
observed as straight with rounded ends (Figure 4.4 (A) and (B)).
56
Figure 4.2: Colony of AB-Cr1 observed under stereo scan microscope using
magnification 50x
Figure 4.3: Colony of ETL-Cr1 observed under stereo scan microscope using
magnification 50x
Figure 4.4: Digital photos of bacterial isolates AB-Cr1 (A) and ETL-Cr1 (B) under
phase-contrast microscope using magnification 100x with oil
immersion.
57
4.3.2.2 Biochemical Characterization
Table 4.2 summarized the results obtained from biochemical tests performed
on the bacterial isolates AB-Cr1 and ETL-Cr1. Both of the isolates tested were
identified as gram negative and non-motile, rod-shaped bacteria formed in singly,
pairs or more than three cells of long chain. The rods were straight with rounded
ends.
Table 4.2: Results for biochemical tests of the selected isolates.
Tests
AB-Cr1
ETL-Cr1
Gram stain
Oxidase
Catalase
Motility
O/F Glucose & Sucrose
Triple Sugar Ion
Nitrate Reduction
Nitrite Reduction
Indole
Gelatin Liquefaction
Urease
Citrate
Negative, long rod
Positive
Positive
Non-motile
Positive
Alkaline/acid
Positive
Negative
Positive
Positive
Positive
Negative, long rod
Positive
Positive
Non-motile
Positive
Alkaline/acid
Positive
Positive
Negative
Positive
Negative
Positive
From the results of biochemical tests and their morphological characteristics,
it was possible to suggest that the isolates AB-Cr1 and ETL-Cr1 might belong to the
genus of Actinobacillus and Aeromonas, respectively. However, the species of these
isolates were difficult to identify by only referring to the biochemical tests.
Even though the genus of the bacteria were successfully identified by the
biochemical test method, the bacteria name were retained as AB-Cr1 and ETL-Cr1,
respectively throughout the study due to the limitation of biochemical test alone in
order to confirmed that these cultures were belong to the genus of Actinobacillus and
Aeromonas, respectively. Further characterization has to be carried out based on the
analysis of the 16S rRNA gene sequence method.
CHAPTER 5
PRODUCTION OF BIOSURFACTANT BY PURE AND MIX BACTERIAL
CULTURES IN SHAKE FLASKS
5.1
Introduction
Majority of the biosurfactant-producing organisms required water-insoluble
substrate as carbon source during biosynthesis [Mulligan, 2004]. There were also
some biosurfactants that have been reported to be produced using water-soluble
compounds such as glucose, sucrose or ethanol as substrate [Desai and Banat, 1997].
However, the biosurfactant in the glucose media was cell-bound and could be
subsequently extracted with organic solvents. This opens some new possibilities in
utilizing waste and cheap carbohydrate media for biosurfactant production [Kosaric
et al., 1987]. Cooper (1986) concluded that the available carbon source, particularly
the carbohydrate used, has influenced the structure and yield of biosurfactant
produced, which in turn alters its surfactant properties. Changing the substrate and
growth conditions also resulted in modification of the polar group in a biosurfactant
[Cooper, 1986].
The best biosurfactant-producing bacteria have been screened (Chapter 4) and
two bacterial strains coded AB-Cr1 and ETL-Cr1 were selected to be studied in
further. In this chapter, growth of both selected strains was optimized and
biosurfactant production was observed under optimum growth conditions. The
effects of glucose and/or crude oil addition on biosurfactant production were also
observed with both isolates. The ability of these isolates to produce biosurfactant in
pure and mix cultures was studied, respectively. All experiments were carried out in
shake flasks.
59
5.2
Methodology
5.2.1
Optimization of Growth
Growth of two potential biosurfactant-producing bacteria, strain AB-Cr1 and
ETL-Cr1 selected previously (Section 4.3.1) was optimized. Optimization was
carried out as a factor of initial glucose concentration (Section 5.2.1.1), pH (Section
5.2.1.2) and temperature (Section 5.2.1.3). All experiments were set up in triplicate
and analysis at variance (ANOVA) was carried out to determine significancy of
differences in the specific growth rates obtained.
5.2.1.1 Effect of Initial Glucose Concentration on Growth
The experiment was set up using 250mL Erlenmeyer flasks containing
100mL of Ramsay medium (Section 3.2.1.1). The medium was inoculated (10% v/v)
with inoculum of strains AB-Cr1 and ETL-Cr1 (Section 3.3.1), respectively. The pH
of growth medium was adjusted to 6.5-6.8 [Ramsay et al., 1983] and the cultures
were incubated with shaking (200rpm) at 37ºC. The medium was supplemented with
0, 1, 3, 5, 8 and 10mM glucose (each was in triplicate). Samples were taken out at
regular intervals to analyze for growth by measuring the optical density at 600nm
(Section 3.4.1.1). The bacterial cell dry weight was determined as mentioned in
Section 3.4.1.2. The specific growth rates (µ) of cultures were then calculated
respectively, based on the plot of ln [cell dry weight at log phase] versus time (hour).
Analyses of variance (ANOVAs) were performed to determine significancy of the
difference between calculated µ obtained from experiments.
5.2.1.2 Effect of Initial pH on Growth
Ramsay medium (Section 3.2.1.1), supplemented with the optimum initial
glucose concentration obtained previously (Section 5.2.1.1) was used in this
experiment. The medium was inoculated (10% v/v) with inoculum of strains AB-Cr1
and ETL-Cr1 (Section 3.3.1), respectively. Cultures were incubated with shaking
60
(200rpm) at 37ºC. Samples were taken out at regular intervals to analyze for growth.
The initial pH of growth medium was set at 5.0, 6.0, 6.5, 7.0, 7.5 and 8.0 (each was
in triplicate). The specific growth rates of cultures were calculated respectively, as
mentioned in Section 5.2.1.1.
5.2.1.3 Effect of Temperature on Growth
Pure culture of isolates AB-Cr1 and ETL-Cr1 were inoculated (10% v/v) into
Ramsay medium (Section 3.2.1.1), adjusted to the optimum pH (Section 5.2.1.2) and
supplemented with the optimum initial glucose concentration (Section 5.2.1.1). The
same experimental set up (Section 5.2.1.2) was used in this study except that the
incubation temperature was varied. The temperature was set at 30º, 37º, 45º and
55ºC. All experiments were carried out in triplicate. Samples were taken out at
regular intervals to analyze for growth. The specific growth rates of cultures were
calculated respectively, as mentioned in Section 5.2.1.1.
5.2.2 Biosurfactant Production under the Optimized Growth Condition
Production of biosurfactant by both bacterial isolates, AB-Cr1 and ETL-Cr1
were monitored under the optimized growth conditions (Section 5.2.1). The
experiment was set up using 250mL Erlenmeyer flasks containing 100mL of Ramsay
medium (Section 3.2.1.1), adjusted to optimum pH (Section 5.2.1.2) and
supplemented with the optimum initial glucose concentration (Section 5.2.1.1),
respectively. The medium was then inoculated (10% v/v) with inoculum of strains
AB-Cr1 and ETL-Cr1 (Section 3.3.1), respectively. Cultures were incubated with
shaking (200 rpm) at their optimum temperature (Section 5.2.1.3). All experiments
were carried out in triplicate. Samples were taken out at regular intervals to analyze
for growth (Section 3.4.1.2), glucose consumption (Section 3.4.2), surface and
interfacial tension (Section 3.4.3.1 and 3.4.3.2) and the amount of biosurfactant
produced (Section 3.5.1 and 3.5.2). The specific growth rates of cultures were
calculated as mentioned in Section 5.2.1.1, respectively.
61
5.2.3 Effect of Glucose and Crude Oil on Biosurfactant Production
Production of biosurfactant and the growth of AB-Cr1 and ETL-Cr1 isolates
were studied in Ramsay medium (Section 3.2.1.1) in the presence of both glucose
and crude oil and either in the presence of glucose or crude oil in the medium. The
same experimental set up (Section 5.2.2) was used in this study with the addition of
crude oil (5% v/v). The crude oil was autoclaved (121ºC, 101.3kPa for 15 minutes)
separately prior to add into the growth medium respectively. All experiments were
carried out in triplicate. Samples were taken out at regular intervals to analyze for
growth (Section 3.4.1.2), glucose consumption (Section 3.4.2), surface tension
(Section 3.4.3.1), and the amount of biosurfactant produced (Section 3.5.1 and 3.5.2).
5.2.4
Production of Biosurfactant by Bacterial Mix Cultures
The study of biosurfactant production by bacterial mix cultures was carried
out as a comparative estimation of the bacterial potential in producing biosurfactant
in relation with those in pure cultures. The growth of bacterial mix cultures (AB-Cr1:
ETL-CR1) and its production of biosurfactant were examined in shake flasks using
Ramsay medium (Section 3.2.1.1) added with glucose and glucose plus crude oil as
carbon sources, respectively. Both bacterial strains were mixed at the ratio of 1:1
(AB-Cr1: ETL-Cr1) under the above mentioned conditions.
Inoculum was first prepared for each strain as mentioned in Section 3.3.1.
Equal volumes of culture broth from the isolates were served as inoculum to prepare
the bacterial mix culture of 1:1 system. The bacterial mix cultures were grown in
Ramsay medium, containing 3mM glucose as the sole source of carbon and in
medium containing both 3mM glucose and crude oil (5% v/v). The same
experimental set up (Section 5.2.2) was used in this study (each was in triplicate).
Samples were taken out at regular intervals to analyze for growth as mix culture
(Section 3.4.1.2), glucose concentration (Section 3.4.2), surface and interfacial
tension (Section 3.4.3.1 and 3.4.3.2) and the amount of biosurfactant produced
(Section 3.5.1 and 3.5.2).
62
5.3
Results and Discussion
5.3.1 Optimization of Growth
Growth of bacterial isolates AB-Cr1 and ETL-Cr1 was optimized as a factor
of initial glucose, pH and temperature, respectively. Results obtained from this study
were recorded and discussed in the following sections (Section 5.3.1.1-5.3.1.3).
5.3.1.1 Effect of Initial Glucose Concentrations on Growth
In this study, growth of both bacterial isolates coded AB-Cr1 and ETL-Cr1
was carried out in batch culture system. Bacteria were grown in Ramsay medium,
supplemented with various initial concentration of glucose (Section 5.2.1.1). Figure
5.1 and 5.2 showed the effect of various initial glucose concentrations on growth of
bacterial isolates AB-Cr1 and ETL-Cr1, respectively. Results indicated that both
isolates shared similar pattern of growth, though the lag growth phase of isolate ETLCr1 (1 hour) was found shorter than that of isolate AB-Cr1 (2 hours). Similarly, the
concentration of 3 to 10mM glucose added into the medium resulted in the
production of biomass significantly higher (~5 g/L (AB-Cr1) and ~6 g/L (ETL-Cr1))
than those in 1mM glucose (~3 g/L (AB-Cr1) and ~4 g/L (ETL-Cr1)). However, the
maximum biomass produced in medium supplemented with glucose ranging from 3
to 10mM was found not significantly different for both isolates, respectively (Table
5.1).
63
0.6
OD 600nm
0.5
0.4
0.3
0.2
0.1
0
0
1
2
3
4
Time, h
5
6
7
8
Fig. 5.1: Growth curve of AB-Cr1 grown in Ramsay medium pH 6.5-6.8 at 37ºC as a
factor of initial glucose concentrations. Glucose concentrations are
0mM,
1mM,
3mM,
5mM,
8mM and
10mM.
0.7
OD 600nm
0.6
0.5
0.4
0.3
0.2
0.1
0
0
1
2
3
4
Time, h
5
6
7
8
Fig. 5.2: Growth curve of ETL-Cr1 grown in Ramsay medium pH 6.5-6.8 at 37ºC as
a factor of initial glucose concentrations. Glucose concentrations are
0mM,
1mM,
3mM,
5mM,
8mM and
10mM.
Both isolates also indicated that addition of at least 1mM glucose was able to
accelerate their growth. This was based on the increase in their maximum specific
growth rates (µ) (Figure 5.3) and biomass (X max) (Table 5.1) when 1mM glucose was
initially supplemented into the growth medium. Both isolates showed approximately
1.9 and 2.5 times higher biomass produced and grew at about 2.5 and 2.9 times faster
when the lowest concentration of glucose was initially added in the culture of ABCr1 and ETL-Cr1, respectively. However, both isolates did not show an obvious
effect upon the addition of 3, 5, 8 and 10mM of glucose (Figure 5.3).
Specific Growth Rates, h -1
64
0.4
0.3
0.2
0.1
0
0
2
4
6
[Initial glucose], mM
8
10
Fig. 5.3: The specific growth rates of AB-Cr1
and ETL-Cr1
grown in
Ramsay medium pH 6.5-6.8 at 37ºC, as a factor of initial glucose
concentrations.
Table 5.1 showed the mean values of the specific growth rates for isolates
AB-Cr1 and ETL-Cr1, obtained from a triplicated experiment. The analysis of
variance (ANOVA) indicated that the values of specific growth rates between isolate
AB-Cr1 and ETL-Cr1 were significantly different (p<0.05) under all conditions
tested. Both isolates exhibited a relatively good growth in the presence of between
1mM to 10mM glucose as carbon source. However, both isolates were similarly
showed the maximum growth in the presence of 3mM glucose, with the specific
growth rates of 0.319h-1 and 0.306h-1 for isolates AB-Cr1 and ETL-Cr1, respectively.
At the optimum concentration of glucose, these isolates grew approximately 1.3 (ABCr1) and 1.2 (ETL-Cr1) times faster compared to those grown on the same medium
supplemented with 1mM glucose.
Growth of both isolates was also found dependent on the glucose supplement,
or else halted (by approximately 75%) in its absence. The dependency of both
isolates on glucose for growth was further confirmed by the very limited growth (µ =
0.099h-1 and 0.089h-1 for isolates AB-Cr1 and ETL-Cr1, respectively) observed in
the glucose-free medium (Figure 5.3).
65
Table 5.1: Specific growth rates and maximum biomass of AB-Cr1 and ETL-Cr1
grown at 37ºC, pH 6.5-6.8 in medium supplemented with various initial
glucose concentrations.
AB-Cr1
ETL-Cr1
Glucose
concentration (mM)
µ (h-1)
Xmax
µ (h-1)
Xmax
0
0.099
1.59
0.089
1.62
1
0.248
2.97
0.259
4.07
3
0.319
4.87
0.306
5.82
5
0.312
4.77
0.298
5.81
8
0.298
4.93
0.292
5.91
10
0.286
4.56
0.306
5.91
5.3.2
Effect of Initial pH on Growth
The maximum growth rates previously obtained with both isolates grown in
the medium supplemented with 3mM of glucose and was chosen for further
experiments. The effect of pH was studied in the pH ranging from 5 to 8.
Figure 5.4 exhibited the effects of pH on the growth rates of isolates AB-Cr1
and ETL-Cr1. The optimum pH for growth of both isolates was found at pH 7.0,
respectively. At this pH, the maximum specific growth rate (µ) and the maximum
biomass obtained were 0.369h-1 and 5.60 g/L (AB-Cr1) and 0.323h-1 and 6.02 (ETLCr1), respectively (Table 5.2). Both isolates also showed higher biomass (1-1.1 times
higher) and grew significantly faster (1-1.2 times faster) than those observed in
cultures grew in the same medium initially adjusted to pH 6.5-6.8 (Section 5.3.1).
The specific growth rates of both isolates were found affected by the pHs tested
at similar pattern. However, both isolates showed some distinctions when grown at
pH between 7.5 to 8.0. Isolate ETL-Cr1 was found more alkaline-tolerant in
comparison with that of isolate AB-Cr1 (Figure 5.4). This was clearly showed by the
similar specific growth rates of isolate ETL-Cr1 at pH 7.5 and 8.0 (µ 0.256h-1 and
0.250h-1, respectively). This was in contrast to that of isolate AB-Cr1, whose growth
was drastically decelerated by the increase of pH from 7.0 (0.369h-1) to 7.5 (0.285h-1)
and 8.0 (0.190h-1). At pH 8.0, isolate ETL-Cr1 grew at the rate of approximately two
66
times slower than that observed at the optimum pH condition. Both isolates were not
acid-tolerant bacteria as the decreased of pH (from pH 7.0 to pH 5.0) results in
drastic decrease in the specific growth rates. Their growth rates at the lowest pH
tested were approximately 1.8 to 2.3 times slower compared to those in the optimum
Specific Growth Rates, h-1
pH.
0.4
0.3
0.2
0.1
4
5
6
7
8
pH
Fig 5.4: Growth optimization of isolates AB-Cr1
and ETL-Cr1
grown at
0
37 C in medium supplemented with 3mM glucose, as a factor of pH.
The maximum biomass produced in medium initially adjusted to pH 5.0-8.0
was found significantly different for both isolates, respectively (Table 5.2). The
production of biomass of AB-Cr1 isolate was found significantly higher (~4.5-5.6
g/L) in the medium adjusted to initial pH of 6.5 to 7.5, than those in the medium
adjusted to initial pH of 5.0-6.0 (~3.0-3.5 g/L) and 8.0 (~2.8 g/L), respectively. This
result indicated that AB-Cr1 isolate was able to grow well in medium initially
adjusted to nearly the neutral pH.
However, ETL-Cr1 isolate was more pH-tolerant compared to AB-Cr1
isolate. This isolate was able to produce higher maximum biomass (~4.7-6.0 g/L) in
medium initially adjusted to pH ranging from 6.0 to 8.0. At the lowest initial pH
tested, the maximum biomass of ETL-Cr1 isolate was halted by approximately 4557%.
67
Table 5.2: Specific growth rates and maximum biomass of AB-Cr1 and ETL-Cr1
grown in Ramsay medium supplemented with 3mM glucose adjusted to
various initial pH.
AB-Cr1
ETL-Cr1
pH
µ (h-1)
Xmax
µ (h-1)
Xmax
5.0
0.159
3.00
0.176
2.58
6.0
0.283
3.54
0.286
5.56
6.5
0.305
4.49
0.288
5.18
7.0
0.369
5.60
0.323
6.02
7.5
0.285
4.66
0.256
4.69
8.0
0.190
2.83
0.250
4.77
5.3.3 Effect of Temperature on Growth
Batch culture studies were performed by varying the temperature ranging from
30ºC to 55ºC. The effect of temperature on the growth rates of both isolates AB-Cr1
Specific Growth Rates, h-1
and ETL-Cr1 were shown in Figure 5.5.
0.4
0.3
0.2
0.1
0
20
30
40 o
Temperature, C
50
60
Fig. 5.5: Growth optimization of isolates AB-Cr1
and ETL-Cr1
grown
in medium supplemented with 3mM glucose at pH 7.0, as a factor of
temperature.
The optimum temperature for growth of both isolates was observed to be at
37ºC, with the maximum specific growth rates (µ) and biomass (X max) (Table 5.3). At
68
the optimum temperature, the maximum specific growth rate and the maximum
biomass obtained were 0.369h-1 and 5.60 g/L (AB-Cr1) and 0.323h-1 and 6.02 (ETLCr1), respectively (Table 5.3).
Table 5.3: Specific growth rates and maximum biomass of isolates AB-Cr1 and
ETL-Cr1 grown in medium supplemented with 3mM glucose at pH 7.0,
incubated at various temperatures.
AB-Cr1
ETL-Cr1
Temperature (ºC)
µ (h-1)
Xmax
µ (h-1)
Xmax
30
0.236
4.25
0.258
4.05
37
0.369
5.60
0.323
6.02
45
0.283
4.43
0.212
4.59
55
0.101
1.63
0.120
2.24
A decrease or increase in the incubation temperature may lead to the changes
of microbial metabolism, thus alter the growth rates of the microbes [Guerra-Santos
et al, 1986]. In this study, both isolates showed similar rates of growth at 30ºC and
45ºC. Both isolates showed higher biomass (1.3-1.5 times higher) and grew
significantly faster (1.3-1.6 times faster) at the optimum temperature than those
observed in cultures grown in the same medium incubated at 45º and 30ºC,
respectively (Table 5.3).
Both isolates also showed similar pattern of growth with the increase of
incubation temperature from 45ºC to 55ºC. Growth was observed to be drastically
decelerated (1.8-2.8 times slower) with lower biomass produced (2.1-2.7 times
lower) when grows from 45ºC to 55ºC by ETL-Cr1 and AB-Cr1 isolates,
respectively.
At the lower temperature (30ºC), slower rate of growth (compared to those at
37ºC) could be possibly due to low metabolism rates of these isolates. However, at
the temperature higher than the optimum, slower rates of growth observed was
majorly due to the incapability of isolates to tolerate with excess heat supplied for
growth and lead to disruption of bacterial cellular.
69
5.3.2
Biosurfactant Production under the Optimized Growth Conditions
In this study, biosurfactant production by both isolates AB-Cr1 and ETL-Cr1
under the optimized growth conditions was monitored. Both isolates were grown in
Ramsay medium supplemented with 3mM glucose, adjusted to an initial pH 7.0 and
incubated at 370C. The relationship between cell growth, glucose consumption and
biosurfactant production of AB-Cr1 and ETL-Cr1 isolates were as showed in Figures
5.6 and 5.7, respectively.
Biomass &
Biosurfactant, g/L
0.3
0.2
0.1
[Glucose], g/L
0.4
10
9
8
7
6
5
4
3
2
1
0
0
0
1
2
3
4
5
Time, h
6
7
8
9
Fig. 5.6: Relationship of growth
, glucose consumption
and biosurfactant
production
by AB-Cr1 isolate grown in Ramsay medium
supplemented with 3mM glucose, adjusted to initial pH 7.0 and incubated
at 37ºC.
The production of biosurfactant by AB-Cr1 was found associated to cell
growth. At the exponential growth phase (between 2-5 h), this bacteria showed a
paralled relationship to the production of biosurfactant and correlated to the glucose
utilization. Glucose was drastically consumed during the exponential growth phase at
the rate of 0.093 gh-1 and followed by slower rate of 0.043 gh-1 during the stationary
growth phase. Biosurfactant production was also found gradually increased during
the stationary growth phase where culture was in their limiting condition. Growthassociated production of biosurfactant has been reported for Bacillus licheniformis
JF-2 [Lin et al. 1993] and Bacillus subtilis [Cooper et al. 1981].
The maximum biosurfactant produced was 8.76 g/L during the stationary
growth phase (at 8 hours incubation). Maximum production of biosurfactant was
70
found paralleled to the maximum reduction of surface tension (Appendix C). Surface
tension of culture medium drastically reduced (from 58mN/m to 37mN/m) during the
maximum cell growth (maximum growth of biosurfactant) followed by gradual
decrease during prolonged incubation prior to reach the maximum reduction of
32mN/m at 8 hours of incubation.
0.4
6
0.3
5
4
0.2
3
2
0.1
[Glucose], g/L
Biomass &
Biosurfactant, g/L
7
1
0
0
0
1
2
3
4
5
Time, h
6
7
8
9
Fig. 5.7: Relationship of growth
, glucose consumption
and biosurfactant
production
by ETL-Cr1 isolate grown in Ramsay medium
supplemented with 3mM glucose, adjusted to initial pH 7.0 and incubated
at 37ºC.
Figure 5.7 showed the trend of biosurfactant production by isolate ETL-Cr1.
Similar to that of isolate AB-Cr1, this isolate also showed a growth-linked
biosurfactant production. This was indicated by the increase of biosurfactant
concentration during the exponential growth phase of this isolate. Although higher
amount of biomass (Xmax = 5.97 g/L) was produced by this isolate, the maximum
biosurfactant formation was relatively lower (Pmax = 6.33 g/L @ 8 hours), compared
to the maximum biosurfactant formation by AB-Cr1 (Pmax = 8.76 g/L @ 8 hours).
The specific rate of biosurfactant formation (qp) for isolate ETL-Cr1 was also found
lower (qp = 0.31 g/g/h) than that of isolate AB-Cr1 (q p = 0.53 g/g/h). The highest
reduction of surface tension occurred during the maximum production of
biosurfactant (8 hours) from 62mN/m to 48mN/m.
A drastic utilization of glucose supplied was observed after 2 hours of
incubation for both isolates (Figures 5.6 and 5.7). During the maximum production of
biomass (t = 5 hours), approximately 60% of glucose was consumed giving the yield
71
of biomass on glucose (Yx/s) of 24.91 and 26.14 for AB-Cr1 and ETL-Cr1 isolates,
respectively. The glucose was almost depleted during the maximum production of
biosurfactant (t = 8 hours), giving the Yp/s of 22.53 and 16.16 g biosurfactant/ g
glucose for isolates AB-Cr1 and ETL-Cr1, respectively. During this time, the pH of
the culture medium fell from 7.0 to approximately 6.0 and 5.8 for AB-Cr1 and ETLCr1, respectively. This was possibly due to the production of weak organic acid such
as acetic acid following the catabolism of glucose [Hommel and Ratledge, 1993].
5.3.3
Production of Biosurfactant in Crude Oil-containing Medium
The ability of both isolates AB-Cr1 and ETL-Cr1 to grow in Ramsay medium
(Section 3.2.1.1) supplemented with crude oil has been observed though quantitative
analysis was not carried out. In this study, the effect of crude oil addition on
biosurfactant production by both isolates was monitored in the medium
supplemented with either (i) glucose or (ii) crude oil or (iii) glucose + crude oil,
respectively. The bacteria were incubated at their optimum conditions i.e: at 37ºC in
the medium initially adjusted to pH 7.0.
This study was also carried out to determine whether microbes isolated from
the oil processing wastes were capable of producing a good surface-active compound
when grown in the medium containing crude oil. A neutral pH of 7.0 used in this
study has also been reported to be optimal for biodegradation of crude oil [Rahman et
al., 1999]. A temperature of 37ºC was previously reported suitable for the growth of
bacterial isolate in the presence of crude oil as the rate of hydrocarbon metabolism
was maximum for most mesophilic bacteria at this temperature [Atlas, 1981].
The correlations between biosurfactant production and growth, changes of pH
and substrate consumption in glucose + crude oil medium were monitored, as
illustrated in Figure 5.8 (A) and (B), for both isolates AB-Cr1 and ETL-Cr1,
respectively. Results also showed that both isolates were able to grow and produce
biosurfactant with crude oil as their sole source of carbon (Figure 5.9). However,
growth and biosurfactant production were found less than those observed in the
medium added with either glucose or glucose and crude oil (Table 5.4). This could be
explained by the fact that glucose was the most preferable carbon source for growth
72
of bacteria in comparison with crude oil due to the complex structure of the crude oil
itself and often more difficult to be degraded by microbes [Hommel, 1990].
(A)
(B)
Fig. 5.8: Relationship of growth
, glucose consumption
, pH
, surface
tension
and biosurfactant production
for isolates AB-Cr1 (A)
and ETL-Cr1 (B), grown in Ramsay medium supplemented with glucose
and crude oil, respectively.
The potential ability of both isolates to grow on hydrocarbon has been
initially observed through the BATH test (Section 4.3.1.1). However, their
73
dependency to the addition of prefixed organic carbon for crude oil consumption was
yet to be observed. This particular study has showed that growth of these isolates on
crude oil was not dependent on the addition of prefixed organic carbon. This was
proved by a significant growth observed in the medium added with crude oil as the
sole source of carbon (Figure 5.9) though at the slower specific growth rates (0.010
and 0.004 h-1 for isolate AB-Cr1 and ETL-Cr1, respectively) compared to those
grown in the medium with glucose as the sole source of carbon (0.366 and 0.319 h-1
for isolate AB-Cr1 and ETL-Cr1, respectively). In medium supplemented with both
glucose and crude oil, growth of both isolates was found faster (0.086 and 0.062 h-1
for isolate AB-Cr1 and ETL-Cr1, respectively) compared to those grown in the
medium with crude oil as sole source of carbon.
Biomass & Biosurfactant, g/L
5
4
3
2
1
0
0
20
40
Time, h
60
80
100
Fig. 5.9: Relationship of growth (opened symbol) and biosurfactant production
(closed symbol) by isolates AB-Cr1
,
and ETL-Cr1
,
grown in Ramsay medium supplemented with 5% (v/v) crude oil.
However, in contrast to those observed in glucose (Figure 5.6 and 5.7) and
crude oil (Figure 5.9) medium, an extended period of stationary growth phase
occurred during prolonged incubation of cultures in medium containing glucose and
crude oil (Figure 5.8). A slight increased in biomass was observed (at 72 h) in culture
of AB-Cr1 with the highest biomass of 4.74 g/L at 96 h of incubation (Figure
5.8(A)). An extension of the stationary growth phase could be due to the utilization
of crude oil as their energy source after glucose was came to depletion.
74
The phenomena of delayed in crude oil utilization by several microbes has
been reported due to a number of different biochemical reaction involved in alkane
utilization including their terminal hydroxylation and the -oxidation [Witholt et al.,
1990]. In addition, the presence of glucose in the crude oil medium resulted to
catabolite repression, which also delayed crude oil utilization by the bacterial strains.
The machinery for glucose catabolism was constitutively expressed, therefore
glucose would be instantly consumed by the microbes in comparison to other carbon
sources including crude oil, which catabolism machinery was induced only by its
presence [Hommel and Ratledge, 1993].
The maximum production of biosurfactant by both isolates in glucose + crude
oil medium (P max = 8.26 and 7.18 g/L for isolates AB-Cr1 and ETL-Cr1,
respectively) was found similar to those obtained in glucose medium (Pmax = 8.76
and 6.33 g/L for isolates AB-Cr1 and ETL-Cr1, respectively). However,
biosurfactant production by both isolates in glucose + crude oil medium occurred in
two phases (Figure 5.8). The first phase of biosurfactant production was observed
during their exponential growth phase where glucose was drastically utilized as both
carbon and energy sources.
Depletion of glucose in the growth medium was particularly the limiting
factor for growth of both isolates and resulted to the stationary growth phase. Drastic
increased in biosurfactant production was obtained within 24 hours of incubation
with the maximum production of 5.54 and 4.42 g/L at the maximum velocity (Vmax)
of 0.123 and 0.069 h-1 for both isolates AB-Cr1 and ETL-Cr1, respectively. This was
followed by a gradual increased of biosurfactant concentration within 36 hours
period prior to the second phase of biosurfactant production during their stationary
phase of growth. This resulted to an increased of biosurfactant concentration up to
8.26 and 7.18 g/L for isolates AB-Cr1 and ETL-Cr1, respectively. It was possible to
assume that the production of biosurfactant during this phase occurred as a result of
crude oil utilization due to the depletion of glucose in the growth medium.
The presence of biosurfactant produced during the first phase might
potentially involved in assisting the adhesion of microbial cells to the crude oil by
lowering the interfacial tension between the two phases (aqueous-oil), thus making
the non-soluble substrate (crude oil) more readily available to these bacteria
[Fiechter, 1992]. This was further supported by an observation to the ability of the
75
biosurfactant produced by both isolates to cause emulsification of hydrocarbon. It has
been reported that the utilization of hydrocarbons by microbe was made easier in the
form of their emulsions [Hommel, 1990].
It could be concluded that the main physiological role of biosurfactant
produced was to permit microbes to grow on water-immiscible substrates, thus
increasing the bioavailability of the substrate for uptake and metabolism.
Both isolates showed a similar ability to produce biosurfactant in the crude oil
medium. The maximum biosurfactant produced were 4.51 and 4.20 g/L for isolates
AB-Cr1 and ETL-Cr1, respectively. These amounts were approximately two times
lower than those produced in glucose + crude oil medium. Therefore, it was possible
to suggest that the glucose added into the medium has a significant role in inducing
biosurfactant production by both isolates. Enhanced biosurfactant production has also
been studied by the addition of several organic acids including lactic, acetic and
pyruvic acids [Desai et al., 1994]. Utilization of crude oil was highly energy
consuming process, which involved several different complex biochemical reactions.
Therefore, less energy could be obtained for growth and biosurfactant production by
these isolates grown in crude oil medium not supplemented with glucose.
The production of biosurfactant was also qualitatively analyzed by measuring
the reduction of surface tension. Drastic reduction of surface tension was observed
during the exponential growth phase of bacteria at all conditions tested. However,
this method was limited by the formation of micelle at a certain concentration of
biosurfactant produced [Lin et al., 1993]. Therefore, reduction of surface tension
could no longer be observed although further increased in biosurfactant concentration
was obtained during prolonged incubation of cultures, such as those observed in
glucose + crude oil medium (Figure 5.8).
A slight decreased in pH was also observed during growth and biosurfactant
production by isolates AB-Cr1 and ETL-Cr1, respectively in glucose + crude oil
medium. However, the pH was found increasing up to ~ 6.6-6.8 during the stationary
growth phase of both isolates. Catabolism of glucose was commonly resulted to the
production of acids such as pyruvic and lactic acids as by product though its
accumulation was not observed as the bacteria was able to consume it as an
alternative source of energy for survival [Hommel and Ratledge, 1993].
76
In this experiment, the production of weak organic acid and its consumption
might have some relation to the mechanism of biosurfactant synthesis. It has been
previously reported that the biosurfactant was consisted of at least one portion of
lipid which formation was initially involved consumption of acetate in the form of
acetic acid and transfer of Co-enzyme A (CoA) to the lipid backbone chain for
further elongation [Hommel and Ratledge, 1993]. However, the relationship between
pH changes and biosurfactant production could not yet be concluded as more detail
analysis on the role of organic acid in biosurfactant production was required.
Table 5.4 summarized the kinetic analysis of growth and biosurfactant
production for both isolates AB-Cr1 and ETL-Cr1 grown in the medium
supplemented with either glucose or crude oil or both glucose and crude oil.
77
Table 5.4: Kinetic analysis of growth and biosurfactant production for isolates AB-Cr1 and ETL-Cr1 grown at 370C, in medium
supplemented with either glucose or crude oil or both glucose and crude oil.
ETL-Cr1
(3mM glucose
+ crude oil 5%)
0.062
3.5 @ 48h
7.18 @ 108h
0.07
AB-Cr1
(Crude oil 5%)
ETL-Cr1
(Crude oil 5%)
0.319
5.97 @ 5h
6.33 @ 8h
0.79
AB-Cr1
(3mM glucose
+ crude oil 5%)
0.086
4.74 @ 96h
8.26 @ 108h
0.08
0.010
2.23 @ 72h
4.51 @ 96h
0.05
0.004
1.99 @ 72h
4.20 @ 96h
0.04
16.16
26.14
0.97
0.31
18.5
21.44
12.51
1.71
0.15
22.5
20.14
7.88
2.56
0.16
15.5
n.d
n.d
2.38
0.02
15.5
n.d
n.d
2.46
0.01
14.5
Isolates
AB-Cr1
(3mM glucose)
ETL-Cr1
(3mM glucose)
µ (h-1)
Xmax (g/L)
Pmax (g/L)
Productivity
(g/L/h)
Yp/s (g/g)
Yx/s (g/g)
Yp/x (g/g)
q p (g/g/h)
STmax (mN/m)
0.366
5.62 @ 5h
8.76 @ 8h
1.10
22.53
24.91
1.45
0.53
32
n.d : not determined
78
In general, both isolates required to the addition of glucose for growth and
biosurfactant production. The presence of crude oil in the medium resulted to less
biomass (up to 4.74 g/L) produced by both isolates in comparison to those grown in
the medium supplemented with glucose only (between 5.0-6.0 g/L). The inhibition of
biomass production might be due to a requirement for direct physical interaction of
the bacterial cells with the hydrophobic substrate in order to initiate the crude oil
degradation [Rosenberg and Rosenberg, 1981]. Furthermore, the physiology of crude
oil itself that contains mutagenic, carcinogenic and growth inhibiting compounds
might contributed to the low value of biomass and biosurfactant obtained [Van Dyke
et al., 1991].
The yield of biomass based on substrate consumption (Yx/s) was found higher
in glucose medium (~24-26 g/g) than those in glucose + crude oil medium. However,
the yield of biosurfactant production based on the substrate consumption (Yp/s) was
not significantly differed in both medium tested. Both isolates showed an efficient
production of biosurfactant based on the maximum biomass (Yp/x) in the medium
supplemented with crude oil either as the sole source of carbon or as an alternative
carbon source.
The relationship between Yp/x and µ would yield the specific rates of product
formation (q p). The correlation between q p and µ could be used to conclude the
nature of product formation either growth-associated or non growth-associated.
Figure 5.10 showed that the changes of qp was directly related to µ for both isolates.
Therefore, this indicated to the growth-associated production of biosurfactant by
these isolates.
79
Specific Rates of Product
Formation, g/g/h
0.6
0.5
(iii)
0.4
0.3
(ii)
0.2
0.1
(i)
0
0
0.1
0.2
0.3
Specific Growth Rates, h-1
0.4
Fig. 5.10: Relationship between specific growth rates (µ) of isolates AB-Cr1
and ETL-Cr1
with the specific rates of product formation (q p) in
medium supplemented with either (i) crude oil, or (ii) both glucose and
crude oil, or (iii) glucose.
The productivity of biosurfactant was also observed to be 14-22 and 11-20
times greater in the medium containing glucose as the sole source of carbon for both
isolates AB-Cr1 and ETL-Cr1, respectively. Both strains that isolated from crude oil
were found to produce biosurfactant in such a good yield, thus increasing the
mobility and solubility of hydrophobic contaminants which is essential for effective
microbial degradation. It could be used as for the treatment of hydrocarboncontaminated soils and in aquatic environments. Therefore, the conditions of the
culture medium in the presence of crude oil have to be further optimized in order to
increase the growth and the activity of biosurfactant produced.
5.3.4
Production of Biosurfactant by Bacterial Mix Cultures
In this study, the production of biosurfactant by bacterial mix culture systems
at the ratios of 1:1 (AB-Cr1: ETL-Cr1) grown in Ramsay medium (Section 3.2.1.1)
supplemented either with glucose or both glucose and crude oil were monitored. The
relationship between cell growth and biosurfactant production by the bacterial mix
culture systems grown in medium supplemented with either glucose or glucose and
crude oil were as showed in Figure 5.11. In this study, growth of both isolates was
80
not individually monitored due to the difficulty to differentiate their colonies on solid
medium. Therefore, no data on the survival of individual strain was reported in this
6
7
5
6
5
4
4
3
3
2
2
1
Biosurfactant, g/L
Biomass, g/L
study.
1
0
0
0
50
100
Time, h
150
Fig. 5.11: Relationship of growth (opened symbol) and biosurfactant production
(closed symbol) by bacterial mix culture system 1:1, grown in Ramsay
medium supplemented with glucose
,
and glucose + crude oil
,
.
Results showed that both isolates AB-Cr1 and ETL-Cr1 were able to grow as
a mixture and produced biosurfactant in the medium supplemented with either
glucose or both glucose and crude oil as their carbon source(s). However, growth in
the bacterial mix culture system was found lower than those observed in single
culture of both isolates in the medium supplemented with glucose as the sole carbon
source (Section 5.3.2). This was indicated by the maximum biomass obtained in the
mix culture system which was ~1.5 times lower than those obtained the single culture
of both isolates.
The specific growth rate of the bacterial mix culture system was found 1.7
time faster in the glucose medium (0.243h-1) compared to those grown in the glucose
+ crude oil medium (0.096h-1). This result was in good agreement with the data
previously obtained (Section 5.3.2 and 5.3.3) that both isolates grew faster in the
medium containing glucose as the sole source of carbon. Drastic increased in
biomass (exponential growth phase) was observed within 7-12 hours of incubation
with the maximum biomass of approximately 3.0 g/L and 4.4 g/L for both mix
81
culture systems in glucose and glucose + crude oil medium, respectively (Figure
5.11). The depletion of glucose (Appendix D) in the medium resulted to the early
death phase occurred in both bacterial systems. This could possibly due to the
competition between both isolates in the mix culture systems for their survival in
carbon limited growth condition (depletion of glucose).
However, growth of the mix culture system in medium supplemented with
glucose and crude oil was observed occurred in two phases (Figure 5.11). The first
phase of growth was observed within the 12 hours of incubation when glucose was
drastically utilized as both carbon and energy sources. This was followed by a slight
decreased of growth by both isolates within 50 hours period of incubation prior to the
second phase of growth. This resulted to a maximum biomass of 1.6 times higher
(4.99 g/L) compared to those obtained in the medium supplemented with glucose as
the sole source of carbon (3.06 g/L). It was possible to assume that the second phase
of growth by both isolates in the mix culture system occurred as a result of crude oil
utilization as an alternative carbon source due to the depletion of glucose in the
growth medium. Increased in biomass in the second phase was than followed by its
decreased due to death phase. The toxicity of hydrocarbon or its related compounds
to microbial cells has been widely reported elsewhere [Edwards et al., 2003].
The production of biosurfactant by the bacterial mix culture systems was
found associated to cell growth (Figure 5.11) similar to those observed in the pure
culture system (Section 5.3.3). Drastic increased in biosurfactant production was
obtained during the maximum growth of both isolates in both mix culture systems.
This was followed by a gradual increased of biosurfactant concentration until the
maximum production of biosurfactant obtained within 72-120 hours of incubation.
In general, the maximum production of biosurfactant by bacterial mix culture
system 1:1 in glucose medium (Pmax = 6.36 g/L) was found similar to those obtained
in glucose + crude oil medium (Pmax = 6.75 g/L). Table 5.5 summarized the kinetic
analysis of growth and biosurfactant production for the bacterial mix culture system
1:1 grown in the medium supplemented with either glucose or both glucose and
crude oil.
82
Table 5.5: Kinetic analysis of growth and biosurfactant production for bacterial mix
culture system 1:1 (AB-Cr1: ETL-Cr1) grown at 37ºC, in medium
supplemented with either glucose or both glucose and crude oil.
Carbon sources
µ (h-1)
Xmax (g/L)
Pmax (g/L)
Productivity (g/L/h)
Yp/s (g/g)
Yx/s (g/g)
Yp/x (g/g)
qp (g/g/h)
ST max (mN/m)
IFT max (mN/m)
Glucose
Glucose + Crude Oil
0.243
3.06 @ 7h
6.36 @ 72h
0.09
12.09
30.32
1.48
0.36
13
9
0.096
4.99 @ 72h
6.75 @ 120h
0.06
14.70
11.42
1.29
0.12
24
10
In general, both isolates were able to grow well as a mixture in medium
supplemented with glucose and crude oil. The presence of crude oil in the medium
resulted to high biomass (~ 5.0 g/L) produced by both isolates in the mix culture
system in comparison to those grown in the medium supplemented with glucose
only. This result was in contrast to those obtained in single culture of both isolates
(Section 5.3.3), which high biomass were produced in the medium supplemented
with glucose as the sole source of carbon. It was possible to assume that the presence
of both isolates as a mixture in the growth medium might have a great interest in
increasing the direct physical interaction of the bacterial cells with the hydrophobic
substrate in order to initiate the crude oil degradation. It has been indicated earlier
using BATH test that both isolates have different capacity to utilize hydrocarbon
therefore they might have been interacted in such a way to benefit one another
[Rosenberg, 1984; Zhang and Miller, 1994].
The yield of biomass based on substrate consumption (Yx/s) was found higher
for the bacterial mix culture system 1:1 grown in glucose medium (30.32 g/g) than
those obtained in the system grown in glucose + crude oil medium (11.42 g/g).
However, the yield of biosurfactant production based on the substrate consumption
(Yp/s) and based on the maximum biomass (Yp/x) was not significantly different in the
mix culture system, grown in medium supplemented with either glucose or both
glucose and crude oil.
83
Comparisons between Yx/s and Yp/s values indicated that substrate utilization
was slightly more efficient for biosurfactant production compared to biomass in the
medium supplemented with both glucose and crude oil. However, the presence of
glucose as the sole source of carbon was able to increase the specific rates of
biosurfactant production (qp) by almost three times.
The maximum reduction of surface tension was found two times higher in the
bacterial mix culture system grown in medium supplemented with both glucose and
crude oil compared to those grown in the medium supplemented with glucose only
(Table 5.5). The presence of hydrocarbon in the mix culture system was found to
deliver a great effect in their surface activity. According to Lin et al. (1993), it is
possible that the inactivation of the biosurfactant is not the result of degradation, but
is caused by the binding of the molecules in the form of micelle, which reduce its
amphiphilic character and its ability to lower the surface tension.
The present study showed that the bacterial mix culture system 1:1 grown in
medium supplemented with both glucose and crude oil was able to produce
biosurfactant at a slightly higher concentration compared to the system grown in
medium supplemented with glucose only (Table 5.5). However, the efficiency of this
system to reduce surface tension (24mN/m reduction) of the medium was not as good
as those obtained in the single culture of AB-Cr1 isolate grown in the medium
supplemented with glucose as the sole source of carbon (32mN/m reduction). Thus,
further study will be carried out to study the production of biosurfactant by strain
AB-Cr1 in a bioreactor (Chapter 6).
CHAPTER 6
PRODUCTION OF BIOSURFACTANT BY STRAIN AB-Cr1 IN
BIOREACTOR
6.1
Introduction
At present, biosurfactants have not yet been employed extensively in industry
due to many reasons. The complexity of fermentation broth compositions, a
relatively low productivity of biosurfactant and poor product recovery process were
amongst the many reasons which sat the biosurfactant as only an alternative to the
chemical surfactant [Deleu and Paquot, 2004]. In general, it was necessary to
optimize the conditions for biosurfactant production because the quality and the
quantity of biosurfactants were strongly dependant on factors, such as fermentor
design, pH, temperature, nutrient composition and types of substrate used [Mulligan
and Gibbs, 1993]. It has also been reported that the induction of biosurfactant
production was best observed in the nitrogen-limited medium [Desai and Banat,
1997] though the truth of this statement would be tested in this study, using a locally
isolated bacterial strain AB-Cr1.
In the present work, the strain AB-Cr1 has been observed with its best
biosurfactant-producing ability (Chapter 5), and was selected for further study in a
bioreactor. Some physiological conditions for biosurfactant production were
investigated in a bioreactor as a factor of temperature (Section 6.2.1.1), initial
glucose concentration (Section 6.2.1.2), controlled pH (Section 6.2.1.3) and initial
NH4NO3 concentration (Section 6.2.1.4).
85
6.2
Methodology
6.2.1
Batch Fermentation
Small-scale batch fermentations were carried out in a 2-liter Biostat B
fermentor (B.Braun Biotech International GmbH) with a working volume of 1.5 liter.
The cultures were stirred at 200rpm, initially saturated with 100% oxygen and
continually aerated at 0.5vvm. The bioreactor was equipped with the temperature,
pO2 and pH controller system.
For inoculation, an overnight culture (Section 3.3.1) grown in Ramsay
medium (Section 3.2.1.1) was added to the fermentor to a final concentration of 10%
v/v.
6.2.1.1 Effect of Temperature on Biosurfactant Production.
The influence of temperature on growth and biosurfactant production by ABCr1 isolate was studied using Ramsay medium (Section 3.2.1.1), supplemented with
3 mM glucose and adjusted to initial pH 7. Temperature was varied at 30º, 37º, 40º
and 45ºC during each batch of fermentation. All experiments were carried out in
duplicate and those with significantly similar results were recorded and presented as
their average values, respectively. Samples were periodically taken over 4 days of
fermentation. Approximately, 40 mL aliquots of whole broth were sampled and
centrifuged at 5000 rpm at 4ºC for 20 minutes prior to analyze for glucose (Section
3.4.2), surface and interfacial tension (Section 3.4.3.1-3.4.3.2) as well as
biosurfactant production (Section 3.5.1-3.5.2). Growth was monitored by measuring
the optical density of the 3 mL aliquots of culture broth at 600 nm (Section 3.4.1.1)
and the bacterial cell dry weight was determined as mentioned in section 3.4.1.2. The
specific growth rates (µ) of culture were then calculated based on plots of ln [cell dry
weight] versus time (hour). Changes of the level of oxygen and pH in the medium
were recorded accordingly as displayed on the controller unit, respectively.
86
6.2.1.2 Effect of Initial Glucose Concentration on Biosurfactant Production
Production of biosurfactant by AB-Cr1 isolate was performed with various
initial concentrations of glucose (0, 3, 5, 10, 20 mM). All experiments were carried
out in duplicate using similar set up described in Section 6.2.1 except that the initial
concentration of glucose was varied between each batch. In addition, the optimum
temperature obtained (Section 6.2.1.1) was also used throughout this experiment.
Samples were taken out at regular intervals to analyze for growth (Section 3.4.1.13.4.1.2), glucose (Section 3.4.2), surface and interfacial tension (Section 3.4.3.13.4.3.2) as well as biosurfactant production (Section 3.5.1-3.5.2). Changes of the
level of oxygen and pH in the medium were recorded accordingly as displayed on the
controller unit, respectively. The growth rates of cultures were calculated as
mentioned in section 6.2.1.1.
6.2.1.3 Effect of Controlled pH Condition on Biosurfactant Production
The effect of hydrogen ion concentration on the production of biosurfactant
was studied. The experiment was performed by inoculating the AB-Cr1 isolate (10%
v/v) in Ramsay medium (Section 3.2.1.1) supplemented with the optimum initial
glucose concentration (Section 6.2.1.2) and incubated at the optimum temperature
(Section 6.2.1.1). The pH of growth medium was maintained at 6.5, 7.0 and 7.5 (each
was in duplication) by automatically adding either 1 N HCl or 1 N NaOH solutions
into the bioreactor according to the signal received from the pH controller unit.
Samples were taken out at regular intervals to analyze for growth (Section 3.4.1.13.4.1.2), glucose (Section 3.4.2), surface and interfacial tension (Section 3.4.3.13.4.3.2) as well as biosurfactant production (Section 3.5.1-3.5.2). Changes of the
level of oxygen and pH in the medium were recorded accordingly as displayed on the
controller unit, respectively. The growth rates of culture were calculated as
mentioned in section 6.2.1.1.
87
6.2.1.4 Effect of Initial NH4NO3 Concentrations on Biosurfactant Production
The same experimental set up (Section 6.2.1) was used in this experiment
using Ramsay medium with yeast extract excluded, supplemented with the optimum
initial glucose concentration (Section 6.2.1.2) and adjusted to the optimum pH
(Section 6.2.1.3). The initial NH4NO3 concentrations in the Ramsay medium were
varied from 0 to 25 mM. The medium was inoculated with AB-Cr1 isolate (10% v/v)
and incubated at the optimum temperature (Section 6.2.1.1). Samples were taken out
at regular intervals to analyze for growth (Section 3.4.1.1-3.4.1.2), glucose (Section
3.4.2), surface and interfacial tension (Section 3.4.3.1-3.4.3.2) as well as
biosurfactant production (Section 3.5.1-3.5.2). Changes of the level of oxygen and
pH in the medium were recorded accordingly as displayed on the controller unit,
respectively. The growth rates of culture were calculated as mentioned in section
6.2.1.1.
6.3
Results and Discussion
6.3.1 Effect of Temperature on Biosurfactant Production
Temperature was one of the major factors that would give direct effects on
bacterial metabolism and thus affecting its producing ability, particularly of primary
metabolites [Guerra-Santos et al., 1986]. The production of biosurfactant was
claimed to be sensitive to the changes of incubation temperature. It has also been
reported that temperature could significantly affected the yield of biosurfactant, as
well as to alter the composition of biosurfactant produced [Desai and Desai, 1993].
The effect of various temperatures on the maximum production of
biosurfactant and biomass of AB-Cr1 was as showed in Figure 6.1.
88
14
Xmax / Pmax , g/L
12
10
8
6
4
2
0
30
37
40
o
Temperature, C
45
Fig. 6.1: Maximum cell biomass ( ) and biosurfactant ( ) production by AB-Cr1
grown in medium supplemented with 3mM glucose, as a factor of
temperature.
The optimal temperature for growth and biosurfactant production of isolate
AB-Cr1 was found in the culture incubated at 37ºC (Figure 6.1). It was indicated by
the highest production of biosurfactant (P max = 12.45 g/L) and biomass (Xmax = 6.48
g/L) obtained in the culture compared to those incubated at 30º, 40º and 45ºC,
respectively. The maximum specific growth rate of 0.390 h-1 was also significantly
faster than those incubated at other temperature tested (0.158 – 0.224 h-1). This result
was in good agreement with that obtained in shake flasks study (Section 5.3.2).
This isolate was also found more affected by temperature higher than 37ºC
compared to that showed at the lower temperature (30ºC). This was indicated by the
lower specific growth rates (by approximately 1.6 times) in culture incubated at 40º
and 45ºC as well as 3 times decreased in biomass production. The minimum
production of biosurfactant was also obtained (8-9 g/L) in the cultures incubated at
the respective temperatures. Decreased of microbial activities was claimed due to the
high maintenance energy required for repair mechanisms activated as a result of the
thermal denaturation of proteins. At the lower temperature, the rate of protein /
enzyme denaturation was negligible in mesophilic, however cells were affected by
the diffusional limitation of solutes such as substrates into and within the cell
[Scragg, 1988]. As a result, the biomass and biosurfactant yield changes at lower or
higher temperature than the optimum. Thus, at extreme incubation temperatures it
89
would also led to some changes in the microbial metabolism as expressed by lower
production of biosurfactant [Guera-Santos et al., 1986].
The correlation between biosurfactant production and growth, substrate and
oxygen consumption and pH changes were as illustrated in Figure 6.2(A) and (B).
Similar to that observed in the shake flask study (Chapter 5), biosurfactant
production was growth-associated in all conditions tested. Therefore, this experiment
also indicated that temperature was on its role to affect only the productivity of
biosurfactant by isolate AB-Cr1 and not resulted in changes to the type of metabolite
it was. Temperature was observed to play its role in changing the pattern of
biosurfactant production from primary to secondary metabolites in many
thermophilic bacteria [Banat, 1993].
100
14
12
60
8
6
40
pO2, %
Biomass &
Biosurfactant, g/L
80
10
4
20
2
0
0
10
20
30
40
50
60
Time, h
70
80
90
0
100
(A)
7
0.4
6.5
pH
[Glucose], g/L
0.3
0.2
6
0.1
0
0
20
40
60
Time, h
(B)
80
5.5
100
90
Surface Activity, mN/m
70
60
50
40
30
20
10
0
0
20
40
Time, h
60
80
100
(C)
Fig. 6.2: Relationship between biosurfactant production
, growth
and
oxygen consumption
(A), glucose consumption
and pH
(B), surface
, interfacial
and spreading tension
(C) by ABCr1, grown in medium supplemented with 3mM glucose adjusted to initial
pH 7.0 and incubated at 37ºC.
Drastic increased in biosurfactant concentration and biomass were also found
related to the maximum consumption of glucose and oxygen (Figure 6.2(A) and (B)).
During the first 5 hours incubation, the concentration of biosurfactant and biomass
increased exponentially with drastic reduction of glucose (34%) and oxygen (70%)
concentrations in the growth medium. Total oxygen consumption was observed in
parallel to the consumption of glucose within 10 hours of incubation. At this point,
oxygen was totally used as the end terminal electron acceptor as the rate of microbial
metabolism was at the maximum utilizing the available substrate for growth and
biosurfactant production. Similar correlation between growth and the consumption of
substrate and oxygen was commonly observed in many aerobic microorganisms
[Guerra-Santos et al., 1986]. The pH of the culture medium was reduced to 5.7
during this phase, as a result of weak acid production during glucose catabolism.
The maximum production of biosurfactant (12.45 g/L) was observed during
48 hours incubation at 37ºC. During this point, the pH was observed slightly
increased from 5.7 to 6.3 and the oxygen level was also increased and stabilized at
the level of ~ 70%. The second phase production of biosurfactant observed with ABCr1 grown in glucose was also in common with some other bacteria from the genus
91
of Bacillus and Pseudomonas [Kluge et at., 1988; Mata-Sandoval et al., 2000]. It was
claimed that increased in the concentration of biosurfactant during the stationary
growth phase has some relation to the consumption of organic acid produced during
active catabolism of glucose [Lang and Wagner, 1993]. Therefore, this might
explained the increased of pH observed during the maximum production of
biosurfactant at 48 hours incubation. Low consumption of oxygen was possibly due
to low metabolism rate of the organic acid(s) as an alternative source of energy and
carbon [Hommel and Ratledge, 1993]. There was gradual reduction in biosurfactant
concentration observed after 48 hours incubation. According to Lin et al. (1993), the
observed disappearance of biosurfactant might be related to the development of
competence. There are 3 possible mechanisms responsible for the decline in the
biosurfactant concentration in stationary phase i.e. (i) the biosurfactant was degraded
by the enzymes in the culture, or (ii) the biosurfactant might be adsorbed on the cell
surface, or (iii) it was reinternalized and processed intracellularly.
The greatest reduction in surface and interfacial tension of the medium
controlled at 37ºC were also observed during the log phase of cell growth. Surface
tension dropped from 69.5mN/m to 31.5mN/m whereas the interfacial tension
dropped from 18mN/m to 4.5mN/m during the end of log phase (within 8 hours of
incubation) and stabilized over the next 40 hours to a value of 38mN/m and 9mN/m
for surface and interfacial tension, respectively (Figure 6.2(C)). Such interfacial
tension values are considered significant for oil mobilization [Li et al., 1984]. This
result also supported the argument that claimed the production of biosurfactants has
an important role in microbial enhance oil recovery [Banat et al., 2000].
On the other hand, the maximum surface and interfacial tension reduction of
the medium controlled at 30ºC also showed quite an acceptable values compared to
the cell-free culture grown at 40ºC and 45ºC (Figure 6.3). The biosurfactant
recovered was also comparably high (P max = 11.57 g/L) though it was not as high as
the amount of biosurfactant obtained from culture at 37ºC. During this point, the
surface and interfacial tension of the cell-free culture at 30ºC was dropped to
37.5mN/m and 8mN/m, respectively.
92
Surface Activity Reduction, mN/m
40
35
30
25
20
15
10
5
0
30
37
40
Temperature, o C
45
Fig. 6.3: Surface ( ) and interfacial tension ( ) reduction of the cell-free culture of
AB-Cr1, grown in medium supplemented with 3mM glucose, as a factor of
temperature.
The ability of biosurfactant to form stable emulsion was also determined by
the calculation of its spreading tensions (Section 3.4.3.3). Biosurfactant from the
cultures of 30º and 37ºC was found able to form a very stable emulsion as the value
of spreading tension was significantly low (< 10mN/m) throughout the fermentation
period. Therefore, this suggested that this bacterium was able to create the emulsion
condition necessary for efficient uptake of hydrocarbon substrate when cells were
grown on such substrate [Ramsay et al., 1983].
Considering the complete consumption of glucose supplied and higher
production of biosurfactant compared to biomass, it was possible to suggest the
efficiency of substrate consumption for biosurfactant production (Yp/s = 33.92 g/g)
was much higher than that for biomass production (Yx/s = 23.46 g/g) at the incubation
temperature of 37ºC. This was also observed at other temperatures tested (Figure
6.4). It might indicated that the carbon source supplied was mostly used for
biosurfactant production compared to the cell growth. Thus, a temperature of 37ºC
was chosen for further experiment to determine the effect of initial glucose
concentration on biosurfactant production by isolate AB-Cr1.
93
35
Yield Coefficients, g/g
30
25
20
15
10
5
0
30
37
40
Temperature, o C
45
Fig. 6.4: The yield coefficients for biosurfactant and biomass production by AB-Cr1
grown in medium supplemented with 3mM glucose, as a factor of
temperature. Symbol: Yp/s , Yx/s and Yp/x .
6.3.2
Effect of Initial Glucose Concentrations on Biosurfactant Production
Glucose as a source of carbon could be an important key to regulate
biosurfactant synthesis [Desai and Desai, 1993]. There were many evident on the
importance of carbon and its concentration in the production of surface-active
compound by microbes [Desai and Banat, 1997]. This section would be focused on
the effects of different initial substrate concentrations to the production of
biosurfactant based on the reduction in surface and interfacial tension as well as the
dry weight of the biosurfactant produced.
Figure 6.5 showed the maximum cell biomass and biosurfactant production as a
result of growth on various initial glucose concentrations for isolate AB-Cr1.
94
14
Xmax / Pmax, g/L
12
10
8
6
4
2
0
0
3
5
[Glucose], mM
10
20
Fig. 6.5: Maximum biomass ( ) and biosurfactant ( ) production by AB-Cr1
grown at 37ºC, as a factor of various initial glucose concentrations.
As shown, biosurfactant production was strongly affected by the initial
concentration of glucose added to the medium. Amongst all the glucose
concentrations tested, the addition of 3mM glucose was found adequate to stimulate
the maximum production of biosurfactant (12.45 g/L). Whereas, culture of AB-Cr1
grown in the presence of 20mM glucose caused a highly significant induction for the
production of maximum biomass instead of biosurfactant. No significant different
was observed in the culture supplemented with 5 and 10mM glucose. However,
reduction of biosurfactant production was very significant (~ 40% reduction based on
the maximum Pmax (12.45 g/L) in the presence of 3mM glucose). Only 10.31 g/L of
biosurfactant produced in the presence of 20mM glucose compared to that observed
in the medium supplemented with 3mM glucose.
It was also indicated that, the excess of glucose in the medium (above 10mM
glucose) would induced for cell growth better than for the production of
biosurfactant. In the presence of 5 and 10mM glucose, the efficiency of glucose
utilization for both biomass and biosurfactant production was found similar. This was
indicated by the similar amount of both obtained from the respective culture as well
as their lower Yp/s and Yx/s values (Table 6.1).
95
Table 6.1 summarized the kinetic analysis performed for the growth and
biosurfactant production by isolate AB-Cr1 grown in the medium supplemented with
various initial concentration of glucose.
Table 6.1: Kinetic analysis for growth and biosurfactant production by AB-Cr1
grown at 37ºC, in medium supplemented with various initial glucose
concentrations.
[Glucose]
mM
0
3
5
10
20
µ (h-1)
Xmax (g/L)
0.0988
1.59 @ 5h
0.3901
6.48 @ 7h
0.2189
5.5 @ 10h
Pmax (g/L)
2.69 @
60h
0.04
12.45 @
48h
0.26
6.98 @
50h
0.14
0.1629
6.24 @
16h
6.92 @
22h
0.31
0.1609
13.25 @
48h
10.31 @
48h
0.21
1.59
2.69
1.69
4.2
33.92
23.46
1.92
38
8.09
6.79
1.27
31.5
3.89
3.83
1.11
28.5
2.96
3.81
0.78
33
6.5
13.5
13
11
11.5
Productivity
(g/L/h)
Yp/s (g/g)
Yx/s (g/g)
Yp/x (g/g)
ST max
(mN/m)
IFTmax
(mN/m)
The efficiency of biosurfactant production using glucose as the sole source of
carbon and energy was also supported by its higher value of Yp/s (33.92 g/g) in the
medium with 3mM glucose added compared to those in the medium supplemented
with higher concentration of glucose (Yp/s < 10 g/g). From the Yx/s (23.46 g/g) value
obtained based on biomass production in the presence of 3mM glucose, it was also
possible to suggest that the utilization of glucose in this culture was more efficient
for the production of biosurfactant compared to biomass. The Yx/s value was
approximately 1.5-fold lower than the Yp/s value. In the presence of 20mM glucose,
both Yp/s and Yx/s values were approximately 11.5 and 6.2-fold lower than those
obtained in the medium added with 3mM glucose. In addition, it was also possible to
suggest that in the glucose rich medium, the efficiency of biomass production was
better than for the production of biosurfactant. This was indicated by the lower Yp/s
value (2.96 g/g) compared to Yx/s (3.81 g/g). It has also been reported that in a
condition where excess substrate was available, higher yield of biomass would be
96
expected in comparison with the production of biosurfactant [Brakemeier et al.,
1998]. This was also as a result to the inhibition of biosurfactant production due to
substrate saturation [Lang and Wagner, 1993]. In addition, biosurfactant production
was commonly induced in the limiting growth conditions as a factor of substrate and
nitrogen concentrations [Desai and Banat, 1997].
Slight differences in the maximum cell biomass and biosurfactant production
could be observed as the initial glucose concentration increased above the optimum
level [Guerra-Santos et al., 1986]. Should the initial substrate concentration was
increased to a value considerably higher than the minimum saturating concentration,
the growth rate of microbes might began to fall due to substrate inhibition, which
may caused by the high osmotic stress imposed on the cells resulted in cell
dehydration and followed by diffusion problems [Scragg 1988].
From the results, it was also observed that the maximum production of
biosurfactant in 3mM glucose was also resulted to the maximum reduction of surface
and interfacial tension. This was in contrast to the biosurfactant produced in the
medium with other concentration of glucose tested which reduction of both surface
and interfacial tension were lower. Both growth and biosurfactant productions were
dependent on the presence of glucose as carbon and energy sources and there was
slow growth and less biosurfactant was produced in the medium not supplemented
with glucose. Carbon source particularly plays an important role in determining the
pathways involved in biosynthesis of biosurfactant [Mulligan and Gibbs, 1993] either
by induction or repression [Cameotra and Makkar, 1998]. It was suggested that both
the lipogenic pathway and the formation of sugar portion would regulate a sugar lipid
type of surfactant synthesized from a carbohydrate by glycolytic metabolism. In
addition, it might also indicated to whether the biosurfactant produced was
extracellular or intracellular [Fiechter, 1992].
Since high initial glucose concentrations (5 - 20mM) resulted in the lower
yields of biosurfactant, results suggested that the addition of 3mM glucose was
adequate to obtain an efficient production of biosurfactant by isolate AB-Cr1.
Therefore, further experiment will be conducted using the medium supplemented
with 3mM glucose and incubated at 37ºC.
97
6.3.3
Effect of Controlled pH Condition on Biosurfactant Production
This experiment was conducted in order to assess whether biosurfactant
production could be further enhanced using controlled pH strategy during
fermentation. Therefore, changes of pH during glucose metabolism in the growth
medium would be automatically corrected by the addition of either acid or alkali. In
this study, pH of the medium was controlled at 6.5, 7.0 and 7.5, respectively. It has
been reported that pH played an important role in affecting biosurfactant production
through their effects on cellular growth and metabolic activity [Desai and Banat,
1997].
Figure 6.6 showed the effects of pH controlled strategy on the production of
biosurfactant and growth of isolate AB-Cr1.
8
Xmax / Pmax, g/L
6
4
2
0
6.5
7
pH
7.5
Fig. 6.6: Maximum biomass ( ) and biosurfactant ( ) production by AB-Cr1
grown in medium supplemented with 3mM glucose at 37ºC, as a factor of
pH.
Results indicated that the pH controlled strategy was not successful to
enhance the production of biosurfactant. This was showed by the significantly lower
concentration of biosurfactant obtained from the pH-controlled experiments
compared to that in the culture which pH was not controlled (Section 6.3.2). The
maximum biosurfactant production of 7.62 g/L observed in the culture controlled at
pH 7.0 was found approximately 1.6-fold lower to that obtained in the non-pH
98
controlled system (12.45 g/L). At the higher pH (pH 7.5), biosurfactant production
was found reduced to 6.32 g/L. At the pH higher than the optimum, microbial
metabolism was regulated for its survival and more energy would be channeled for
biomass production, thus reduction of biosurfactant production occurred [GuerraSantos et al., 1986].
However, this was in contrast to that observed in the culture controlled at pH
6.5, which indicated to the more efficient production of biomass compared to
biosurfactant. The maximum biomass produced (6.31 g/L) was approximately two
times higher than that observed in pH 7.0 culture (3.51 g/L). It was possible to
suggest that the change of physiological parameters such as temperature and pH
would significantly caused alteration of the enzymatic activity of microbes. Increased
or decreased in pH / temperature might affect substrate and oxygen uptake which was
related directly or indirectly with the regulation of enzymes production for
biosynthesis of microbial surfactant and biomass [Hommel and Ratledge, 1993]. In
other study by Guerra-Santos et al. (1986) the claimed that controlled pH condition
would cause accumulation of significant amount of organic acid from glucose
catabolism throughout fermentation period. This might result to alteration of
membrane permeability of the cell which could also led to toxicity in relation with
the accumulated organic acid. Therefore, microbial production of biosurfactant
would be suppressed significantly [Hommel and Ratledge, 1993].
The highest reduction of surface and interfacial tensions observed in the
culture controlled at pH 7.0 (Figure 6.7) was also in correlation with the highest
concentration of biosurfactant obtained in the system. It has also been reported that
the higher concentration of biosurfactant presence in the solution could be indicated
by the higher reduction of surface tension obtained in comparison with a solution
containing less biosurfactant [Hua et al., 2003]. The reduction of interfacial tension
indicated to the possible efficiency of biosurfactant for particular application such as
for hydrocarbon degradation, increasing oil mobility in MEOR process, pipeline
cleaning and etc. [Banat et al., 2000]. The ability of the biosurfactant to reduce the
interfacial tension up to 8.2mN/m suggested that it was acceptable for industrial
cleaning application of oily surfaces and hydrocarbon degradation [Mulligan, 2004].
Figure 6.7 showed the reduction of surface and interfacial tension by the
biosurfactant produced under controlled pH condition. Although production of
99
biosurfactant at controlled pH 7.5 was relatively low compared to biosurfactant
produced at pH 7.0, the surface and interfacial tension reduction of the growth
medium at pH 7.5 showed a significant values in producing a relatively good
biosurfactant (>20mN/m and >10mN/m for surface and interfacial tension reduction,
respectively). Based on this fact, the biosurfactant produced by the culture controlled
at pH 6.5 was not considered as good biosurfactant for applications.
Surface Activity Reduction, mN/m
40
35
30
25
20
15
10
5
0
6.5
7
pH
7.5
Fig. 6.7: Surface tension ( ) and interfacial tension ( ) reduction of the cell-free
culture of AB-Cr1, grown in the medium supplemented with 3mM glucose
at 37ºC, as a factor of pH.
However, the surface and interfacial tension of the culture medium were
found increased at the later phase of growth (Figure 6.8(B)). The changes of surface
activity were closely related to the changes in biosurfactant concentration presence in
the culture [Hua et al., 2003]. There were two possible explanations for the
increment of surface and interfacial tension observed, i.e: (i) declined in the
concentration of biosurfactant due to its degradation via microbial metabolism, and
(ii) the production of other soluble metabolic by products which interfere with the
measurement of both surface and interfacial tension [Lin et al., 1993].
100
100
9
8
80
6
60
5
4
40
pO2, %
Biomass &
Biosurfactant, g/L
7
3
2
20
1
0
0
20
40
Time, h
60
80
0
100
(A)
70
Surface Activity, mN/m
60
50
40
30
20
10
0
0
20
40
60
80
100
Time, h
(B)
Fig. 6.8: The relationship between biosurfactant production
, growth
and
oxygen consumption
(A), surface
and interfacial tension
(B) by AB-Cr1 grown in medium at controlled pH 7.0 and incubated at
37ºC.
Figure 6.8(A) showed the relationship between biosurfactant production,
growth and consumption. Similar to those observed in the previous experiment
(Chapter 5), the production of biosurfactant was directly proportional to cell growth,
indicated to primary metabolite. Similar pattern of growth, biosurfactant production
and oxygen consumption were observed to those discussed in Section 6.3.1.
However, this result was in contrast due to the lower maximum production of
biosurfactant obtained in the pH controlled system. Increased in biosurfactant
101
production during the stationary growth phase was found less than that observed in
the culture which pH was not controlled (Figure 6.2 (A)). This might also related to
the inability of the cells to consume organic acid produced from glucose catabolism
which resulted to limited production of biosurfactant [Hommel and Ratledge, 1993].
Table 6.2 summarized the kinetic analysis for growth and biosurfactant
production by isolate AB-Cr1 in the effect of pH control.
Table 6.2: Kinetic analysis for growth and biosurfactant production by AB-Cr1
grown in the medium controlled at various pH values, supplemented
with 3mM glucose and incubated at 37ºC.
pH
6.5
7.0
7.5
µ (h-1)
Xmax (g/L)
Pmax (g/L)
Productivity (g/L/h)
Yp/s (g/g)
Yx/s (g/g)
Yp/x (g/g)
ST max (mN/m)
IFTmax (mN/m)
0.378
6.31 @ 5h
5.57 @ 60h
0.09
15.82
53.3
0.88
17.3
11
0.292
3.51 @ 8h
7.62 @ 48h
0.16
20.76
53.37
2.17
37.8
10.5
0.257
5.07 @ 72h
6.32 @ 84h
0.08
17.67
14.18
1.25
32.5
10.8
From this analysis, it was found that the condition best for microbial growth
was not necessarily supported for the highest biosurfactant production under
controlled pH condition. The best condition for growth under controlled pH
condition was at pH 6.5 based on the highest specific growth rate obtained (0.378 h1
). It was observed that both yield coefficient and productivity of biosurfactant by the
culture controlled at pH 7.0 were higher than that those observed in other pH tested.
The productivity of biosurfactant at pH 7.0 reached to value of 0.16 g/L/h, which is
almost two times than that the productivity of biosurfactant at pH 6.5 and 7.5.
However, this was far lower than that productivity observed in the culture which ph
was not controlled (Table 6.1).
The efficiency of biomass and product (biosurfactant) formation based on
substrate consumed were as showed in Table 6.2. It was observed that the yield of
biosurfactant production based on the substrate consumption (Yp/s) was found higher
in the medium at controlled pH 7.0 (20.76 g/g) than those at other controlled pH
(~15-18 g/g). However, the yield of biomass based on substrate consumption (Yx/s)
102
was not significantly differed by the culture controlled at pH 7.0 and 6.5 (Yx/s = ~53
g/g). The Yx/s value was found lower by the culture controlled at pH 7.5 (Yx/s = 14.18
g/g) as indicated by the maximum biomass obtained at the longer fermentation period
(t = 72 hours).
6.3.4
Effect of Initial NH4NO3 Concentrations on Biosurfactant Production
Nitrogen whether it is organic or inorganic forms were very important in the
cellular metabolism thus affecting its producing ability of surface-active compounds.
It has been reported that NH4NO3 could be the best nitrogen source for the
production of biosurfactant by facultative aerobes [Desai and Banat, 1997]. It is very
water-soluble, therefore could be easily supplied to the cell. So this section was
focused to study the production of biosurfactant in medium added with various
concentrations of NH4NO3, supplemented with 3mM glucose, adjusted to initial pH
7.0 and incubated at 37ºC. Yeast extract which contributed to organic nitrogen source
was excluded from the medium.
The effect of various NH4NO3 concentrations on the maximum production of
biosurfactant and biomass of AB-Cr1 was as showed in Figure 6.9.
In general, the exclusion of yeast extract from the medium was not effective
for enhanced production of biosurfactant by isolate AB-Cr1. This was indicated by
the lower Pmax value (~ 4-6.5 g/L) obtained compared to that observed in the
presence of both yeast extract and NH4NO3 (Pmax = 12.45 g/L) as nitrogen sources.
Amongst all the NH4NO3 concentrations tested, the addition of 15mM NH4NO3 was
found adequate to stimulate the maximum production of biosurfactant (6.47 g/L).
Whereas, culture of AB-Cr1 grown in the presence of 25mM NH4NO3 caused a
significant reduction of biosurfactant production (5.54 g/L). However, increased in
NH4NO3 concentration would significantly stimulated growth of the bacterium. This
result was in good agreement with Sudhakar et al. (1996) that excessive nitrogen
concentration might induce the growth of bacteria, though the production of
biosurfactant was suppressed.
103
7
Xmax / Pmax, g/L
6
5
4
3
2
1
0
0
5
15
25
[NH4NO3], mM
Fig. 6.9: Maximum biomass ( ) and biosurfactant production ( ) by AB-Cr1
grown in medium supplemented with 3mM glucose at 37ºC, as a factor of
various initial NH4NO3 concentrations.
It was also indicated that, the efficiency of biosurfactant producing ability by
isolate AB-Cr1 was found similar in the presence of 5 and 25mM NH4NO3.
However, biosurfactant production was significantly halted in the nitrogen-free
medium (~ 4 g/L), thus indicated to the requirement for nitrogen supply. It was
observed that both biomass and biosurfactant production were dependent on the
availability of NH4NO3 supplied into the medium, or else halted in its absence. The
dependency of isolate AB-Cr1 on NH4NO3 for biomass and biosurfactant production
was further confirmed by the very limited growth and biosurfactant production (Xmax
= 1.05 g/L and Pmax = 3.81 g/L) observed in the nitrogen-free medium (Figure 6.9).
According to Cameotra and Makkar (1998), nitrogen in the form of NH4NO3 played
an important role in the biosynthesis of protein in the cells, as well as in the
regulation of biosurfactant synthesis.
The correlation between biosurfactant production, growth and oxygen
consumption, surface and interfacial tension were as illustrated in Figure 6.10(A) and
(B).
104
100
7
80
5
4
60
3
40
pO2, %
Biomass &
Biosurfactant, g/L
6
2
20
1
0
0
20
40
60
80
0
100
Time, h
(A)
70
Surface Activity, mN/m
60
50
40
30
20
10
0
0
20
40
60
80
100
Time, h
(B)
Fig. 6.10: The relationship between biosurfactant production
, growth
and
oxygen consumption
(A), surface
and interfacial tension
(B), by AB-Cr1 grown in medium supplemented with 15mM NH4NO3
and incubated at 37ºC.
The production of biosurfactant by isolate AB-Cr1 was found associated to
cell growth. There was a paralleled relationship between the cell growth and the
biosurfactant production. According to Desai and Desai (1993), nitrate as a nitrogen
source caused the production of biosurfactant during the exponential growth phase,
whereas the ammonium caused growth-associated biosurfactant production.
105
Drastic increased in biosurfactant concentration and biomass were also found
related to the maximum consumption of oxygen (Figure 6.10(A)). During the first 10
hours incubation, the concentration of biosurfactant and biomass increased
exponentially with drastic reduction of oxygen concentration followed by its
depletion in the growth medium. At this point, oxygen was totally used as the end
terminal electron acceptor as the rate of microbial metabolism was at the maximum,
utilizing the available substrate for growth and biosurfactant production.
It was also observed that the oxygen was totally used within the next 50 hours
incubation. High consumption of oxygen during this period might also due to high
metabolism rate of the organic acid(s) presence as an alternative source of energy and
carbon under glucose-limiting condition. Glucose was totally consumed within 10
hours of fermentation period. The maximum production of biosurfactant (6.47 g/L)
was observed during 60 hours incubation at 37ºC at which point the oxygen level
started to increase until 96 hours fermentation period. Increased in the oxygen was
directly related to decrease in biomass, thus less consumption of oxygen was due to
low metabolism rates [Guerra-Santos et al., 1986; Hommel and Ratledge, 1993].
The highest reduction in surface and interfacial tension of the medium were
also observed during the log phase of cell growth. Surface tension dropped from
70mN/m to 53.1mN/m whereas the interfacial tension dropped from 18.9mN/m to
14.1mN/m during the end of log phase (within 10 hours of incubation) and stabilized
over the next 70 hours to a value of 47.5mN/m and 10.7mN/m for surface and
interfacial tension, respectively (Figure 6.10(B)).
Figure 6.11 showed the yield coefficients for biosurfactant and biomass
production by isolate AB-Cr1 grown in the medium supplemented with various
initial NH4NO3 concentrations.
106
18
Yield Coefficients, g/g
16
14
12
10
8
6
4
2
0
0
5
15
25
[NH4NO3], mM
Fig. 6.11: The yield coefficients for biosurfactant and biomass production by ABCr1 grown in medium supplemented with 3mM glucose and incubated at
37ºC, as a factor of various initial NH4NO3 concentrations. Symbols: Yp/s
, Yx/s
and Yp/x .
Considering the complete consumption of glucose supplied and higher
production of biosurfactant compared to biomass, it was possible to suggest the
efficiency of substrate consumption for biosurfactant production (Yp/s = 16.37 g/g)
was much higher than that for biomass production (Yx/s = 8.73 g/g) in the medium
supplemented with 15mM NH4NO3. This was also observed at other NH4NO3
concentrations tested (Figure 6.11). The Yx/s value was approximately 1.3-1.9 times
lower than the Yp/s value. It might indicate that the carbon source supplied was
mostly used for biosurfactant production compared to the cell growth.
The efficiency of biosurfactant production was also supported by its higher
value of Yp/s (16.37 g/g) in the medium with 15mM NH4NO3 added compared to
those in the NH4NO3-free medium (Yp/s ~ 10 g/g). However, the greatest efficiency
of biomass production was observed in medium supplemented with 5mM NH4NO3
(Yx/s = 12.08 g/L).
Therefore, 15mM NH4NO3 was found to be the best concentration of
NH4NO3 (without the addition of yeast extract) for maximum production of
biosurfactant (6.47 g/L) by isolate AB-Cr1 grown in the medium supplemented with
3mM glucose as source of carbon and incubated at 37ºC. However, the amount of
biosurfactant produced was not as high as the amount of biosurfactant produced in
107
the medium supplemented with both yeast extract and 25mM NH4NO3 (Section
6.3.1). It was possible to suggest that for maximum biosurfactant production in the
medium supplemented with both NH4NO3 and yeast extract as the sources of nitrogen
would be more preferable. Yeast extract as an organic nitrogen source, showed a
significant effect on the production of biosurfactant by isolate AB-Cr1. This study
was also in contrast with those previously studied the conditions for biosurfactant
production that claimed the yeast extract-free nitrogen-limited condition would
significantly enhanced biosurfactant production [Haferbegr et al., 1986; Hommel and
Ratledge, 1993].
CHAPTER 7
CHARACTERIZATION OF CRUDE BIOSURFACTANT
7.1
Introduction
Several types of biosurfactants have been isolated and characterized based on
their biochemical nature and the microbial species producing them. They include
glycolipids, lipoproteins-lipopeptides, phospholipids, polysaccharide-protein
complexes, and those containing fatty acids and neutral lipids [Cooper, 1986; Jenny
et al., 1991]. Physicochemical properties such as surface tension reduction, stability
of the emulsion formed as well as pH and heat stability are very important in
searching for a potential biosurfactant for specific applications.
There are several studies of biosurfactants that both lowered surface tension
and stabilized emulsions [Cooper and Zajic, 1980; Cooper and Goldenberg, 1987]. A
preliminary analysis of the hydrophilic moiety of the biosurfactant produced by both
AB-Cr1 and ETL-Cr1 isolates could also lead to the identification of a complex
polymer of protein, carbohydrate and lipid. Whereas, the lipophilic portion of the
amphiphilic molecule was formed by fatty acids of different chain lengths and
characterized as their methyl ester.
This study was mainly focused on characterizing the crude biosurfactant
produced by AB-Cr1 and ETL-Cr1 isolates by several methods. The properties of the
crude biosurfactant obtained were studied because no further purification had been
carried out due to several limitations and time required to purify the biosurfactant.
109
7.2
Methodology
7.2.1
Emulsification Activity Tests
7.2.1.1 Assay of Emulsification
The emulsifying activity of the biosurfactant was determined by using the cellfree culture broth (Section 3.5.1) adjusted to pH 1, 4, 7, 10 and 13. The assay was
carried out by adding kerosene (3mL) to the sample fluid (3mL) in a test tube. The
tubes were then vigorously vortexed for 2 minutes and allowed to settle for 24 hours
before the percentage of volume occupied by the emulsion was determined. The
emulsification index, Ei was determined by the following equation [Cooper and
Goldenberg 1987];
Ei = Height of the emulsion layer
Total height
x 100
7.2.1.2 Assay of Emulsion Stability
The stability of emulsified solutions (Section 7.2.1.1) was allowed to stand
for 24 hours at room temperature. The absorbance was read at the wavelength of
540nm at 24 hours interval for 4 days. Stability of emulsions was expressed as decay
constant (Kd) obtained from the slope value of a plot between log [absorbance]
versus time (hour) [Kim et al., 2000].
7.2.2
Critical Micelle Concentration (CMC) Determination
Surface tension was determined as described in Section 3.4.3.1. The dried
extracted biosurfactant (Section 3.5.1-3.5.2) was dissolved in distilled water and
serially diluted (100-10-3) prior to the measurement of surface tension. The CMC was
obtained from a plot of the surface tension as a function of the biosurfactant
110
concentrations. The concentration at which micelles began to form was represented
as the CMC. Above this concentration, there would be no increment could be
detected for the reduction of surface tension.
7.2.3
Stability Studies
Stability studies were done using the cell-free culture broth by centrifugation
at 5000 rpm, 4ºC for 30 minutes. The cell-free broths were heated in a boiling water
bath (100ºC) for up to 70 minutes. Sample was drawn out at 10 minutes interval and
was cooled to room temperature prior to surface tension measurement (Section
3.4.3.1). To study the pH stability of the biosurfactant produced, the pH of the cellfree culture broth was adjusted to 1, 3, 5, 7, 9, 11 and 13 and the surface tension was
measured, respectively.
7.2.4
Thin Layer Chromatography (TLC)
Biosurfactant produced in various media were extracted as described in
Section 3.5.1 and were characterized by thin layer chromatography using silica-gel
60 F254 plates (20cm x 20cm, Merck). The development of solvent systems used was
differed based on the components tested. The TLC plates were spotted with
biosurfactant extracts and developed with the following: solvent 1, petroleum etherdiethyl ether-acetic acid (80:20:1) for neutral lipids, solvent 2, chloroform-methanolwater (65:25:4) for polar lipids, solvent 3, n-buthanol-acetic acid-water (4:1:1) for
amino acids and solvent 4, ethyl acetate-acetic acid-methanol-water (12:3:3:2) for
carbohydrate compounds.
After developing, the spots were visualized with standard reagents. The lipid
components were detected as yellow spots after placing the plates in a closed jar
saturated with iodine vapors. Using the ninhydrin solution followed by heating at
90ºC for 5 minutes, generated a red or purple color when the compound had an amine
function. Carbohydrate components were detected as red spots on the plates after
spraying with an α-naphthol solution followed by concentrated sulphuric acid, and
111
heated for 5 minutes at 100ºC. These components were identified by comparison
with published reports and against various amino acids and carbohydrates standards
(Sigma Chemicals Ltd.).
7.2.5
Fourier Transform Infrared (FTIR)
The biosurfactant extracted using chloroform-methanol (2:1) method (Section
3.5.1-3.5.2) was first dried with sodium sulphate, Na2SO4 to remove residual water
before being evaporated on a rotary evaporator. The FTIR spectra were recorded on
the Shimadzu FTIR-8300 Spectrophotometer, in the 4000-400 cm-1 spectral regions
at a resolution 2 cm-1. Each spectrum was scanned 10 times using a Mull technique.
7.2.6 Fatty Acid Analysis
The fatty acids composition of biosurfactant extracts were analyzed by GCMS (HP 5890(II) gas chromatograph combined with HP 5989A mass spectrometer)
of fatty acids methyl esters.
The GC was equipped with flame ionization detector and HP-wax columncrosslinked polyethylene glycol (30m x 0.25mm x 0.15µm). The carrier gas was
helium at 6ºC / min. The operating temperature of injector was 250ºC and that of the
detector was 270ºC. The column temperature was set at 50ºC for the first minute,
then increased to 200ºC at a rate of 4ºC min-1 and remained at the final temperature
for 20 minutes.
Fatty acids were released from the biosurfactant extracts by alkaline
hydrolysis followed by methylation. Approximately 10 mg biosurfactant extracts
(Section 3.5.1-3.5.2) were saponified with 1mL NaOH 60mM in methanol (40% v/v)
at 37ºC for 18 hours. Methylation of organic extracts was carried out by adding 2mL
methanol solution followed by acidification with HCl 1M before it was kept at 600C
for 30 minutes. After twofold extraction with MTBE-hexane (1:1), the fatty acids
methyl esters obtained were analyzed by GC-MS.
112
7.3
Results and Discussion
7.3.1
Emulsification Activities
There have been many studies on mesophilic microorganisms that produce
bioemulsifiers [Cooper and Goldenberg, 1987; Rosenberg, 1993]. Most of the
bioemulsifier had been identified as polymeric substances consisting of
polysaccharides that were able to stabilize emulsions and lower the surface tension
significantly [Neu and Poralla, 1990]. Usually, these were complex polymeric
compounds. Their emulsification properties are important to influence primarily the
structure of the water miscible and water immiscible components of biosurfactants.
The effect of pH on the emulsifying activity of the cell-free biosurfactants
extracts was determined after 24 hours of incubation at room temperature. Results
showed that the biosurfactants produced by both isolates AB-Cr1 and ETL-Cr1 have
higher emulsifying activity at pH 10 with the Ei of more than 60%. Figure 7.1 shows
the results of the emulsion test performed on shake flask samples after the removal of
biomass and pH adjustment.
70
Ei (%)
50
30
10
-10
0
5
pH
10
15
Fig. 7.1: Effect of pH on the activity of the emulsifier produced by AB-Cr1
and ETL-Cr1
isolates.
It was observed that there was strong pH dependence in the ability of the
biosurfactant to emulsify. This was also reported by Cooper and Goldenberg, 1987
for biosurfactant produced by Bacillus sp. The biosurfactant produced by both
isolates used in this study were very effective emulsifiers at pH 10. They emulsified
113
more than 60% of the kerosene added which corresponded to complete
emulsification of the oil phase. According to Willumsen and Karlson (1997), a good
bioemulsifier was designated as those having Ei of more than 50%. On the other
hand, the emulsion index obtained was found less than 10% at pH 13 (Table 7.1),
whereas no emulsion was obtained at the pH below 10 and above 13. Table 7.1
summarized the results obtained from this experiment.
Table 7.1: Emulsification activity and stabilization of bioemulsifiers by isolated
biosurfactants.
pH
Isolates
Emulsification Index, Ei (%)
Decay constant (Kd)
1
AB-Cr1
-
-
ETL-Cr1
-
-
AB-Cr1
-
-
ETL-Cr1
-
-
AB-Cr1
-
-
ETL-Cr1
-
-
10
AB-Cr1
ETL-Cr1
63.63
60.56
-0.06
-0.14
13
AB-Cr1
ETL-Cr1
8.67
7.69
-0.07
-0.05
4
7
The stability of the emulsions has been reported to be importance for both the
performance and the effectiveness of the emulsifier [Willumsen and Karlson, 1997].
The emulsion stabilizing capacity of biosurfactant could be characterized by studying
the decay pattern of the emulsion formed (Appendix C). A stable biosurfactant was
indicated by the decay constant (Kd) value of less than -2.00 obtained from the slope
value of the emulsion decay pattern [Kim et al., 2000]. In this study, stable and
compact emulsions of kerosene-crude biosurfactant fluid were observed after 24
hours and they were found stable up to 4 days. The stabilization properties (based on
Kd values) of biosurfactant produced by both isolates were found equally good at
both pH 10 and 13.
From these results, it could be possible to suggest that the biosurfactant
produced were emulsifier-type biosurfactant and would be useful in the applications
114
for biodegradation of hydrocarbon or other water-immiscible substrates as well as
enhanced oil recovery. This property was important to reduce the capillary forces that
entrapped oil within the pores of the rock as well as a mobility control agent to
improve the sweep efficiency of a water flood in petroleum industry [De Acevedo
and McInerney, 1996].
7.3.2
Critical Micelle Concentration (CMC)
Micellization was an important phenomenon in surfactant chemistry because
it affects many interfacial phenomena such as surface and interfacial tension
reduction that do not directly involve micelles. It is a characteristic property of a
biosurfactant. In aqueous solution, biosurfactant tended to aggregate and formed
colloidal sized clusters known as micelles. At very low concentrations, individual
molecules are present singly. As the biosurfactant concentration increase, a point
called the critical micelle concentration (CMC) is reached (Figure 7.2).
Fig. 7.2: Schematic diagram of the variation of surface tension, interfacial tension
and the CMC point with surfactant concentration [Mulligan et al., 2001].
CMC was defined as the lowest concentration of the surfactant required to
initiate micelle formation. It was proportional to the amount of surfactant present and
was generally used to measure the efficiency of a surfactant. There will be no further
decrease in surface tension after this point although more surfactant is present in the
medium. At the CMC, sudden changes in surface tension, electrical conductivity,
115
detergency, viscosity, density and osmotic pressure could be observed [Margaritis et
Surface tension, mN/m
al., 1979; Kim et al., 2000].
70
60
CMC
50
40
30
0
0.5
1
[Biosurfactant], g/L
1.5
Fig. 7.3: Surface tension of a solution against the concentration of the biosurfactant
produced by AB-Cr1
and ETL-Cr1
, grown in medium
supplemented with glucose as sole source of carbon.
The CMC of biosurfactant in the culture medium absence of crude oil was
found lower than that observed in the culture medium with the presence of crude oil.
As showed in Figure 7.3, the CMC values of crude biosurfactant extract from ABCr1 and ETL-Cr1 in the medium not supplemented with crude oil were found to be
approximately 0.1 and 0.2 g/L, respectively, and the surface tension (γ cmc) at that
point were 36.7mN/m and 40.2mN/m, respectively. These results indicated that both
isolates produced an efficient biosurfactant in the medium absence of crude oil.
Minor concentrations of biosurfactant are required for maximal surface activity
(43.5-48.5%).
Efficiency was measured by the surfactant concentration required to produce
a significant reduction in the surface tension of water. Whereas, effectiveness is
measured by the min value to which the surface tension can be reduced [Kim et al.,
2000]. Therefore, some important characteristic of potent biosurfactant were their
abilities to lower the surface tension in aqueous solutions and to possess a low CMC.
In this study, biosurfactant produced by both isolates AB-Cr1 and ETL-Cr1
grown in medium supplemented with glucose as the sole source of carbon might also
reduced the surface tension of the water from 71.2mN/m to 33.1mN/m and 35mN/m,
respectively at a biosurfactant concentration of 1.5 g/L. Thus, biosurfactant produced
by both AB-Cr1 and ETL-Cr1 isolates were also an effective biosurfactant as well as
116
an efficient biosurfactant in the medium absence of crude oil. From an environmental
point of view, low values of CMC and high capacity to reduce surface tension are
among the optimal characteristics that surfactants should have in order to promote
remediation of contaminated subsurface environments by increasing the hydrocarbon
mobility [Mata-Sandoval et al., 1999; Banat et al., 2000].
In contrast, the biosurfactant produced by both isolates in the medium with
crude oil added have a lower capacity to reduce surface tension. As shown on Figure
7.4, the CMC of the biosurfactant produced in the presence of crude oil were
determined to be approximately 1.0 and 1.2 g/L for AB-Cr1 and ETL-Cr1,
respectively. As the biosurfactant concentrations reached the CMC values, the
surface tension of water were significantly reduced (40%) from 71.2mN/m to
42.1mN/m and 42.3mN/m by AB-Cr1 and ETL-Cr1, respectively. These surface
tension values were found to be stable at increasing concentration of biosurfactant
Surface tension, mN/m
presence.
75
65
CMC
55
45
35
0
0.5
1
[Biosurfactant], g/L
1.5
Fig. 7.4: Surface tension of a solution against the concentration of the biosurfactant
produced by AB-Cr1
and ETL-Cr1
, grown in medium
supplemented with glucose and crude oil.
7.3.3
Stability Studies
Biosurfactant produced by both isolates, AB-Cr1 and ETL-Cr1 was
considerably pH and thermally stable as shown in Figure 7.5 and 7.6.
117
Surface tension, mN/m
44
42
40
38
36
34
32
0
2
4
6
8
10
12
14
pH
Fig. 7.5: The pH stability test of biosurfactant produced by AB-Cr1
and ETLCr1
grown in medium supplemented with glucose, based on the
change of surface tension values.
Surface tension, mN/m
40
39
38
37
36
35
34
33
0
10
20
30
40
50
Time of heating, min
60
70
Fig. 7.6: Thermal stability test of biosurfactant produced by AB-Cr1
and ETLCr1
grown in medium supplemented with glucose, based on the
change of surface tension values.
The surface activity of the biosurfactant produced by AB-Cr1 was remained
stable at pH ranging from 3.0 to 13.0 at room temperature with a minimum deviation
in surface tension. Variation in pH has no appreciable effect on surface activity
except at pH 1.0, which a negligible loss in activity (3mN/m) was observed. This
result was found in good agreement with those observed previously (Chapter 5 and
6), that AB-Cr1 produced a good biosurfactant in terms of their pH stability because
most known biosurfactants were found less stable over such an extreme pH range
[Kim et al., 2000]. In contrast, biosurfactant produced by ETL-Cr1 was only stable at
pH ranging from 5.0 to 9.0. Below and above these pH values, a slight decrease in
activity was observed (Figure 7.5).
118
When the cell-free culture broth of both isolates were heated at 100 0C for
different time intervals, they were found capable of retaining their surface activity
(33.4-35.1mN/m and 36-37.5mN/m for AB-Cr1 and ETL-Cr1, respectively) for 1
hour. After 1 hour of heating, the biosurfactant produced was lack of their surface
property. The loss of surface activity was observed by the increased value of surface
tension measured (Figure 7.6).
The high stability of the biosurfactant produced by AB-Cr1 at a wide range of
pH and temperature made it very suitable for the extreme conditions encountered in
the field application such as for in situ microbial enhanced oil recovery (MEOR). In
addition, the high pH stability properties of the biosurfactant produced could also be
of potential applications in acidophilic and alkalophilic environments.
7.3.4
Thin Layer Chromatography (TLC)
Preliminary analysis of the biosurfactant produced by AB-Cr1 and ETL-Cr1
isolates in the presence or absence of crude oil indicated the presence of different
lipid compounds, identified on the basis of their Rf values (Table 7.2). Ninhydrin and
α-naphthol-sulphuric acid positive fractions were also detected, indicating to the
presence of amino acid and carbohydrate substances in the biosurfactant extracts.
Table 7.2: TLC analysis of biosurfactant produced by AB-Cr1 and ETL-Cr1 isolates
based on the Rf values.
Rf
Components
/ Isolates
AB-Cr1
(- C/O)
ETL-Cr1
(- C/O)
AB-Cr1
(+ C/O)
ETL-Cr1
(+ C/O)
Neutral lipid
Polar lipid
0.65
0.68
0.67
0.92
Amino acid
0.49
0.67
0.85
-
-
0.65
0.83
0.69
0.72
0.83
-
0.37
0.60
0.78
0.71
Carbohydrate
0.22
119
The crude extract of lipid fractions in biosurfactant produced by both AB-Cr1
and ETL-Cr1 isolates also showed the presence of polar lipid compounds with the Rf
values of 0.65 and 0.68, respectively. The polar group presence in the lipid extracts
was then identified as amino or carbohydrate compounds based on their reaction with
the mobile phases (Section 7.2.4) used in the analysis.
The lipid fractions of biosurfactant produced by AB-Cr1 grown in medium
supplemented with glucose give three typical amino acid spots, revealed after the
ninhydrin staining with the Rf values as showed in Table 7.2. By comparing the data
obtained with the amino acid standards and reference from the previous study [Neu
and Poralla, 1990; Kluge, et al., 1988], it can be observed that AB-Cr1 produced
biosurfactant that contain Val, Leu and Glu amino acids. These amino acids were
found similar to that observed in surfactin, a lipopeptide surfactant produced by
Bacillus subtilis ATCC 2132 [Kluge et al., 1988]. No spots detected in the
carbohydrate analysis for biosurfactant produced by AB-Cr1 in the absence of crude
oil. This result had suggested that AB-Cr1 produced a lipopeptide type of
biosurfactant. A lipopeptide biosurfactant is well known as a potent surfactant with
high antibiotic activity that was useful for some therapeutic applications such as
inhibiting fibrin clot formation and hemolysis [Jenny et al., 1991; Banat et al., 2000].
However, the polar lipid in crude biosurfactant produced by ETL-Cr1 grown
in the crude oil-free medium showed with α-naphtol-sulphuric acid positive spots at
the Rf values of 0.37, 0.60 and 0.78, indicated that the biosurfactant contains a
mixture of carbohydrate substance. No detectable amount of amino acid was
obtained in the biosurfactant produced by ETL-Cr1 grown in both medium
supplemented or not with crude oil. This result suggested that ETL-Cr1 produces a
glycolipid type of biosurfactant. A glycolipid biosurfactant have been widely studied
in the application of environmental control. They were used intensively in oil storage
tank cleaning by reducing the viscosity of heavy oils, thereby facilitating recovery,
transportation and pipelining [Bertrand et al., 1994].
Table 7.2 also indicated that the biosurfactant produced by isolate AB-Cr1
grown in the presence of both glucose and crude oil as carbon sources were isolated
as mixtures of lipoprotein and glycolipid indicated by the positive spots in the
analysis of all compounds tested (Table 7.2). Neutral lipid biosurfactants were also
detected in the extracts from AB-Cr1 and ETL-Cr1 at Rf 0.67 and 0.69, respectively.
120
The mixture of biosurfactant could be the result of indiscriminate extraction of lipids
from cellular membranes by the hydrocarbon phase presence in the medium [Cooper,
1986].
7.3.5
Fourier Transform Infrared (FTIR)
Functional groups of the biosurfactant produced by both isolates were analyzed
by FTIR spectroscopy to further confirm that the biosurfactant produced were
lipopeptide and glycolipid type of biosurfactant. Figure 7.7 showed the infrared
spectrum of the biosurfactant, produced by AB-Cr1 isolate grown in medium
containing glucose as the sole source of carbon. Strong bands characteristic for
peptides at 3275 cm-1 (band A), at 1695 cm-1 (band D), and at 1537.5 cm-1 (band E)
resulting from the N-H stretching mode, the stretching mode of C=O bond and the
deformation mode (combined with the C-N stretching mode) of the N-H bond,
respectively were observed, indicating that this compound is a lipopeptide.
Fig. 7.7: Infrared spectrum of the surface-active fraction extracted from culture of
AB-Cr1 grown in medium supplemented with glucose as the sole source of
carbon.
121
Other bands observed in the FTIR spectrum of the biosurfactant produced by
AB-Cr1 were found quite similar to the spectrum of the biosurfactant produced by
ETL-Cr1 grown in medium containing glucose as sole carbon source (Figure 7.8).
The aliphatic nature was clearly showed by several C-H stretching bands of CH2 and
CH3 groups observed in the region 3000-2800 cm-1 (bands B and C). The
deformation vibrations (C-CH3 and C-CH2) at 1454-1362 cm-1 (bands F and G) also
confirm the presence of those alkyl groups.
Fig. 7.8: Infrared spectrum of the surface-active fraction extracted from culture of
ETL-Cr1 grown in medium supplemented with glucose as the sole source
of carbon.
Carbonyl (C=O) stretching band was also found at 1735 cm-1 (band D) in the
spectrum of biosurfactant produced by ETL-Cr1 (Figure 7.8), which is characteristic
for ester compounds. Both spectra showed the C-O deformation vibration bands
appear at 1084-1152 cm-1 (bands H and I) was also proved for the presence of ester
carbonyl group.
One of the characteristics of the biosurfactant produced by ETL-Cr1 isolate
was the presence of hydroxyl (-OH) group involved in hydrogen bonds in the region
of 3226 cm-1 (band A, Figure 7.8). However, biosurfactant produced by ETL-Cr1
lack of peptide characteristic bands as observed in the spectrum of biosurfactant
122
produced by AB-Cr1, indicating that the product contains aliphatic hydrocarbons as
well as glycolipid like moiety of biosurfactant.
Figure 7.9 and 7.10 showed the spectra of biosurfactant produced by AB-Cr1
and ETL-Cr1 isolates, respectively in the medium containing both glucose and crude
oil as carbon sources.
Fig. 7.9: Infrared spectrum of the surface-active fraction extracted from culture of
AB-Cr1 grown in medium supplemented with both glucose and crude oil
as carbon sources.
123
Fig. 7.10: Infrared spectrum of the surface-active fraction extracted from culture of
ETL-Cr1 grown in medium supplemented with both glucose and crude oil
as carbon sources.
Both biosurfactant produced by isolates AB-Cr1 and ETL-Cr1, grown in
medium containing both glucose and crude oil produced similar types of
biosurfactant as indicated with the analysis of IR spectra (Figure 7.9 and 7.10). The
biosurfactants produced have a characteristic of ester compound that contains
aliphatic hydrocarbon moiety. In the region 2850-2930 cm-1 (bands B and C) were
observed several C-H stretching bands of CH2 and CH3 groups. The deformation
vibrations at 1370-1470 cm-1 (bands F and G) confirm the presence of those aliphatic
chains (-CH2-, -CH3). The presence of C=O stretching band at 1695-1740 cm-1 (band
D) indicated to the characteristic of an ester compound. Similar to the biosurfactant
produced in medium containing only glucose as the sole source of carbon, the C-O
deformation vibrations were also observed in the biosurfactant produced by both
isolates, grown in medium containing both glucose and crude oil as source of carbon
(ν, 1028-1100 cm-1).
However, there was a clear difference between the characteristics of the
biosurfactant produced by both isolates grown in the medium containing crude oil. In
the region ~1630 cm-1 (band E, Figure 7.9) was observed a C=C stretching band
presence in the biosurfactant produced by AB-Cr1. The presence of the C=C bond
also confirmed by the deformation vibration observed at ~800 cm-1 (band I).
124
7.3.6 Fatty Acid Analysis
The presence of fatty acids in the lipophilic portion of the biosurfactant
extracts produced by both isolates was analyzed by combined GC-MS. The
chromatograms obtained from both AB-Cr1 and ETL-Cr1 isolates exhibited two
peaks, identified as fatty acids with different retention times (Figure 7.11 and 7.12).
Fig. 7.11: GC-MS chromatogram of fatty acid methyl ester from a culture medium of
AB-Cr1.
125
Fig. 7.12: GC-MS chromatogram of fatty acid methyl ester from a culture medium of
ETL-Cr1.
Table 7.3 summarized the relative positions of the peaks and their
composition in lipid obtained from GC-MS analysis for fatty acid methyl esters
presence in the biosurfactant produced.
Table 7.3: Relative positions of peaks from GC-MS for methyl esters of fatty acids.
Isolates
AB-Cr1
ETL-Cr1
Types of fatty acids
produced
Pentadecanoic acid
Octadecanoic acid
Pentadecanoic acid
Heptadecanoic acid
Relative positions of
the peaks
Composition (%)
in lipid
15.52
17.70
15.51
17.69
36.90
42.18
8.81
5.34
The results of GC-MS analysis showed in Table 7.3 indicated that both ABCr1 and ETL-Cr1 isolates produced 2 types of saturated fatty acids in the non-polar
chain of the biosurfactant. One of the fatty acids presences in crude biosurfactant
extracts was generally similar and differed only in the proportion of the fatty acid.
AB-Cr1 isolate produced biosurfactant that contain a significant amount of
pentadecanoic acid (36.90% of the total), whereas ETL-Cr1 was distinguished by a
126
lower concentration (8.81%) of this compound. Although the composition of
pentadecanoic acid in the lipid portion of the biosurfactant produced by ETL-Cr1
was much lower than that in AB-Cr1, it was found to be the predominant saturated
fatty acid compared to the other fatty acid obtained.
The chromatograms of both isolates exhibited another peak at nearly the same
retention time (~17.70 min) but different type of fatty acid as identified by MS. The
peaks were identified as saturated, long chain fatty acids namely octadecanoic acid
presence in biosurfactant from AB-Cr1 and heptadecanoic acid from ETL-Cr1.
Octadecanoic acid (42.18%) was presence as the predominant saturated fatty acid
compared to pentadecanoic acid (36.90%) produced by AB-Cr1. Figure 7.13 – 7.15
showed the structure of fatty acids detected in the biosurfactant of both isolates.
Fig. 7.13: Structure of pentadecanoic acid (C15H30O2).
Fig. 7.14: Structure of octadecanoic acid (C18H36O2).
Fig. 7.15: Structure of heptadecanoic acid (C17H34O2).
127
The result verified by MS spectra (Figure 7.16-7.18) showed the
characteristic base peak at m/z 74 typical for saturated fatty acid methyl esters due to
the ions formed up on an
cleavage with simultaneous migration of one hydrogen
atom from the lost fragment, known as the McLafferty rearrangement. Signals at m/z
29 and 43 were due to the bond cleavage that lead to the formation of ethyl- (-C2H5 +)
and propyl- (-C3H7+) group. The presence of the molecular ions (M+) at m/z 270 and
298 defined the chain length to be C17 and C19, respectively due to the presence of
methylated ester of the fatty acids.
Fig. 7.16: Mass spectrum of pentadecanoic acid from a culture of AB-Cr1.
Fig. 7.17: Mass spectrum of octadecanoic acid from a culture of AB-Cr1.
Fig. 7.18: Mass spectrum of heptadecanoic acid from a culture of ETL-Cr1.
CHAPTER 8
GENERAL DISCUSSION AND CONCLUSION
8.1
Conclusion
Ten bacterial strains previously isolated from petrochemical waste water were
selected for the screening of biosurfactant producer(s), via four different methods
commonly used and described elsewhere. Two mesophilic isolates coded AB-Cr1,
isolated from Titan Petrochemical (M) Sdn. Bhd. and ETL-Cr1, isolated from Exxon
Mobile Oil Refinery were chosen to be the most potential biosurfactant producers
among other isolates tested. Both isolates showed highest affinity towards
hexadecane (hydrocarbon), capable to destabilize the oil droplet within one hour,
showed -hemolysis on blood agar and lowered the surface tension of water, with the
reduction up to ~35mN/m compared to other isolates tested (16-27mN/m).
From the results of the biochemical tests and their morphological
characteristics, it was possible to suggest that the isolates AB-Cr1 and ETL-Cr1
might belong to the genus of Actinobacillus and Aeromonas, respectively. These
biosurfactant producers were selected to be studied in further for biosurfactant
production in shake flasks.
Both isolates AB-Cr1 and ETL-Cr1 were found grown optimally in Ramsay
medium supplemented with 3mM glucose, adjusted to an initial pH 7.0 and incubated
at 37ºC. The production of biosurfactant by both isolates was found to be growthassociated at all conditions tested. Linear relationship between biosurfactant
production and cell growth indicated by the plot of q p and µ (Figure 5.10) also
129
suggested to the growth-associated production of biosurfactant. Thus, the
biosurfactant was produced by both isolates as primary metabolite. The production in
medium with crude oil (7.18-8.26 g/L) was found similar to that observed in the
absence of crude oil (6.33-8.76 g/L). However, a longer fermentation period was
required by both isolates to achieve maximum production of biosurfactant when
grown in the medium supplemented with crude oil in comparison with those in crude
oil-free medium (Section 5.3.3).
The efficiency of both isolates to produce biosurfactant was indicated by the
comparison in values of productivity and their surface tension reducing ability. In the
medium containing glucose as the sole carbon source, both AB-Cr1 and ETL-Cr1
isolates were capable of performing highest rate of biosurfactant production (1.10
and 0.79 g/g/h, respectively) with high values of surface tension reduction (32 and
18.5mN/m, respectively) compared to those in the medium containing both glucose
and crude oil as carbon sources (0.08 and 0.07 g/g/h, respectively).
This study also showed that the bacterial mixed culture 3:1 (AB-Cr1:ETLCr1) system in the medium containing glucose as sole carbon source can efficiently
produce biosurfactant in higher concentrations (9.35 g/L) compared to those as single
culture (6.33-8.76 g/L) in shake flasks (Section 5.3.2-5.3.4). However, the efficiency
of this system to reduce surface (20.5mN/m reduction) and interfacial tension
(9.5mN/m reduction) of the medium was not as good as those obtained in single
culture of AB-Cr1 (32mN/m reduction) grown in the same condition (Section 5.3.2).
AB-Cr1 isolate was selected to be study of biosurfactant production in a
bioreactor. Similar to that observed in the shake flask experiment, the pattern of
biosurfactant production in bioreactor was also growth-associated. The highest
production of biosurfactant obtained was 12.45 g/L after 48 hours of fermentation, by
AB-Cr1 grown in medium adjusted to an initial pH of 7.0, supplemented with 25mM
NH4NO3 as nitrogen source and 3mM glucose as carbon source, incubated at 37ºC.
The pH controlled strategy was found not suitable for enhanced production of
biosurfactant.
At the optimum condition, the production of biosurfactant by AB-Cr1 in
bioreactor was shown as a biphasic pattern. The first phase of biosurfactant
production occurred during the exponential phase of cell growth followed by the
production at stationary growth phase. The maximum biosurfactant production of
130
approximately 7.03 g/L obtained during the first 25 hours of fermentation. The
second phase of biosurfactant production was observed during the following 25 hours
of fermentation period (t = 25-50 hours) with the maximum production (Pmax) of
12.45 g/L.
TLC characterization of biosurfactant produced by AB-Cr1 and ETL-Cr1
isolates grown in the medium containing glucose as the sole source of carbon
indicated the presence of lipoprotein and non-aromatic glycolipid types of
biosurfactant, respectively. However, biosurfactant produced by both isolates in
crude oil-containing medium are isolated as a mixture of lipoprotein and glycolipid
as a result of indiscriminate extraction of lipids from cellular membranes by the
hydrocarbon phase presence in the medium [Cooper, 1986]. Functional groups of the
biosurfactant as analyzed by FTIR also confirmed to the presence of those
compounds in the biosurfactant extracts.
The GC-MS chromatogram of fatty acid methyl ester derived from the
lipophilic portion of both cultures indicated to the presence of two types of saturated,
long chain fatty acids. The peaks were identified as pentadecanoic acid (C15:0) as well
as octadecanoic acid (C18:0) and heptadecanoic acid (C17:0) presence in the
biosurfactant produced by AB-Cr1 and ETL-Cr1, respectively (Section 7.3.6).
The CMC of the biosurfactant in the crude oil-free medium was found to be
lower than that observed in the crude oil-containing medium. The CMC of the
biosurfactant produced in the presence and absence of crude oil were approximately
(g/L) 1.0 and 0.1 for AB-Cr1, and 1.2 and 0.2 for ETL-Cr1, respectively. The
biosurfactant produced by both isolates in the crude oil-free medium were also an
effective and efficient biosurfactant, as they might reduce the surface tension of the
water from 71.2 mN/m to a minimum value of 33.1-35 mN/m.
The biosurfactant were also found capable of producing a relatively stable
emulsion with hydrocarbon at pH 10. More than 60% of the kerosene was emulsified
by these emulsifier-type biosurfactants. It was also found stable at various pHs (3.013.0 and 5.0-9.0, for AB-Cr1 and ETL-Cr1, respectively) and thermostable for 1 hour
at 100ºC, based on the value of surface tension. The high stability of the biosurfactant
produced by AB-Cr1 at a wide pH range and temperature makes it very suitable for
the extreme conditions encountered in the application field such as for in situ
131
microbial enhanced oil recovery (MEOR) as well as other applications such as
biodegradation and pipeline cleaning.
8.2
Suggestion
The presence study has generated much important information on the trend of
biosurfactant production by the new strains isolated and their physicochemical
properties of the biosurfactant produced. However, the available information is still
limited on their biosynthetic mechanisms and their structural characteristics. Thus,
further studies from different fields are required in order to promote the research and
development of biosurfactant.
The production of biosurfactant can also be studied in fed-batch or continuous
fermentation process. The fed-batch culture would maintain the low level of the
residual substrate concentration in the system. This could avoid the toxic effects of a
medium component, thus enhanced the production of biosurfactant [Stanbury and
Whitaker, 1984]. In the continuous culture, the exponential growth phase might be
prolonged by the addition of fresh medium into the fermentation system. The loss of
cells in the system could be balanced by the formation of new biomass. Thus, high
production of biosurfactant could be obtained during prolonged stationary growth
phase [Stanbury and Whitaker, 1984].
The structural elucidation of biosurfactant can be determined by high
resolution 1H and 13C Nuclear Magnetic Resonance (NMR) Spectroscopy, High
Performance Liquid Chromatography (HPLC) and Fast Atom Bombardment (FAB)
Mass Spectroscopy. By these techniques, the structure of biosurfactant can be
determined in a relatively short time, with very small amount of sample. Once the
structure of biosurfactant had been determined, the biosynthetic pathway of
biosurfactant production could be proposed by enzymatic or radioactively labeled
precursors.
The biosurfactant-producing isolates should be subjected to phylogenetic
analysis in order to further confirm the genus and species of the isolates by 16S
rRNA gene sequence analysis.
132
Research can also be carried out on the bioavailability of biosurfactant and
their effects on biodegradation of contaminant. The study can be done by concerning
the interaction of the biosurfactants and the crude oil contaminant, relationship of
biosurfactant structure and the treatment of petrochemical waste, scale-up of
biosurfactant production and finally, the cost reduction efforts for ex-situ production
of biosurfactant.
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147
APPENDIX A
GRAFT OD600 VERSUS CELL BIOMASS
1
0.9
0.8
OD600
0.7
y = 0.1004x
2
R = 0.9995
0.6
0.5
0.4
0.3
0.2
0.1
0
0
2
4
6
Cell Biomass, g/L
8
10
148
APPENDIX B
GLUCOSE STANDARD CURVE.
1.4
1.2
OD490
1
y = 1.2801x
R2 = 0.9902
0.8
0.6
0.4
0.2
0
0
0.2
0.4
0.6
[Glucose], g/L
0.8
1
149
APPENDIX C
BIOCHEMICAL CHARACTERIZATION METHODS
1.
Gram staining
Principle:
The method is based on the ability of microorganisms to retain the purple
color of crystal violet during the colorization with alcohol. Gram stain is essential in
identifying an unknown bacterium to determine whether it is gram positive or gram
negative. The gram stain arises because of differences in the cell-wall structure of
gram positive or gram negative cell.
Method:
§
The heat-fixed smear of a single colony was covered with crystal violet and
left for 20 seconds.
§
The strain was then washed off using of distilled water. The excess of water
was drained using a soft tissue.
§
The gram’s iodine solution was added to the smear and left for 1 minute.
§
The gram’s iodine was poured off by flooded the smear with 95% off using
distilled water.
§
The smear was covered with safranin for 20 second. Then washed gently for a
few second and blot dry with bibulous paper.
§
Finally, the slide was examined under x1000 immersion oil microscope.
150
Result:
Gram
Positive
Negative
2.
Indication
Pink red colour
Remain in purple colour
Oxidase Test
Principle:
To detect a cytochrome oxidase that catalysed the oxidation of reduced
cytochrome by molecular oxygen. Certain bacteria contain oxidase that will catalyse
the transport of electrons between the electron donors in the bacteria and a redox dye
tetramethyl-p-phenylenediamine dihydrochloride. The dye is oxidized to deep purple
color.
Reagent preparation:
A 1.0g tetramethyl-p-phenylenediamine dihydrochloride (1% w/v) was
dissolved in 100mL of distilled water. The reagent was prepared fresh before testing
of the unknown microorganisms.
Method:
§
A piece of filter paper was placed on a glass slide. A loopful of freshly
prepared oxidase reagent was saturated on it using a loop.
§
A portion of the colony of the microorganisms tested was rubbed gently onto
the impregnated filter paper.
§
Finally, the appearance of an intense dark purple color was observed for the
positive result.
Result:
Dark purple color – positive
No color formed – negative
151
3.
Catalase Test
Principle:
Catalase is an enzyme that decomposes hydrogen peroxide into water and
oxygen. Chemically, catalase is a hemoprotein similar structure to haemoglobin,
except for ferum atoms in the molecule in the oxidized (Fe3+) rather than the reduced
(Fe2+) state. The test demonstrate the presence of catalase, an enzyme that catalyse
the release of oxygen from the hydrogen peroxide in example it has ability to
decompose hydrogen into water and free hydrogen as shown by following equation:
2 H2O2
2 H2O + O2 liberated gas
Reagent preparation:
3mL of 100% hydrogen peroxide solution was added to 97mL of distilled
water. The reagent was kept in dark bottle at 40C and was avoided to expose to the
light.
Method:
§
2-3mL of the reagent solution was poured into a test tube.
§
The test organisms were immersed into the reagent solution using a glass rod.
Active bubbling – catalase produced (positive)
No bubbles release – no catalase produced (negative)
4.
Motility Test
Principle:
Motility test is essential in determining if organism is motile or non-motile.
Bacteria are motile by means of flagella.
Method:
§
A small amount of light microscopic oil was placed near each corner of the
cover glass with a toothpick.
152
§
A loopful of pure culture was placed in centre of cover glass.
§
Depression slide was then pressed against oil on cover glass and quickly
inverted.
§
Finally, the slide was examined under x1000 immersion oil microscope.
Result:
A true motility if it is motile bacteria. An organism is considered motile if
only a few cells are seen moving about.
5.
Oxidation-fermentation Test
Principle:
Glucose can be degraded either by oxidative or by fermentative process with
the formation of pyruvic acid as the key intermediate in both metabolic pathways. In
fermentation, pyruvic acid ultimately transfers its electrons to organic compound
with the formation of a large amount of mixed acids, whereas in oxidation pyruvic
acid further enters the Krebs cycle where it ultimately transfers its electron to oxygen
from water. Citric acid produced in the Krebs cycle is a weak acid compared with the
mixed acid produced by fermentation.
Enteric base carbohydrates media for example Hugh and Leison’s O-F basal
medium will detect acid production by fermentation organism. Oxidative bacteria
produces low amount of acid in enteric media. The sufficient products were produced
from peptone in the medium to neutralize the acid produced by oxidative
metabolism. This is not a problem when large amounts of acid were produced by
active fermenters with the introduction of specially formulated O-F basal media, the
oxidative bacteria can be determined.
153
Medium preparation:
Hugh and Leison’s Medium;
Composition
Peptone
Sodium chloride
di-Potassium hydrogen phosphate anhydrous
Agar
g/L
2.0
5.0
0.3
2.5
Method:
§
The composition of chemical was mixed in water heated to 100 0C to dissolve
it. The pH was adjusted to 7.1.
§
The medium was allowed to cool to 50-550C and then 3mL of the
bromothymol blue was added to the medium.
§
A 50mL amounts was dispensed in screw-cap containers and autoclaves at
1210C for 15 minutes.
§
A 5mL at 10% (w/v) sterile glucose was added aseptically.
§
Then, a 5mL of the amount of medium was dispensed in sterile screw-cap
tubes.
§
The tubes were inoculated by stabbing with a sterile wire to the bottom of the
bottle.
§
Approximately 2mL of sterile method soft paraffin oil was overlaid to all
tubes labeled covered to exclude all oxygen.
§
The tubes was incubated at 370C and examined after 24-48 hours (can be
prolonged in orders to check the acid production).
Result:
Open tube
Yellow
Yellow
Green or blue
Sealed tube
Green
Yellow
Green
Interpretation
Oxidative organism
Fermentative organism
No utilization of carbohydrates
154
6.
Triple Sugar Ion Test
Principle:
To determine the ability of an organism to attack a specific carbohydrate
incorporated in a basal growth medium, with or without the production of gas, along
with the possible hydrogen sulphide (H2S) production.
Medium preparation:
Triple sugar ion agar.
Composition
Beef extract
Yeast extract
Peptone
Lactose
Sucrose
Glucose
Ferrous sulphate, FeSO4
Sodium thiosulphate, Na2S2O3
Sodium chloride, NaCl
Agar
Phenol red
g/L
3.0
3.0
20.0
10.0
10.0
1.0
0.2
0.3
5.0
12.0
0.024
Method:
§
The chemical composition above was dissolved into 1000mL of distilled
water and the pH was adjusted to 7.4 by using H2SO4 (1M) and NaOH (1M).
§
The medium was then distributed into tube approximately 5.0mL per tube to
make long slant agar medium.
§
The medium was then autoclaved and cooled in slanted position with deep
butt after sterilization.
§
The slant was inoculated with the tested bacterium by making a stab butt and
fish-tail slant before incubated for 24 hours at 350C.
155
Result:
Slant
Red color
(alkaline)
Yellow color
(acid)
Red color
(alkaline)
7.
Butt
Yellow color
(acid)
Yellow color
(acid)
Red color
(alkaline)
Interpretation
Fermentation of glucose
Fermentation of both glucose and
lactose
Neither glucose nor lactose fermented
Nitrate/ Nitrite Reduction Test
Principle:
To determine the ability of an organism to reduce nitrate to nitrites or free
nitrogen gas. The reaction usually takes place under anaerobic conditions, in which
an organism derives its oxygen from nitrate.
Composition
Beef extract
Peptone
Potassium nitrate, KNO3
Agar
g/L
3.0
5.0
1.0
12.0
Reagents:
§
§
-Naphthylamine (0.5% v/v)
Sulfanilic acid (0.8% v/v)
Both reagents were mixed in equal parts immediately before testing.
Method:
§
The composition of chemical was mixed in 1000mL of distilled water and
heated to 100 0C to dissolve it.
§
The medium was distributed into tube approximately 5.0mL per tube.
§
The medium was then autoclaved and cooled in slanted position with deep
butt after sterilization.
§
The slant was inoculated with the tested bacterium by making a stab butt and
fish-tail slant before incubated for 24 hours at 350C.
156
§
Ten drops of nitrate reagents were add before attempting to make an
interpretation.
§
If no color change, a pinch of zink dust was add directly to the tube.
Result:
1.
Gas production.
2.
§
Gas is present – positive
§
No gas is present – negative
Phase 1.
§
Pink to deep red color – positive (NO3- reduced to NO2-)
§
No color change – negative (NO3- is not reduced to NO2-)
proceed to phase 2.
3.
Phase 2 (Zink reduction test).
§
No color change – positive (absence of NO2-)
NO2- is reduced to free nitrogen of ammonia
§
8.
Pink to deep red color – negative (NO3- is still present)
Indole Test
Principle:
To determine the ability of an organism to split indole from tryptophane
molecule. Tryptophane is an amino acid that can be oxidized by certain bacteria to
form three major indolic metabolites, indole, skatole (methyl indole) and indoleacetic
(IAA). Various intracellular enzymes involved are collectively called tryptophanase,
a general term used to donate the complete system of enzymes that mediate the
production of indole by hydrolytic activity against substrate tryptophane.
Reagent preparation:
Kovacs’ Indole Test
§
10.0g p-dimethylaminoensaldehyde was dissolved in 150.0mL of pure
isoamyl alcohol.
§
A concentrated HCl was added slowly to the aldehyde-alcohol mixture.
157
Medium preparation:
Composition
Tryptophane
Yeast extract
Sodium chloride
di-Sodium Phosphate (Na2HPO4)
Agar
g/L
2.0
3.0
5.0
1.0
12.0
Method:
§
The chemical composition above was dissolved into 1000mL of distilled
water and the pH was adjusted to 6.8 by using H2SO4 (1M) and NaOH (1M).
§
Then, the medium was distributed into tube approximately 4.0mL per tube to
make long slant agar medium.
§
The medium was then autoclaved and cooled in slanted position after
sterilization.
§
A light inoculum was inoculated onto the medium and incubated at 370C for 2
days.
§
After incubation period, 5 drops of reagent was added directly onto the slant
tube. The reagent was rolled gently over the slant before making an
interpretation.
Result:
Result
Positive
Negative
Variable
9.
Indications
A red ring at the surface of the medium in the alcoholic layer.
No color development at the alcohol yellow, still yellow color
of the reagent.
An orange color at the surface of the medium due to the
development of skatole, a methylated compound which may
be a precursor to indole formation.
Gelatine Liquefaction Test
Principle:
To determine the ability of an organism to produce proteolytic-like enzymes
(gealtinases), which are liquefying gelatin. Naturally occurring proteins are too large
158
to enter a bacterial cell. Therefore in order for a cell to utilize protein, they first must
be catabolized into smaller components. The exocellular proteolytic-like enzymes
gelatinases is a two steps processes and the final result yields a mixture of individual
amino acids.
Protein + H2O
Polypeptides
Polypeptides
Individual amino acid
Method:
§
All the composition of chemical was suspended into 800mL distilled water.
§
The pHs were adjusted to pH 6.8 and then it up to 1L by distilled water.
§
After autoclaving, the media was distributed to bijou bottles in approximately
5mL per bottle.
§
The bottle was kept in upright position at 4-100C.
§
A heavy inoculum culture was stab into the medium to a depth of ½ to 1 inch.
§
Then, the bottle was incubated at 370C for 14 days.
§
After 14 days, the bottle was put into 40C fridge for 1 hour in order whether
gelatine liquefaction had occurred.
Result:
Result
Positive (gelatinase)
Negative (no gelatinase)
10.
Indications
Medium maintains as liquid after keeping in
the fridge for 1 hour.
Medium becomes solid after keeping in the
fridge for 1 hour.
Urease Test
Principle:
To determine the urease enzyme activity by urease-producing bacteria. If the
strain is urease-producer, the enzyme will break down the urea (by hydrolysis) to
give ammonia and carbon dioxide. With the release of ammonia, the medium
becomes alkaline and changed the indicator to red-pink color.
159
Medium preparation:
Composition
Potassium phosphate (KH2PO4)
Sodium chloride
Peptone
Glucose (0.1%)
Urea (Highest purity, 20%)
Phenol red
Agar
g/L
2.0
5.0
1.0
1.0
20.0
0.01
15.0
Method:
§
All the ingredients above except urea dissolved in 900mL of distilled water
and autoclaved at 1210C for sterilized.
§
The urea was rehydrated in 100mL of distilled water and then filters
sterilized.
§
When the medium added aseptically to the medium and distributed
approximately 5mL per tube.
§
The pure colony of bacterium was inoculated into the tube.
Result:
Result
Positive (urease produced)
Negative (no urease produced)
11.
Indications
Red pink color
Yellowish color remained
Citrate Test
Principle:
Simmon’s citrate agar is recommended for the differentiation of the family
Entrobacteriaceae based on weather or not citrate is utilized as the sole carbon
source. The only source of nitrogen is the sodium ammonium phosphate, while the
sole carbon source is sodium citrate. The organism utilizes citrate and produces an
alkaline reaction as indicated by the bromothymol blue, which changes the color
from green to blue.
160
Medium preparation:
Composition
Ammonium dehydrogen phosphate
Sodium ammonium phosphate
Sodium citrate
Magnesium sulphate
Sodium chloride
Agar
g/L
1.0
1.0
2.0
0.2
5.0
20
Method:
§
All the composition of chemical above was dissolved in 1L of distilled water.
§
The pH of the medium was adjusted to 6.8 using H2SO4 (1M) and NaOH
(1M).
§
The medium was then distributed into screw-cap tube and autoclaved at
1210C for 15 minutes.
§
After autoclaving, the medium was allowed to solidify in a slanted position.
§
The slant was inoculated with the tested bacterium and incubated for 4 days at
37 0C.
Result:
Result
Indications
Positive (citrate utilized)
Turbidity and blue color
Negative (citrate not utilized) No growth with no change in color (green)
161
APPENDIX D
PRODUCTION OF BIOSURFACTANT AND SURFACE TENSION
10
70
9
65
Biosurfactant, g/L
8
60
7
6
55
5
50
4
45
3
40
2
35
1
0
30
0
1
2
3
4
5
Time, h
Relationship of biosurfactant production
6
7
8
9
and surface tension reduction
by AB-Cr1 isolate grown in Ramsay medium supplemented with 3mM glucose,
adjusted to initial pH 7 and incubated at 37ºC.
Surface Tension, mN/m
REDUCTION IN THE MEDIUM GROWN WITH AB-Cr1 ISOLATE
162
APPENDIX E
0.4
6
0.3
5
4
0.2
3
2
[Glucose], g/L
Biomass & Biosurfactant, g/L
7
0.1
1
0
0
0
50
Time, h
100
150
(A)
0.5
7
0.4
6
5
0.3
4
0.2
3
2
0.1
1
0
0
0
50
Time, h
100
150
(B)
Relationship of growth
production
, glucose consumption
and biosurfactant
by bacterial mix culture system 1:1, grown in Ramsay medium
supplemented with glucose (A) and glucose + crude oil (B).
[Glucose], g/L
Biomass & Biosurfactant, g/L
8
163
APPENDIX F
DETERMINATION OF DECAY CONSTANT
Time, hrs
-0.6
10
20
30
40
log [Abs]
-0.8
-1
-1.2
-1.4
Determination of decay constants, Kd as an indication to emulsion stability formed
with crude biosurfactant of isolates AB-Cr1
,
and ETL-Cr1
,
at pH
10 (closed symbol) and pH 13 (opened symbol), respectively. The value of Kd was
obtained from the slope of each plot.
164
APPENDIX G
MASS SPECRUMS OF FATTY ACID METHYL ESTERS FROM THE
CULTURE OF AB-Cr1 AND ETL-Cr1 ISOLATES.