THE PERFORMANCE OF PHENOL BIODEGRADATION BY
Candida tropicalis RETL-Cr1 USING BATCH AND FED-BATCH
FERMENTATION TECHNIQUES
PIAKONG BIN MOHD.TUAH
UNIVERSITI TEKNOLOGI MALAYSIA
THE PERFORMANCE OF PHENOL BIODEGRADATION BY Candida tropicalis
RETL-Cr1 USING BATCH AND FED-BATCH FERMENTATION TECHNIQUES
PIAKONG BIN MOHD. TUAH
A thesis is submitted in fulfilment of the
requirements for the award of the degree of
Doctor of Philosophy
Faculty of Science
Universiti Teknologi Malaysia
JANUARY 2006
iii
Dedicated especially to my wife,
Nur Shiqah @Chuah Kim Hong Abdullah and
my children,
Nur Azidah, Nur Sulina and Nurul Atiqah
iv
ACKNOWLEDGEMENT
I wish to extend my deepest appreciation and thank you to both my supervisors;
Assoc. Professor Dr. Noor Aini Abdul Rashid, and Dr. Madihah Md Salleh for their
advice, invaluable comments, guidance and high level inspiration. My appreciation also
goes to Dr. Adibah Yahya, Assoc. Prof. Dr. Zaharah Ibrahim, Dr. Fahrul Zaman Huyop,
Dept. of Biology and Dr. Rosli Md. Illias, Dept. of Bioprocess, Universiti Teknologi
Malaysia for their continuous support and encouragement.
I would like to thank the Dept. of Biology for giving me the opportunity to use the
facilities and lab space. I wish to acknowledge the assistance given by Exxon Mobil Oil
Refinery, Port Dickson, Negeri Sembilan and Titan (Malaysia) Petrochemical Industries,
Pasir Gudang, Johor.
I am fortunate to have the opportunity to work with so many researchers in the
Molecular Biology and Microbiology Lab at Dept of Biology, UTM. I appreciate their
friendship and collective encouragement given to me at the most crucial moments. I am
thankful to Haryati Jamaluddin, Roslindawati Haron, S. Hasila Hamzah, Mohd. Firdaus,
Aishah Husin, Sharifah Norhafizah Syed Muhd. Rafeii, Hasniza Ramli, Norhasniza
Ibrahim, Maihafizah Mohd. Zahari, Rusniza Mohd. Zawawi, Chan Giek Far, Sia Kia
Chuan and Fathul Karim Sharani for their support and for sharing their ideas. My
gratitude also goes to Chong Chun Shiong for assisting with the printing and binding of
the thesis. I also wish to thank the Laboratory Assistants: Puan Fatimah Harun, Puan
Radiah Hassan and En. Mohd. Ruzaini bin Ramli for providing their assistance. I am also
grateful to Dr. Henry Parry and Dr. Alan Scragg, Univeristy West of England, Bristol,
U.K. for their assistance in supplying the relevant literatures.
My sincerest thank you to Universiti Malaysia Sabah for granting me my study
leave and financial support throughout my Ph.D.
My special thank you to my wife, Nur Shiqah @ Chuah Kim Hong Abdullah and
my children; Nur Azidah, Nur Sulina and Nurul Atiqah for their love, understanding,
perseverance and constant prayers. I love you all.
v
ABSTRACT
Phenol is a toxic compound found in many industrial-waste effluents. A locally
isolated yeast strain RETL-Cr1 from the effluent of the Exxon Mobil Oil Refinery
wastewater treatment plant was investigated for phenol degradation using batch and fedbatch fermentation under aerobic condition. Based on a BLASTN search of GenBank, the
complete sequences of ITS1-5.8S rDNA-ITS2 regions and portions of I8S and 28S for
the purified DNA products of RETL-Cr1 shared 98% similarity with C. tropicalis. This
yeast strain RETL-Cr1 was redesignated C. tropicalis RETL-Cr1 and was deposited at
the GenBank under the accession number AY725426. The optimum condition for phenol
degradation was at 30oC, pH 6.5 in RM in the absence of glucose. The highest phenol
biodegradation efficiency in shake-flask cultures with IPC of 3mM was 100% achieving
a degradation rate of 0.0257 g L-1 h-1 at µ 0.3718 h -1 after 14 h cultivation. Degradation
of phenol was faster by 1.5-fold in bioreactor than in shake-flask whereby degradation
rate was improved to 0.0395 g L-1 h-1 at µ 0.5391 h-1 after 10 hours of incubation. When
tested at various IPC (0.0028 – 0.94 g L-1), inhibition was evident at IPC levels above 5
mM (0.470 g L-1). The fed-batch system in a bioreactor offered an 85 times fold
degradation rate (2.3 g L-1 h-1) over shake-flask culture (0.0257 g L-1 h-1) and 61-fold over
2L bioreactor (0.0395 g L-1 h-1) batch system. It was observed that kinetically phenol
degradation by RETL-Cr1 was significantly high in fed-batch culture as indicated by high
degradation rate (2.3 g L-1 h-1) and substrate yield (Yx/s = 0.71-4.48 g g-1). However, a
lower product yield (Ypc/s = 1.6x10-4 – 2.1x10-3 g g-1; Ypc/x = 3.5x10-5 – 1.4 x10-3 g g-1;
YccMA/s = 1.0x10-4 – 2.0x10-4 g g -1; YccMA/x = 4.4x10-5 – 1.8x10-4 g g-1) and productivity
(catechol = 1.2x10-5 – 5.3x10-5 g L-1 h-1; ccMA = 1.4x10-5 – 2.6 x10-5 g L-1 h-1) were
achieved. When catechol and ccMA were analysed to determine whether an ortho or
meta pathway was taken, it was found that these two metabolites were present in low
amounts. This probably indicates further degradation of the metabolites. Hence, RETLCr1 strain metabolizes phenol via ortho-cleavage pathway. The optimum condition for
both phenol hydroxylase and catechol 1,2-dioxygenase were at 30oC, pH 6.5. The most
distinctive feature of this yeast strain is that it has a very high tolerance limit towards
phenol reaching up to 60 mM. Based on the observations, RETL-Cr1 has a good potential
to be used for treatment of phenol in industrial effluent.
vi
ABSTRAK
Fenol adalah sebatian toksik terdapat dalam pelbagai efluen sisa buangan industri.
Yis tempatan strain RETL-Cr1 dipencilkan daripada efluen loji pengolahan air sisa kilang
penapis minyak Exxon Mobil telah dikaji untuk pembiodegradasian fenol menggunakan
fermentasi kultur kelompok dan kelompok suapan dalam keadaan aerobik. Berdasarkan
pencarian pada GenBank, jujukan sepenuhnya kawasan ITS1-5.8S rDNA- ITS2 dan
bahagian-bahagian 18S dan 28S produk DNA RETL-Cr1 menyumbang 98% kesamaan
dengan C. tropicalis. Strain yis RETL-Cr1 ini telah dinamakan semula sebagai C.
tropicalis RETL-Cr1 dan disimpan dalam GenBank di bawah nombor penambahan
AY725426. Keadaan optimum bagi pembiodegradasian fenol adalah pada suhu 30oC, pH
6.5 dalam RM tanpa glukosa. Pembiodegradasian fenol dalam kultur kelompok kelalang
goncangan pada kepekatan fenol permulaan 3 mM adalah 100% mencapai kadar
pendegradasian 0.0257 g L-1 j-1, = µ 0.3718 j -1 selepas 14 jam pengeraman.
Pembiodegradasian fenol didapati 1.5 kali lebih cepat dalam kultur kelompok bioreaktor
berbanding dengan kelalang goncangan dengan pencapaian 0.0395 g L-1 j-1 pada µ 0.5391
j-1 selepas 10 jam pengeraman. Apabila diuji pada pelbagai IPC (0.028–0.94 g L-1), kesan
perencatan adalah jelas apabila kepekatan fenol melebihi tahap 5 mM (0.470 g L-1).
Sistem suapan sesekelompok mencapai 85 kali lebih baik dengan kadar pemdegradasian
2.3 g L-1 j -1 dari sistem kelompok kelalang goncangan (0.0257 g L-1 j-1) dan 61 kali dari
2L bioreaktor. Didapati dari segi kinetik, pembiodegradasian fenol dalam sistem suapan
kelompok adalah bersignifikan tinggi seperti ditunjukkan oleh kadar degradasi (2.3 g L-1
h-1) dan hasil substrat (Yx/s = 0.71-4.48 g g-1) yang tinggi. Walau bagaimanapun hasil
produk (Ypc/s = 1.6x10-4 – 2.1x10-3 g g-1; Ypc/x = 3.5x10-5 – 1.4x10-3 g g-1; YccMA/s =
1.0x10-4 – 2.0x10-4 g g -1; YccMA/x = 4.4x10-5 – 1.8x10-4 g g-1) dan produktiviti (katekol =
1.2x10-5 – 5.3x10-5 g L-1 h-1; ccMA =1.4x10-5 –2.6x10-5 g L-1 h-1) adalah rendah. Apabila
katekol dan ccMA dianalisis untuk menentukan samada laluan ortho atau meta, didapati
amaun kedua-dua metabolit ini adalah rendah. Ini menunjukkan berlakunya proses
pemdegradasian terhadap kedua-dua metabolit ini. Oleh itu, strain yis RETL-Cr1 ini
mendegrad fenol melalui laluan belahan ortho. Keadaan optimum bagi enzim fenol
hidroksilase dan katekol 1,2-dioksigenase adalah pada 30oC, pH 6.5. Ciri tersendiri yis ini
adalah ketolerannya yang tinggi terhadap fenol sehingga mencapai 60 mM. Berdasarkan
kajian ini, RETL-Cr1 berpotensi digunakan untuk rawatan fenol dalam efluen industri.
vii
TABLE OF CONTENTS
CHAPTER
1
2
TITLE
PAGE
TITLE
i
DECLARATION
ii
DEDICATION
iii
ACKNOWLEDGEMENT
iv
ABSTRACT
v
ABSTRAK
vi
CONTENTS
vii
LIST OF TABLES
xiii
LIST OF FIGURES
xvi
LIST OF SYMBOLS
xxi
LIST OF ABBREVIATIONS
xxiii
LIST OF APPENDICES
xxiv
INTRODUCTION
1.1
Introduction
1
1.2
Objectives of study
3
LITERATURE REVIEW
2.1
Phenol
2.1.1
5
Chemical identity, physical and
chemical properties of phenol
2.2
6
Sources of phenol
7
2.2.1
Natural sources
7
2.2.2
Man-made sources
7
viii
2.2.3
2.3
2.4
Endogenous sources
8
Releases of phenol to the environment
9
2.3.1
Air
9
2.3.2
Water
10
2.3.3
Soil
12
Fate of phenol in the environment
12
2.4.1
Air
12
2.4.2
Soil and sediment
13
2.4.3
Water
15
2.5
Hazards of phenol
16
2.6
Microbial degradation
17
2.6.1
Phenol-degrading microorganisms
18
2.6.2
Phenol-degrading Candida tropicalis
26
2.6.3 Aerobic biodegradation of phenol
27
2.6.3.1 Phenol inhibitory levels for phenol
degradation by microorganisms.
2.6.3.2 Phenol degradation lag period (TL)
28
30
2.6.3.3 Intermediates of phenol biodegradation
and metabolic pathway
2.6.4
2.7
3
40
Phenol biodegradation methods
45
2.7.1
Batch fermentation
45
2.7.1.1 Definition
45
2.7.1.2 Advantages and disadvantages
46
Fed-batch fermentation
46
2.7.2.1 Definition
46
2.7.2.2 Advantages and disadvantages
47
2.7.2
2.8
Anaerobic biodegradation of phenol
33
Summary of Literature Review
48
GENERAL MATERIALS AND METHODS
3.1
3.2
Media Preparation
51
3.1.1 Ramsay medium agar
51
Sample Collection
52
ix
3.3
3.4
Bacterial culture preservation
56
3.3.1
Short-term preservation
56
3.3.2
Long-term preservation
56
Phylogenetic analysis of phenol-degrading RETL-Cr1
3.4.1 DNA Extraction
56
3.4.2
57
Electrophoresis
3.4.3 Sequencing and analysis
57
3.5
Sample analysis
58
3.5.1
Determination of biomass concentration
58
3.5.2
Determination of specific growth rate
58
3.5.3
Determination of average phenol degradation
rate
59
3.5.4
Determination of glucose
59
3.5.5
Determination of phenol, catechol –
and cis,cis-muconic acid
4
56
60
ISOLATION, SCREENING AND CHARACTERIZATION
OF POTENTIAL PHENOL-DEGRADERS FROM
PETROCHEMICAL WASTES
4.1
Introduction
61
4.2
Materials and Methods
63
4.2.1
Media preparation
63
4.2.2
Sample collection
63
4.2.3
Isolation of microorganisms
63
4.2.4
Screening for phenol-degrading microorganisms
64
4.2.4.1 Test for growth on RM agar containing
1 mM phenol
64
4.2.4.2 Test of phenol tolerance for selected isolates
65
4.2.4.3 Test for survivality
65
4.2.5 Phenol degradation by selected isolates
66
4.2.6
66
Morphological characterization
4.2.6.1 Colony morphology
66
4.2.6.2 Cellular morphology
66
4.2.7 Biochemical tests
67
4.2.8 Identification of selected isolates
67
x
4.2.8.1 Phylogenetic analysis of phenol-degrading
RETL-Cr1
4.2.9
4.3
Sample Analysis
70
4.2.9.1 Determination of Biomass Concentration
70
4.2.9.2 Determination of average phenol degradation rate
70
4.2.9.3 Determination of Glucose Concentration
70
4.2.9.4 Determination of Phenol Concentration
70
Results and Discussion
70
4.3.1
Isolation and screening for phenoldegrading microorganisms
4.3.2
5
70
Morphological and physiological
characterization of selected strains
77
4.3.3
Biodegradation of phenol by selected strains
81
4.3.4
Characterization and identification of the best
phenol-degrading RETL-Cr1
4.4
67
Conclusions
86
89
BIODEGRADATION OF PHENOL IN BATCH
CULTURES OF YEAST Candida tropicalis RETL-Cr1
5.1
Introduction
91
5.2
Materials and Methods
93
5.2.1
Culture media
93
5.2.2
Batch fermentation: Shake-flask culture
94
5.2.2.1 The effect of temperature on phenol
degradation
94
5.2.2.2 The effect of pH on phenol1
phenol degradation
94
5.2.2.3 Effect of glucose on phenol degradation
94
5.2.3
Batch fermentation: Bioreactor culture
95
5.2.4
Experimental Design
95
5.2.5
Sample Analysis
96
5.2.5.1 Determination of biomass concentration
96
5.2.5.2 Determination of average phenol degradation rate
97
xi
5.2.5.3 Determination of phenol, catechol and
cis,cis-muconic acid
5.3
Results and Discussion
97
97
5.3.1 Optimization of phenol degradation inshake-flask culture
97
5.3.1.1 The effect of temperature on phenol
degradation in shake flask culture
97
5.3.1.2 The effect of glucose on phenol
degradation
104
5.3.1.3 The effect of pH on phenol
phenol degradation
111
5.3.1.4 The effect of initial phenol –
concentration (IPC)
5.3.2
116
Comparison of phenol degradation in shakeflask and bioreactor
126
5.3.3 Time course of phenol degradation by C.
tropicalis RETL-Cr1 under optimum condition
5.4
6
Conclusions
128
130
IMPROVEMENT OF PHENOL BIODEGRADATION IN
FED-BATCH CULTURES OF Candida tropicalis RETL-Cr1
6.1
Introduction
132
6.2
Materials and Methods
133
6.2.1
Fed-batch fermentation
133
6.2.1.1 Batch and Fed-Batch Experimental Design
133
Sample Analysis
136
6.2.2.1 Determination of biomass concentration
136
6.2.2
6.2.2.2 Determination of average phenol degradation
rate
136
6.2.2.3 Determination of phenol, catechol and
cis,cis-muconic acid
6.2.3
6.3
Microscopy observation
Results and Discussion
136
136
136
xii
6.4
7
6.3.1 Batch fermentation
137
6.3.2
138
Fed-batch fermentation
Conclusions
146
PHENOL-METABOLIC PATHWAY OF Candida tropicalis
RETL-Cr1
7.1
Introduction
148
7.2
Materials and Methods
149
7.2.1
Meta-cleavage dioxygenase assays
149
7.2.2
Determination of cis,cis-muconic acid
149
7.2.3
Experimental Design
149
7.3
Results and Discussion
7.3.1
Determination of intermediates
of C. tropicalis RETL-Cr1
7.3.2
8
150
Phenol metabolic pathway of C. tropicalis –
RETL-Cr1
7.4
150
Conclusion
153
156
CONCLUSION AND FUTURE RESEARCH
8.1
Conclusions
157
8.2
Future research
161
REFERENCES
163
APPENDICES
214
xiii
LIST OF TABLES
TABLE
2.1
TITLE
Sources of phenols and other related aromatic
compounds in wastewater
2.2
PAGE
8
Typical levels of phenol concentration in wastewater
of some selected industries
11
2.3
Phenol-degrading microorganisms
20
2.4
Source of origin of phenol-degrading Candida
tropicalis
2.5
27
Phenol inhibitory levels for phenol degradation
by microorganism
29
2.6
Observed phenol degradation lag period (TL)
31
2.7
Intermediates and products produced of phenol
degradation by microorganism
37
2.8
Phenol metabolism pathway of microorganism
38
3.1
Composition of Ramsay Medium (RM)
52
3.2
Oil and petrochemical waste samples collected
53
4.1
Aerobic growth comparison of selected isolates
on RM agar containing 1 mM phenol at 37oC.
73
xiv
4.2
Colony morphology of selected isolates on RM agar
at 37oC after 24 hours incubation isolated from two
sampling locations.
4.3
Biochemical tests, cellular morphology, and Gram
stain reaction of selected strains.
4.4
79
80
Growth kinetics and performance of phenol
degradation at 3 mM IPC by selected isolates
at 37oC, pH 6.5.
5.1
85
Effect of temperature on phenol degradation by
C. tropicalis RETL-Cr1 at different temperature,
pH 6.5. (shake-flask) after 18h incubation.
5.2
Effect of glucose on phenol degradation by
C. tropicalis RETL-Cr1 at 30oC, pH 6.5.
5.3
100
106
Effect of pH on phenol degradation by
C. tropicalis RETL-Cr1 at 30oC after 18h incubation
(RM broth with 3 mM IPC).
5.4
114
The effect of initial phenol concentration (IPC) on
phenol degradation by C. tropicalis RETL-Cr1
at 30oC, pH 6.5 in shake-flask.
5.5
118
Comparison of phenol degradation performance
in shake-flask and bioreactor cultures with an IPC of
3 mM of C. tropicalis RETL-Cr1 at 30oC, pH 6.5.
6.1
127
Kinetic parameters/kinetics of fed-batch
fermentation of phenol degradation by C. tropicalis –
RETL-Cr1.
139
xv
6.2
Kinetic parameters/performance of phenol degradation
in batch and fed-batch fermentation by C. tropicalis
RETL-Cr1.
145
xvi
LIST OF FIGURES
FIGURE
TITLE
2.1
Chemical structure of phenol
2.2
Microbial metabolism of some aromatic compounds
via catechol
2.3
36
42
Phenol degradation pathway, phenol transformation to
benzoate and acetate in the presence of BES.
3.1
34
Postulated pathway of anaerobic phenol metabolism
in the denitrifying bacterium T. aromatica.
2.5
6
The main pathways of phenol degradation under
under aerobic condition.
2.4
PAGE
44
Wastewater treatment system and sampling
points, Titan Petrochemical Sdn Bhd. (TPSB)
Pasir Gudang, Johor
3.2
Waste treatment system and sampling points at
Exxon Mobil Oil Refinery, Port Dickson, N.Sembilan
4.1
55
Schematic representation of the fungal ribosomal genes
containing the primer target areas
4.2
54
68
Experimental design of isolation, screening and
characterization of phenol-degrading microorganisms
from petrochemical wastes.
69
xvii
4.3
Number of strains isolated from petrochemical samples
via plating after enrichment in RM incubated at 37oC.
4.4
71
Growth comparison of selected isolates grown
aerobically in RM broth containing varying initial
phenol concentration as a sole carbon source at 37oC
after 24 h.
4.5
74
Test for phenol tolerance limit of isolate RETL-Cr1
in RM containing 1 mM glucose incubated
at 30oC, pH 6.5 after 96h.
4.6
75
Growth comparison of selected isolates grown
aerobically on RM broth containing 3 mM
phenol at 37oC, pH 6.5.
4.7
Phenol removal efficiency by selected isolates in
RM incubated at 37oC, pH 6.5.
4.8
77
82
Degradation of phenol against time and glucose
utilization by growth pattern of RETL-Cr1 in RM
containing 3 mM phenol at 37oC, pH 6.5.
4.9
Colony morphology of RETL-Cr1 on RM
agar under stereo microscope (x12).
4.10
87
The amplified DNA from C. tropicalis RETL-Cr1
ribosomal gene generated using TS1 and TS4 primers.
4.12
86
Gram morphology of RETL-Cr1 magnified x1000
under light microscopy.
4.11
83
Complete sequence of the 5.8S rDNA (Italics) flanked
by adjacent ITS1 and ITS2 regions of C. tropicalis –
88
xviii
RETL-Cr1.
5.1
Experimental design of phenol degradation by
C. tropicalis RETL-Cr1 in batch culture
5.2
88
96
The effect of temperature on the average phenol
degradation rate of C. tropicalis in the absence of glucose
in RM medium containing 3 mM phenol at pH 6.5 in
shake flask culture.
5.3
98
Hypothetical illustration on PH and C1,2D optimum
activity during phenol degradation by C. tropicalis –
RETL-Cr1 at optimum temperature.
103
5.4
Typical electron and energy flows in a bacterial cell.
104
5.5
Hypothetical Illustration on how glucose may affect the
primary flows of electrons and energy during phenol
degradation by C. tropicalis RETL-Cr1.
5.6
109
Degradation of phenol and utilization of glucose by
C. tropicalis-RETL-Cr1 in RM containing 3 mM phenol
at 30oC, pH 6.5.
5.7
110
The effect of pH on phenol degradation rate of
C. tropicalis RETL- Cr1 in RM containing 3 mM
initial phenol concentration at 30oC.
5.8
112
Hypothetical illustration on how low and high pH may
affect PH and C1,2D activity during phenol degradation
by C. tropicalis RETL-Cr1.
5.9
116
Hypothetical illustration on how high phenol
concentration may affect PH and C1,2D activity during
phenol degradation by C. tropicalis RETL-Cr1.
120
xix
5.10
Hypothetical illustration on how high phenol concentration
may affect the primary flows of electron and energy
during phenol degradation by C. tropicalis RETL-Cr1.
5.11
121
Concentration of intermediates; catechol
and cis,cis-muconic acid and phenol removal
efficiency at various IPC by C. tropicalis RETL-Cr1
5.12
124
Degradation of phenol and production of intermediates;
catechol and cis,cis-muconic acid by by C. tropicalisRETL-Cr1 against time at IPC of 5 mM in RM at 30oC,
pH 6.5 in shake-flask.
5.13
125
Degradation of phenol by C. tropicalis against time in
RM with IPC of 3 mM in the absence of glucose at
at 30oC, pH 6.5.
129
6.1
Fermenter set-up for fed-batch culture.
135
6.2
Time course of phenol degradation in batch culture by
C. tropicalis RETL-Cr1 in RM at 30oC, initial pH 6.5.
6.3
137
Time course of phenol degradation in fed-batch
fermentation by C. tropicalis RETL-Cr1 in RM at 30oC,
initial pH 6.5.
6.4
141
Hypothetical illustration how low pH (3.9) may affect PH,
C1,2D and ccMA lactonizing enzyme (ccMALe) activity
at the end of phenol degradation process by C. tropicalis –
RETL-Cr1 in fed-batch fermentation.
6.5
143
Hypothetical illustration how ccMA may affect the primary
flows of electrons and energy during phenol degradation by
C. tropicalis RETL-Cr1.
144
xx
7.1
Experimental design to postulate possible phenol
metabolic pathway of C. tropicalis RETL-Cr1.
7.2
150
Typical HPLC chromatogram recorded in an aerated
suspension: cis,cis-muconic acid, catechol and phenol
during phenoldegradation by C. tropicalis RETL-Cr1
at initial phenol concentration of 3 mM after
7h incubation.
7.3
151
Time course of phenol degradation in batch system
(shake-flask) using C. tropicalis RETL-Cr1 at IPC
of 3 mM, pH 6.5, and detection of intermediates.
7.4
General principle of aerobic aromatic catabolism
in bacteria.
7.5
152
153
Postulated ortho-pathway for degradation of
phenol by C. tropicalis RETL-Cr1
155
xxi
LIST OF SYMBOLS
Į
-
alpha
ß
-
beta
Ȗ
-
gamma
abs
-
absorbance
o
-
degrees Celsius
g
-
gram
g L-1
-
gram per litre
h-1
-
per hour
L
-
litre
mg L-1
-
milligram per litre
mM
-
millimolar
mL
-
millilitre
nm
-
nanometer
%
-
percent
OD600
-
optical density at 600
S
-
substrate concentration (mg L-1 or g L-1)
So
-
initial substrate concentration (mg L-1 or g L-1)
C
t
time (h)
TL
-
lag period (h)
µ
-
specific growth rate (h-1)
µg L-1
-
microgram per litre
µL
-
microlitre
µm
-
micrometer
% v/v
-
percentage volume per volume
% wt/v
-
percentage weight per volume
Xmax
-
maximum biomass concentration (gdw L-1)
xxii
Yx/s
-
cell mass yield on phenol (g g -1)
Catmax
-
catechol maximum concentration (mg L-1 or g L-1)
Ypc/s
-
catechol yield on phenol (g g -1)
Ypc/x
-
catechol yield on cell mass (g g -1)
ccMAmax
-
cis,cis-muconic acid maximum concentration
(mg L-1 or g L-1)
YccMA/s
-
cis,cis-muconic acid yield on phenol (g g -1)
YccMA/x
-
cis,cis-muconic acid yield on cell mass (g g -1)
xxiii
LIST OF ABBREVIATIONS
ATCC
-
American Type Culture Collection
AGE
-
agarose gel electrophoresis
bp
-
base pairs
C1,2D
-
catechol 1,2-dioxygenase
ccMA
-
cis,cis-muconic acid
ccMALe
-
cis,cis-muconic acid lactonizing enzyme
CFU
-
colony forming unit
CIF
-
constant intermittent feeding
DNA
-
deoxyribonucleic acid
2-HMSA
-
2-hydroxymuconic semialdehyde
IPC
-
initial phenol concentration
HPLC
-
high-performance liquid chromatography
ITS
-
internal transcribed spacer
MCA
-
MacConkey agar
PCR
-
polymerase chain reaction
PH
-
phenol hydroxylase
psi
-
pounds per sq. in
rDNA
-
ribosomal deoxyribonucleic acid
RM
-
Ramsay medium
rpm
-
revolutions per minute
sp.
-
species
pH
-
hydrogen ion concentration
ppm
-
parts per million
RETL-Cr1
-
Ramsay Effluent of Treatment Lagoon-Cream 1
TCA
-
tricarboxylic acid cycle
TSI
-
triple sugar iron
UV
-
ultraviolet
xxiv
LIST OF APPENDICES
APPENDIX
A1
TITLE
PAGE
Plot of OD600 Vs dry weight during batch cultivationCalibration Curve for calculation of dry cell weight of
C. tropicalis RETL-Cr1.
A2
214
Standard Curve use to calculate glucose concentration
using Shimadzu Spectrophotometer Model based on
Sigma® procedure 510 (Sigma® Diagnostics,
St Louis, MO).
215
A3
Determination of glucose using Sigma® Procedure 510
216
A4
HPLC-analytical parameters for determination of
phenol, catechol and cis,cis-muconic acid.
A5
A6
B1
218
Heterotrophic Plate Count – Test Method
APHA 9215
219
Cellular Morphology and Biochemical Tests- Basic
Procedures.
220
Schematic representation for the biodegradation of
phenol by C. tropicalis REL-Cr1 isolated from
C
Exxon Mobil Oil Refinery treatment plant
225
Publications
226
1
CHAPTER 1
INTRODUCTION
1.1
Introduction
Environmental pollution has been considered as a side effect of industrial society.
Soil, lakes, rivers, and seas are highly contaminated with different toxic compounds
(Alexander, 1981). An example of such compound is phenol. Phenol is released into the
environment from industrial discharges (Keith, 1976; Jungclaus et al., 1978; Parkhurst et
al., 1979; Pfeffer, 1979) and spills (Delfino and Dube, 1976). According to Prasad and
Ellis (1978), phenols and its derivatives are among the most frequently found pollutants
in rivers, industrial effluents and landfill run-off waters. Hence, populations residing near
waste disposal sites, landfill sites or phenol spills may be at risk for higher exposure to
phenol than other populations. An example of such spill was one that occurred in June,
2001 when the Indonesian-registered oil tanker MT Endah Lestari capsized off the coast
of Johore, southern Malaysia spilling 600 metric tons of phenol and large amount of
diesel killing thousands of marine life in the nearby fish farming ground.
Nowadays, environmental preservation has become a key issue in a society
because it is often linked to quality of life. The impacts of pollution on the environment
have led to an intense scientific investigation. The removal of phenol from industrial
effluents has attracted researchers from different fields (Yang and Humphrey, 1975;
2
Shingler, 1996). The increasing awareness on the environment in both developed and
developing countries has initiated more studies of possible solutions for treating phenol.
Environmental biotechnology relies on the pollutant-degrading capacities of
naturally occurring microorganisms (Liu and Suflita, 1993). It has been reported to be
advantageous over physical and chemical treatments due to its relatively low cost and has
less ecological impact to the environment (Head, 1998; Edington, 1994). Researchers are
studying pollutant-degrading microorganisms which inhabit polluted environments
(Kumaran, 1980; Kapoor et al., 1998; Yap et al., 1999; Heinaru et al., 2000; Komarkova
et al., 2003; Santos and Linardi, 2004; Margesin et al., 2005) as well as uncontaminated
environment (Bastos et al., 2000a; Koutny et al., 2003). Harnessing the potential of
microbes (Ahmed, 1995; Fulthorpe and Allen, 1995; Bastos et al., 2000b; Ruiz- Ordaz et
al., 2001; Vojta et al., 2002; Páca Jr. et al., 2003) to degrade phenol has been an area of
considerable study to develop bioremediation approaches which has been considered as a
“green option” (Singleton, 1994) for treatment of environmental contaminants.
Many researchers support the biological treatment of phenols. A number of
studies with prokaryotic microorganisms have been carried out for the purpose to
improve the technological processes of biodegradation. Some examples are,
Pseudomonas sp. have demonstrated the ability to mineralize phenol (Ehrhardt and
Rehm, 1989; Hinteregger et al., 1992; Ahmed, 1995; Chitra et al., 1995; Dapaah and
Hill, 1992; Fulthorpe and Allen,1995; Fava et al., 1995; Loh and Wang, 1998),
Alcaligenes sp. (Hill et al., 1996; Valenzuela et al., 1997), Azotobacter sp. (Li et al.,
1991), Rhodococcus sp. (Apajalahti and Salkinoja-Salonen, 1986; Oh and Han, 1997),
Phanerochaete sp. (Perez et al., 1997; Larmar et al., 1990), and Cryptococcus sp.
(Mörsen and Rehm, 1987).
However, according to Katayama-Hirayama et al., (1994) information on
degradation of phenol is limited in the yeast strains. Among the eukaryotic
microorganisms, only some members of yeast genera Candida, Rhodotorula, and
Trichosporon that able to metabolize phenolic compounds as a sole carbon and energy
3
source (Neujahr, 1990; Katayama-Hirayama et al. 1994; Chen et al., 2002). Among the
Candida strain, Candida tropicalis has been the most studied in the biodegradation of
phenol (Shimizu et al., 1973; Kumaran, 1980; Krug et al., 1985; Bastos et al., 2000a;
Chen et al., 2002; Vojta et al., 2002; Yan et al., 2005). However, none of these yeast
strains were isolated from Malaysian environment.
Studies on the naturally pollutant-degrading microorganisms termed as
environmentally relevant microorganisms (ERM), include the isolation of bacteria from
the environment, their classification and physiological characterization, molecular
analysis of their degradative enzymes (Watanabe and Baker, 2000). Biodegradation of
phenol by many microorganisms has been studied in order to understand the nutrient
requirements, environmental physico-chemical factors, and complex biochemistry
involved that may assist in bioremediation of this toxic compound.
1.2
Objectives of the study
The aim of this study is to investigate the ability of locally isolated
microorganisms to degrade phenol with the specific objectives listed below:
1. To isolate, screen and identify phenol-degrading microorganisms from oil, waxy oil
and petrochemical wastes.
2. To optimize and conduct kinetic analyses on the aerobic phenol biodegradation in
batch and fed-batch cultures by potential strains.
3. To postulate possible metabolic pathway of phenol degradation by the microorganism
of interest.
4
4. To identify the potential strain by a molecular mechanisms (PCR amplification of
ribosomal DNA targeting the conserved regions of 5.8S, 18S and 28S using universal
primers ITS1 and ITS4).
5
CHAPTER 2
LITERATURE REVIEW
2.1
Phenol
Petroleum hydrocarbons can be divided into four classes namely saturates,
aromatics, the asphaltenes (phenol, fatty acids, ketones, esters and porphyrines), and
the resins (pyridines, quinolines, carbazoles, sulfoxides and amides) (Colwell and
Walker, 1977). Petroleum products have vast uses in this modern society. Phenol is
an important industrial chemical of environmental concern widely used in many
industries such as coke, refineries, manufacturers of resin, pharmaceuticals,
pesticides, dyes, plastics, explosives and herbicides, and can also occur in their
wastewaters (Lenke et al., 1992; Marvin-Sikkena and de Bont, 1994; Yang et al.,
1998). Phenols are produced in very large quantities for use as solvents, and starting
materials for chemical synthesis (Budavari, 1996).
Phenols and its derivatives are some of the major hazardous compounds in
industrial wastewater (Watanabe et al., 1996b; Peters et al., 1997). For instance,
phenol is released into water from industrial effluent discharges such as petroleum
refinery wastewater (Pfeffer, 1979). For the release in other industrial discharges, see
references Keith, (1976), Jungclaus et al., (1978), Parkhurst et al., (1979), and
Hawthorne and Sievers, (1984). Phenol has been also detected in groundwater as a
6
result of leaching through soil after a spill of phenol (Delfino and Dube, 1976), from
landfill sites (Clark and Piskin, 1977), and from hazardous waste sites (Plumb, 1987).
Phenols have relatively high water solubility and widely known to be acutely
toxic to a range of organisms. It produces undesirable taste, odour, colour to water
and is considered toxic (Klibanov, 1982). Therefore, this compound needs to be
disposed off in a safe and environmentally acceptable way.
2.1.1 Chemical identity, physical and chemical properties of phenol
Phenol, C6H5OH (Pronounced fƝ'nôl') or hydroxybenzene, is an aromatic
molecule containing hydroxyl group attached to the benzene ring structure (Figure 2.1).
Phenol commonly known as carbolic acid (Gardner et al., 1978) has a molecular weight
of 94.11 gm/mole (Lide, 1993). It has a melting point of 43oC and forms white to
colourless crystals (Budavari et al., 1989), colourless to pink solid or thick liquid
(NIOSH, 1985; HSDB, 1998). It has a characteristic of acrid smell and a sharp burning
taste. Phenol have relatively high water solubility and it is soluble in most organic
solvents such as aromatic hydrocarbons, alcohols, ketones, ethers, acids, halogenated
hydrocarbons (Kirk and Othmer, 1980; Lide, 1993.) However, the solubility is limited in
aliphatic solvents. The odour threshold of phenol in air is 0.040 ppm (v/v) (Amoore and
Hautala, 1983) and in water between 1 ppm and 7.9 ppm (w/v) (Baker et al., 1978;
Amoore and Hautala, 1983).
OH
Figure 2.1 Chemical structure of phenol
7
2.2
Sources of phenol
The origin of phenol in the environment is from natural, man-made and
endogenous sources. Phenol is released primarily to the air and water as a result of its
manufacture and use, wood burning and auto exhaust. Phenol mainly enters waters from
industrial effluent discharges.
2.2.1 Natural sources
Phenol is a constituent of coal tar, and is formed during decomposition of organic
materials. Increased environmental levels may result from forest fires (Hubble et al.,
1981). It has been detected among the volatile components from liquid manure at
concentrations of 7-55 ug/kg dry weight (Spoelstra,1978) and has an average
concentration in manure of 30 ug/kg dry weight (RIVM, 1986).
2.2.2 Man-made sources
Man-made sources are from industrial wastes from fossil fuel extraction, chemical
manufacturing processes such as phenol manufacturing plants, pharmaceutical industry,
wood processing industry and pesticide manufacturing plants (Kumaran and Parachuri,
1997). Industrial sources of phenols and other related aromatics are from petroleum
refinery, petrochemicals, basic organic chemical manufacture, coal refining,
pharmaceuticals, tannery and pulp and paper mills (Table 2.1) (Kumaran & Paruchuri,
1997).
8
2.2.3 Endogenous sources
An important additional source of phenol may be the formation from various
xenobiotics such as benzene (Pekari et al., 1992) under the influence of light (Hoshino
and Akimoto, 1978).
Table 2.1: Sources of phenols and other related aromatic compounds in wastewater
(Kumaran and Paruchuri, 1997).
Sources
Significant phenolic compounds
Petroleum refining
Hydrocarbons (alkanes, cycloalkanes, polyaromatic
hydrocarbons), benzenes, substituted benzenes, toluenes,
n-octanes, n-decanes, naphthalenes, biphenyles, phenol,
cyanide, sulphide and ammonia.
Petrochemicals
Naphthalene, hepatanes, benzenes, butadiene, C-4
alcohols, phenol and resorcinol.
Basic organic chemical
Manufacturing
m-amino phenol, resorcinol, dinitrophenol, pnitrophenol,trinitrophenol, benzene sulphonic acids, aniline,
chlorobenzenes, toluene and resorcinol.
Coal refining
Phenol, catechol, o-, m-, p-cresols, resorcinol,
hydroquinone, pyrogallol, polyaromatic hydrocarbons,
pyridine, pycolines, lutidines, xylenes, toluenes, benzoic
acid.
Pharmaceuticals
Toluenes, benzyl alcohols, phenyl acetic acid, chlorinated
products of benzene, chloroform, ether, ethyl alcohol.
Tannery
Tannin, catechin, phenol, chlorophenol, nitrophenols.
Pulp and paper mills
Lignin, vanillin, vanillic acid, dehydrodivanillin, ferulic
acid, cinnamic acid, synringic acid, vieratric acid,
protocatechuic acid, gentisic acid, benzoic acid,
guadiachols, catechol, coniferyl alcohol,
dehydrodihydroconiferyl alcohol, phenyl propionic acid,
phenols and chlorophenols.
9
The annual production of phenols are estimated around 1.25 x 109 kg (BČchard et
al., 1990). In 1995, the total annual capacity of phenol production approached 4.5 billion
pounds (CMR, 1996). The most commonly used production method for phenol is from
cumene (isopropylbenzene) (IARC, 1989). Phenol is also produced from chlorobenzene
and toluene. It is the basic feedstock from which a number of commercially important
materials are made, including phenolic resins, bisphenol A (2, 2-bis-1hydroxyphenylpropane), capro-lactam, alkyl phenols, chlorophenols such as
pentachlorophenol (IARC, 1989). Phenolic resins are used as a binding material in,
insulation material, chipboard and triplex, paints and casting sand foundries. Phenols are
environmental pollutants commonly present in the wastewaters from oil industry.
2.3
Release of phenol into the environment
Man-made phenolic compounds are found in the environment in abundance, due
to agricultural and industrial activities. It has been reported that an estimated total of
23.5 million pounds (10.6 million kg) of phenol was released to the environment from
689 large processing facilities (TRI, 1998). Phenol has been found in surface water,
ground water, soil and sediment (HazDat, 1998).
2.3.1 Air
The estimated releases of phenol of 9.5 million pounds (4.3 million kg) to air
from 635 large processing facilities accounted for about 5% of environmental releases
(TR1, 1998). During manufacturing, phenol is released to the atmosphere from storage
tank vents during transport loading (Delaney and Hughes, 1979).
Other major sources of release to the atmosphere are from residential wood
burning and automobile exhaust (Scow et al., 1981). Phenol has been detected in the
exhaust gases of private cars at concentration of 0.3 ppm (approximately 1.2 mg/m3) to
10
1.4-2.0 ppm (5.4-7.7 mg/m3) (Kuwata et al., 1980; Verschueren, 1983).
Phenol has been detected from other sources such as emissions from waste
incinerator plant at 0.36 ppb (Jay and Stieglitz, 1995), in cigarette smoke and plastics
(Graedel, 1978). It has been identified in cigarette smoke, in quantities that are
comparable to an average emission of 0.4 mg/cigarette (Groenen, 1978). Emission gases
from all material incinerators and home fires, especially wood-burning, may contain
substantial quantities of phenol (Den Boeft et al., 1984).
Volatilization from environmental waters and soil has been shown to be a slow
process and not expected to be a significant source of phenol in the atmosphere.
2.3.2 Water
It has been reported that an estimated of 72,550 pounds (32,650 kg) of phenol
releases to water from 230 large processing facilities accounted for about 0.3% of total
environmental releases (TRI, 1998).The most common anthropogenic sources of phenol
in natural water include coal tar (Thurman, 1985) and waste water from manufacturing
industries such as resins, plastics, fibers, adhesives, iron, steel, aluminum, leather, rubber,
and influents from synthetic fuel manufacturing (Parkhurst et al., 1979). Phenol is also
released from paper pulp mills (Keith, 1976) and wood treatment facilities (Goerlitz et
al., 1985). Phenol has been detected in the effluent discharges of a variety of industries.
Levels of phenol concentration in wastewater from selected industries are shown in Table
2.2.
11
Table 2.2: Typical levels of phenol concentration in wastewater of some selected
industries
Selected industry
Phenol concentration
(mg L-1)
Reference
Phenol production
3,000-4000
Godjevargova et al., 2003
Pulp and paper
33.1-40
Peralta-Zamora et al., 1998;
Minussi et al., 1998
Textile
12.3
Kunz et al., 2001
Olive oil mill
3000-10,000
Klibanov et al., 1983;
Borja et al., 1992;
Hamdi, 1992;
Martinez-Neito et al., 1992;
Knupp et al., 1996;
Robards and Ryan, 1998
Coal conversion plant
4-4780
Parkhurst et al., 1979
Shale oil wastewater
4.5
Hawthorne and Sievers, 1984
Ash-heap water (oil shale)
500
Kahru et al., 1998
Phenolic resins production
1200->10,000
Patterson, 1985;
Kavitha and Palanivelu, 2004
Methyl violet and cumenephenol production
310-660
Kanekar et al., 1999
Chemical specialitiesmanufacturing
0.01-0.30
Jungclaus et al., 1978
Petroleum oil refinery
33.5
Pfeffer, 1979
Other release of phenol results from commercial use of phenol and phenolcontaining products, including slimicides, general disinfectants (Hawley, 1981; Budavari
et al., 1989), medicinal preparations such as ointments, ear and nose drops, cold sore
lotions, mouthwashes, gargles, toothache drops, analgesic rubs, throat lozenges (USEPA,
1980), and antiseptic lotions (Musto et al., 1977). It has been estimated that 3.8 kg/day of
12
phenol release to seawater from municipal treatment facilities (Crawford et al., 1995).
Animal and decomposition of organic wastes are the two natural sources of phenol in
aquatic media.
2.3.3 Soil
In 1996, the estimated releases of 159,059 (71,577 kg) of phenol to soil from 102
large processing facilities accounted for about 0.7% of total environmental releases (TRI,
1998). Phenol are released to the soil during its manufacturing process, loading and
transport when spills occur, and when it leaches from hazardous wastes sites and landfills
(Xing et al., 1994). According to ASTDR, (1998) generally the data on concentrations of
phenol found in soil at sites other than hazardous sites are lacking. This may be due to a
rapid rate of biodegradation and leaching. Phenol can be expected to be found in soils
that receive continuous or consistent releases from a point source. Phenol that leaches
through soil to groundwater spends at least some time in that soil as it travels to the
groundwater. Phenol has been found in groundwater, mainly at or near hazardous wastes
sites.
2.4
Fate of phenol in the environment
2.4.1
Air
There has been no data have been found concerning wet and dry deposition of
phenol. Dry deposition (by particle deposition) is expected to be negligible since phenol
in air is almost exclusively in gas phase (IPCS, 1994). The theoretical deposition rates for
phenol were estimated assuming a behavior similar to SO2 (IPCS, 1994), and when
comparing with the rate of reaction of phenol with hydroxyl radicals, it was concluded
that most phenol in the atmosphere is degraded chemically, rather than transported
(RIVM, 1986).
13
Phenol absorbs light in the region of 290-330 nm (Sadtler, 1960), and therefore
might photodegrade directly in the atmosphere (Howard, 1989). Phenol may react in air
with hydroxyl and NO3 radicals, and undergo other photochemical reactions to form
dihydroxy-benzenes, nitrophenols, and ring cleavage products (Atkinson et al., 1979;
Bruce et al., 1987). The gas-phase reaction of phenol with photochemically produced
hydroxyl radicals is probably the removal mechanism in the atmosphere. The half-life of
phenol in air is 5 h based on its estimated reaction rate with hydroxyl radicals (RIVM,
1986). Howard, (1989) estimated a half-life of 15 h for reaction of phenol with hydroxyl
radicals in air.
The reaction of phenol with nitrate radicals during the night may be a significant
removal process; a half-life of 15 min has been estimated at an atmospheric concentration
of 2x108 nitrate radicals per cm3 (Atkinson et al., 1987; Howard, 1989). The reaction of
phenol with nitrate radicals present in the atmosphere during smog episodes may
decrease the half-life of phenol in polluted atmospheres. The above data indicates that
phenol has a short half-life in the atmosphere, probably less than 1 day (ATSDR, 1998).
It has been concluded that deposition may contribute to the disappearance of
phenol from the atmosphere. A relatively high concentration has been found in rain water
(Leuenberger et al., 1985). The fact that it has been detected in rain water, some phenol
may wash out of the atmosphere in limited amounts because of the short atmospheric
half-life of phenol (ATSDR, 1998). Therefore, transport of phenol from air to soil and
water is likely (RIVM, 1986).
2.4.2 Soil and sediment
Partition coefficient (KOC) values of phenol for two silt loams were reported to be
39 and 91 dm3/kg. Based on this KOC values, phenol would be expected to be highly
mobile in soil, and therefore may leach to groundwater (Howard, 1989). This was
confirmed by Scott et al., (1982) who found that low adsorption of phenol to two sterile
14
silt loams (pH 5.4, organic matter content 1.1 and 3.6, respectively). The moderately low
sorption partition coefficient (1.21-1.96) suggests that sorption to sediment is not an
important transport process. Ehrlich et al., (1982) reported that there was very little
sorption of phenol onto aquifers materials thus suggested that phenol sorption to
sediments may be minimal.
Based on the soil sorption coefficient, phenol released to soil is expected to leach
to ground water. However, the rate of phenol degradation in the soil may be rapid, except
in cases of large releases such as spills or continuous releases such as leaching from
landfill sites, the probability of groundwater contamination may be low (Ehrlich et al.,
1982). According to Xing et al., (1994), sorption coefficient for phenol by soil increases
with increasing soil organic matter which may indicate that soil organic matter may act as
a primary phenol sorbent in soil. Volatilization from dry-near surface soil should be
relatively rapid (Howard, 1989).
Phenol is degradable in soil under both aerobic and anaerobic conditions. The
half-life of phenol in soil is generally less than 5 days (Baker and Mayfield, 1980; HSDB,
1998) but acidic soils and some surface soils may have half-life between 20 and 25 days
(HSDB, 1998). Haider et al., (1974) found that mineralization in alkaline soil under
aerobic conditions was 45.5 after 3 days, 48% after 7 days and 65% after 70 days. The
half-lives for degradation of low concentration of phenol in silt loam soils were between
2.7 and 3.5 hours (Scott et al., 1982). Phenol degradation under anaerobic conditions is
slower. However, phenol can be degraded completely in soil under both aerobic and
anaerobic conditions, and phenol is not expected to be absorbed to sediment (HSDB,
1998).
Plants can readily uptake phenol (Cataldo et al., 1987) however, bioaccumulation
does not take place due to high rate respiratory decomposition of phenol to CO2 (ATSDR,
1998).
15
2.4.3 Water
Phenol is highly soluble in water and relatively low in vapour pressure at room
temperature. With these properties, phenol is expected to end up largely in the water
phase upon distribution between air and water.
Phenol absorbs light in the region of 290-330 nm (Sadtler, 1960), thus it might
photodegrade directly in surface waters. Phenols react relatively rapid in sunlit natural
water via reaction with photochemically produced hydroxyl radicals and peroxy radicals.
According to Mill and Mabey (1985), the typical half-lives for hydroxyl and peroxyl
radical reactions are on the order of 100 and 19.2 hours of sunlight, respectively. The rate
constant for the reaction of phenol with ozone in water has been reported to range from
1.5x10-5 to 6x10-5 milliseconds-1 (Beltran and Alvarez, 1996).
Phenol has been detected in ground water as a result of leaching through soil from
a spill of phenol (Delfino and Dube, 1976), from landfill sites (Clark and Piskin, 1977),
and from hazardous waste sites (Plumb, 1987). Phenol is readily biodegradable in natural
water, provided the concentration is not high enough to cause inhibition. A complete
degradation of phenol is less than 1 day in water from lakes (Rubin and Alexander, 1983)
and river after 2-4 days depending on the temperature (Ludzack and Ettinger, 1960).
However, degradation of phenol was reported to be slower in salt water, and a half-life of
9 days has been reported in an estuarine river (Lee and Ryan, 1979).
While the evidence cited above suggest that phenol can rapidly degraded in
natural water, but it may still present in the environment is because the exact conditions
under which phenol is rapidly degraded are not present in all instances. In some situation,
phenol concentration may be too high or the population of microorganisms may not be
present in sufficient numbers for significant biodegradation to occur.
Phenol was found not to bioconcentrate in aquatic organisms. Reported log
bioconcentration factors (BCF) in fish for phenol include 0.28 for goldfish (Kobayashi et
16
al., 1989) and 1.3 for golden orfe (Freitag et al., 1984). According to Nicola et al., (1987)
the highest mean level of phenol detected in bottom fish was 0.14 ppm.
2.5
Hazards of phenol
Aromatic hydrocarbons are not as readily biodegradable as the normal and
branched alkanes, they are somewhat more easily degradable than the alicyclic
hydrocarbons (Perry, 1984; Leahy and Colwell, 1990). Many of these compounds are
toxic and some are known or suspected carcinogens (Verschueren, 1977; Klibanov, 1982;
Kuhn et al., 1989; Nicell et al., 1993; Bryant and Schultz, 1994; Sheeja and Murugesan,
2002). The presence of phenol in drinking water and irrigation water represents a serious
health hazards to humans, animals, plants and microorganisms (Shailubhai, 1986;
Salonen et al., 1989; Sharma et al., 1997).
Phenol concentrations greater than 50 ppb are toxic to some form of aquatic life
and ingestion of 1 g of phenol can be fatal in human beings (Seetharam and Saville,
2003). Continuous ingestion of phenol for a prolonged period of time causes mouth sore,
diarrhea, excretion of dark urine and impaired vision at concentrations levels ranging
between 10 and 240 mg L-1 (Barker et al., 1978). Lethal blood concentration for phenol is
around 4.7 to 130 mg/100 ml. Phenol affects the nervous system and key organs, i.e.
spleen, pancreas and kidneys (Manahan, 1994).
Phenol is lethal to fish even at relatively low levels, e.g. 5-25mg/L, depending on
the temperature and state of maturity of rainbow trout (Brown et al., 1967). Phenolic
compounds are also responsible for several biological effects, including antibiosis
(Rodriguez et al., 1988; Gonzalez et al., 1990), ovipositional deterrence (Girolami et al.,
1981) and phytotoxicity (Capasso et al., 1992).
Phenol is classified as a priority pollutant owing to their high toxicity and wide
spread environmental occurrence (USEPA, 1984a, 1984b). Various regulatory authorities
17
have imposed strict limits to phenol concentration in industrial discharges. Phenol is
released into the environment is regulated by many countries (CEPA, 2001; USEPA,
1998, Sa and Boaventura, 2001). For drinking waters, it has been prescribed a guideline
concentration of 1 µg L-1 (WHO, 1994). In Malaysia, the Environmental Protection Act,
1974 establish a phenol concentration of 0.001 mg L-1 for Standard A, 0.1 mg L-1 for
standard B, and 5 mg L-1 other than standard A and B as the limit for wastewater
discharges into inland waters. Therefore, the disposal of phenol has become a major
global concern (Percival and Senior, 1998).
2.6
Microbial degradation
Microbial degradation of chemicals in the environment is a route for their
removal. The microbial degradation of these compounds is a complex series of
biochemical reactions and often different when different microorganisms are involved.
The interdependence of biodegradation, biotransformation and biocatalysis has been
reviewed by Parales et al., (2002). Microbial degradation of pollutants is crucial in order
to predict their longevity and long term effects and also important in the actual
remediation process (Landis and Yu, 2003).
In aerobic respiration, oxygen acts as the electron acceptor. Molecular oxygen is a
reactant for oxygenase enzymes and is incorporated into the final products. In anaerobic
respiration, different inorganic electron acceptors are possible such as NO3-, SO42-, S0,
CO2 and Fe3+. Most of the documentation on microbial degradation of organic pollutants
in nature is focused on aerobic transformation. Many synthetic compound compounds
accumulate in nature because the release rates exceed the rates of microbial and chemical
degradation (Harms and Bosma, 1997). In addition, many microbial transporters and
catabolic are regulated, i.e. they are only synthesizes in response to the presence a certain
concentration of their substrate (Spain et al., 1980; Spain and van Veld, 1983).
18
There are two major reasons for low degradation rates have been identified. First,
the biochemical potential to degrade certain compound is limited. This is more likely that
less chemicals resembles natural compounds (Reineke and Knackmuss, 1978; Alexander,
1981; Van deer Meer et al., 1992). Secondly, the pollutant or other substrates, e.g.
appropriate electron acceptors are unavailable to the microflora (Lyngkilde and
Christiansen, 1992; Mihelcic et al., 1993; Bosma et al., 1996).
In the natural environment, the rate of degradation can be depended on physical,
chemical and biological factors which may differ among ecosystems (Melcer and Bridle,
1985). Alexander (1994) reported that for a microbial transformation to occur, a number
of conditions must be satisfied. These include: 1) Microorganisms must exist with the
required enzyme to catalyze the specific transformation. There are unspecific enzymes
that can attack several types of substrates, while other enzymes can only catalyze the
breakdown of one specific bond in a specific compound. Duetz et al., (1994) reported
that different bacterial strains may also degrade same compound by different degradation
patterns, depending on the types of enzymes used. Many degradation pathways are
achieved only by the synergistic relationship of several species (Lappin et al., 1985), 2)
The chemical must be made available for the microorganism. The inaccessibility may be
resulted from the chemical existing in a different phase from the bacteria, for example, in
a liquid phase immiscible with water, or sorbed to a solid phase, 3) The success of the
degrading strains to proliferate will depend on their ability to compete for the organic
compound, oxygen and other environmental factors.
2.6.1. Phenol-degrading microorganisms
Microorganisms that can degrade phenol were isolated as early as 1908 (Evans,
1947). The key components of microbial communities responsible for degradation of
phenolic wastes are Pseudomonads species. Their physiological and genetic basis of
phenol degradation has been described by many researchers ( Kotturi et al., 1991; Nurk et
al., 1991; Topp and Akhtar, 1991; Kiyohara et al., 1992; Motzkus et al., 1993; Arquiaga
et al., 1995; Puhakka et al., 1995; Srivastava et al., 1995; Buitron and Gonzalez, 1996;
19
Loeser et al., 1998).
Phenols are metabolized by microorganisms from a variety of different genera
and species, as shown in Table 2.3. Bacteria, fungi, yeast and algae have been reported to
be capable of degrading phenol. As shown in Table 2.3, Pseudomonas putida has been
extensively investigated and has been reported to be capable of high rates of phenol
degradation (Hutchinson and Robinson, 1988). According to Whiteley et al., (2001)
isolates that were able to utilize phenol as a sole carbon source predominantly belonged
to Pseudomonas pseudoalcaligenes. The earlier reports on the decomposition of phenolic
compounds by yeasts were by strains belonging to the genera Oospora, Saccharomyces,
Candida, Debaryomyces and Trichosporon cutaneum (Harris and Ricketts, 1962;
Henderson, 1961; Neujahr and Varga, 1970; Neujahr et al., 1974; Hashimoto, 1970,
1973). Among the yeast strains, Candida tropicalis has been the most studied and able to
degrade phenol, phenol derivatives and aliphatic compounds at a relatively high phenol
concentration (Krug et al., 1985; Chang et al., 1995,1998; Ruiz-Ordaz et al., 1998, 2000).
According toYap et al., (1999) mutant strain of Comamonas testosteroni E23 has been
regarded as the best phenol degrader of all phenol degrading strains reported up to date.
20
Table 2.3: Phenol-degrading microorganisms
Microorganism Degradation parameters Performance
(A, AN,FC, IC, SS, MS) (mg L-1 h-1)
Reference
A. Bacteria
Acinetobacter sp.
7.7
36-38
16.7
4.2
6.9-12.2
4.2
20-33
Tibbles and Baecker, 1989b
Hao et al., 2002
Hao et al., 2002
Beshay et al., 2002
Beshay et al., 2002
Abd-EL-Haleem et al., 2003
Abd-EL-Haleem et al.,2003
- A. calcoaceticus AH A,FC,SS
7.6-25
Nakamura and Sawada, 2000
- A. johnsonii
11.8
Heilbuth et al., 2003
Achromobacter sp.E1 A,FC,MS
0.5
Watanabe et al., 1996a
Alcaligenes faecalis A,FC,SS
4.7-7.4
Bastos et al., 2000b
- A. sp. E2
-A. sp. R5
-A. strain P5
A,FC,MS
A,FC,MS
A,FC,SS
0.4
0.8
0.1-0.2
Watanabe et al., 1996a
Watanabe et al., 1996a
Baek et al., 2001
Arthrobacter sp.
A,FC,SS
A,FC,MS
A,FC,SS
83
62.5
9.4
Kar et al., 1996
Kar et al., 1996
Tibbles and Baecker, 1989b
Azoarcus sp.
AN,FC,SS
1.8
Shinoda et al., 2000
Azospirillium –
brasilense
A,FC,SS
1.0
Barkovskii et al., 1995
- A. sp. W-17
A,FC,SS
A,FC,SS
A,FC,MS
A,FC,SS
A,IC,SS
A,FC,SS
A,IC,SS
A,FC,SS
B. thermoleovoransA2
A,FC,SS
7.8-19.6
Mutzel et al., 1996
A = aerobic; AN = anaerobic; FC = free cells; IC = immobilized cells; SS = single
substrate; MS = mixed substrates
21
Table 2.3: Phenol-degrading microorganism -
continue
Microorganism Degradation parameters Performance
Reference
(A, AN, FC, IC, SS, MS) (mg L-1 h-1)
Burkholderiacepacia G4
A,FC,SS
A,FC,MS
A,FC,MS
A,FC,MS
Comamonastestosteroni
-P15
-E23 (mutant)
A,FC,SS
A,FC,SS
Halomonas sp
A,FC
249
218
141
153
44
40-70
Moustafa El-Sayed, 2003
Moustafa El-Sayed, 2003
Moustafa El-Sayed, 2003
Moustafa El-Sayed, 2003
Yap et al., 1999
Yap et al., 1999
8
Hinteregger and Streichsbier,
1997
Halophilic bacteria
CA00, CA08, SL03,
SL08, SP04
A,FC,SS
3-4
Peyton et al., 2002
Iron-reducingorganism GS-15
2
Lovley and Lonergan, 1990
Magnetospirillum sp. AN,FC,SS
2
Shinoda et al., 2000
Micrococcus sp.
A, FC,SS
5
Tibbles and Baecker, 1989b
Nocardia sp.
A,FC,SS
15
Tibbles and Baecker, 1989b
Ochrobactrum tritici
A,FC,SS
3
El-Sayed et al., 2003.
AN, FC,MS
Phormidium –
valderianumBDU 30501
A,FC,SS
<1
Shashirekha et al., 1997
A = aerobic; AN = anaerobic; FC = free cells; IC = immobilized cells; SS= single
substrate; MS = mixed substrates
22
Table 2.3: Phenol-degrading microorganism –
continue
Microorganism
Reference
Degradation parameters
Performance
(A, AN, B, FC, IC, SS, MS) (mg L-1 h-1)
Pseudomonas sp.
A,FC,MS
14-28
Kang and Park, 1997
-P. pictorumNICM-2077
A,FC,SS
20-46
Sheeja and Murugesan, 2002
A,IC,SS
A,FC,SS
A,FC,SS
A,IC,SS
A,IC,SS
17
5.0
30-40
20-110
250-297
Loh and Liu, 2001;
Collins and Daugulis, 1997a
Mordocco et al., 1999
Mordocco et al., 1999
Hannaford and Kuek, 1999
152-238
Hughes and Cooper, 1996
A,F,SS
A,I,SS
A,FC,SS
A,IC,SS
4.2
3.5
14
3-4
Gonzalez et al. 2001a
Gonzalez et al. 2001a
Tarighian et al. 2001
Gonzalez et al. 2001b
A,FC,SS
A, B,SS(SFPB)
14
70-58
Daraktchiev et al, 1996
Daraktchiev et al, 1996
A,FC,SS
8-22
Wang and Loh, 1999
3-8
10-18
Abuhamed et al., 2003
Abuhamed et al., 2004
- P. putida
ATCC 11172
- P. putida –
ATCC 12633
- P. putida –
ATCC 17484
-P. putida –
ATCC 21812
- P. putida –
ATCC 49451
- P. putida F1ATCC 700007
- P. putida F1
A,FC,SS,(SCF)
A,FC,SS
A,FC,SS
A,FC,SS
0.5
Reardon et al., 2000
A,FC,MS
0.8-0.9
Reardon et al., 2000
A,FC,MS
1
Reardon et al., 2002
A = aerobic; AN = anaerobic; B = Biofilm; FC= free cells; IC = immobilized cells; SS =
single substrate; MS= mixed substrates; SCF = self-cycling fermentation; SFPB = semifixed packing bioreactor
23
Table 2.3: Phenol-degrading microorganism-
continue
Microorganism Degradation parameters
Reference
Performance
(A, AN, FC, IC, SS, MS) (mg L-1 h-1)
A,FC,SS
34-63
- P. putida BH
- P. putida BH(ps10-45)(GEM) A,FC,SS
Soda et al., 1998
40-91
Soda et al., 1998
A,FC,SS
A,IC, SS
1-20
1-12
Chung et al., 2003
Chung et al., 2003
- P. putida –
DSM 548
A,FC,SS
2
Monteiro et al., 2000
- P. putida EKII
A,FC,SS
8-13
Hinteregger et al., 1992
A,FC,SS
A,IC,SS
4-9
5-10
A,IC,SS
3-11
A,IC,SS
10-19
Bandhyopadhyay et al., 1998
Bandhyopadhyay et al.,
2001
Mahadevaswamy et al.,
2004
Banerjee et al., 2001
A,FC,SS
3
Kotturi et al., 1991
A,FC,SS
38-48
Ahamad and Kunhi, 1996
A,FC,MS
<1
Kim et al., 2002
Ralstonia eutrophaATCC 17697
A, FC,SS
50
Léonard et al., 1999
18
86
Pai et al., 1995
Pai et al., 1995
- P. putida –
CCRC14365
- P. putida –
MTCC 1194
- P. putida Q5
- P. stutzeristrain SPC2
P. testosteroniCPW301
Rhodococcussp. DCB-p0610
A,FC,SS
A,IC,SS
- R. erythropolisUPV-1
A,FC,SS
14-27
Prieto et al., 2002
A,IC,SS
20
Prieto et al., 2002
A,FC,SS
3-13
Hidalgo et al., 2002
Sulfate-reducingbacteria
AN, FC, SS
1
Boopathy, 1995
A = aerobic; AN = anaerobic; FC = free cells; IC = immobilized cells; SS = single
substrate; MS= mixed substrates.
24
Table 2.3: Phenol-degrading microorganismMicroorganism
Degradation parameters
(A, AN, FC, IC, SS, MS)
continue
Performance Reference
(mg L-1 h-1)
Mixed bacterialcultures
Mixed bacteria
0.8-2
Ha et al., 2000
Mixed methanogeniccultures
AN,FC, SS
<1-4
Karlsson et al., 1999
Arthrobacter sp +,
B. cereus, C.
Freundii, M. agilis,
P. putida b. B
A,FC, SS
7-14
Kanekar et al., 1999
Bacteria + E. coliATCC 33456
AN, FC,MS
<1
Chirwa and Wang, 2000
Clostridium ghonii,
C. hastiforme, C. glycolicum)
AN, FC,SS
<1
Létourneau et al., 1995
P. putida F1 +
B. strain JS150
A,FC,SS
2-3
A,FC,MS
1
Rogers and Reardon, 2000;
Reardon et al., 2002,
Reardon et al., 2002
AN,FC,SS
0.8-1
Boopathy, 1997
SRB and AUMB
A,B,(GAC),MS
B. Fungi
Aspergillusterreus
A,FC,SS
3-7
Garcia et al., 1997, 2000
-A. niger
A,FC,SS
8
Garcia et al., 2000
- A. LA2,
LA3, AE5
A,FC,SS
<1-4
Santos and Linardi, 2004
Fusarium
- F. FE11, FE16
A,FC,SS
<1-4
Santos and Linardi, 2004
A = aerobic; AN = anaerobic; B= Biofilm; FC = free cells; IC = immobilized cells; SS =
single substrate; MS = mixed substrates; SRB = sulfate-reducing bacteria; AUMB=
acetate-utilizing methanogenic bacteria; GAC = granular activated carbon
25
Table 2.3: Phenol-degrading microorganism -
continue
Microorganism Degradation Parameters Performance
Reference
Coprinus sp.
C. cinereus
C. micaceus
A,FC,SS
A.FC,SS
A,FC,SS
0.8
0.8
0.8
Guiraud et al., 1999
Guiraud et al., 1999
Guiraud et al., 1999
Graphium LE6,
LE11,LA1, LE9,
LA5,FIB4,AE2
A,FC,SS
4
Santos and Linardi, 2004
Geotrichumcandidum
A,FC,SS
<1-3
Garcia et al., 1997, 2000
Penicillium
AF2, AF4,FIB9
A,FC,SS
<1-4
Santos and Linardi, 2004
Pleurotus
ostreatus
AN,FC,SS
6-13
Fountoulakis et al., 2002
Phanerochaetechrysosporium
A,FC,SS
8
Garcia et al., 2000
A,FC,SS
A,FC,SS,(RBMBC)
7-29
99-191
Bastos et al., 2000a
Ruiz-Ordaz et al., 2001
A,FC,SS,(FB)
157
Komarkova et al., 2003
A,FC,SS
A, IC, SS
0.9-10
6-7
Chen et al., 2002
Chen et al., 2002
(A, AN, FC, IC, SS, MS) (mg L-1 h-1)
C. Yeast
Candida tropicalis
- C. tropicalis
-C. tropicalisCt2
-C. tropicalis –
NCYC 1503
-C. tropicalis
A,FC,SS
30
Yan et al., 2005
A = aerobic; AN = anaerobic; FC = free cells; IC = immobilized cells; SS = single
substrate; MS = mixed substrates; FB = Fed-batch; RBMBC = repeated batch multistage
bubble column.
26
Table 2.3: Phenol-degrading microorganism Microorganism
Degradation Parameters
continue
Performance Reference
(A, AN, FC, IC, SS, MS) (mg L-1 h-1)
Rhodotorulaglutinis ATCC
28052
A,FC,SS
26
Katayama-Hirayama et al.,
1994
Trichosporoncutaneum R57
(mutant)
A,FC,SS
50-63
Alexieva et al., 2004
D. Alga
Ochromonas danica
A,FC,SS
24
Semple and Cain, 1995
A,FC.MS
12
Semple and Cain, 1995
A = aerobic; AN = anaerobic; FC = free cells; IC = immobilized cells; SS = single
substrate; MS = mixed substrates
2.6.2
Phenol-degrading Candida tropicalis
An investigation on the origin of the phenol-degrading Candida tropicalis by
other researchers previously was attempted, as shown in Table 2.4.
Yeasts are widely distributed in nature and have extremely diverse metabolic
capabilities and can utilize a wide range of nutrients under a variable of environmental
conditions (Tornai-Lehoczki et al., 2003). Among the yeast species, Candida tropicalis
utilized a very large variety of carbon sources including many sugars, disaccharides,
alkanes, alkane derivatives, fatty acids and phenols (Kurihara et al., 1992; Kawachi et al.,
1997). Other industrial importance of Candida tropicalis are production of xylitol
(Yahashi et al., 1996; Azuma et al., 2000; Walther et al., 2001; Lima et al., 2003), crude
oil-utilizers (Murzakov et al., 2003), and production of microbial protein and fodder
yeast (Stanton and Dasilva, 1978).
27
Table 2.4: Source of origin of phenol degrading Candida tropicalis
Candida tropicalis
Source of origin
Reference
C. tropicalis
activated sludge
Yan et al., 2005
C. tropicalisYMEC14
olive mill wastewater
Ettayebi et al., 2003
C. tropicalis ct2
C. tropicalisNCYC 1503
C. tropicalis
activated sludge of an industrialwastewater treatment plant
Komarkova et al., 2003
Vojta et al., 2002
Bryndová, 2002
NA
Chen et al., 2002
soil from pristine Amazonrain forest
Bastos et al., 2000b
C. tropicalis
NA
C. tropicalis CHP4
phenol-bearing industrialwastes
Klein et al., 1979
Neujahr and Gaal, 1973
Stephenson, 1990
Kumaran, 1980
C. tropicalis H15
NA
Krug et al., 1985
Krug and Straube, 1986
C. tropicalis 708
NA = not available
NA
Shimizu, 1973
As shown in Table 2.4, C. tropicalis capable of degrading phenol were found both
in contaminated and pristine ecosystem.
2.6.3 Aerobic biodegradation of phenol
Microorganisms have been isolated that grow on phenol (Murray and Williams,
1974; Hutchinson and Robinson, 1988). Microorganisms that can degrade phenol were
28
isolated as early as 1908 (Evans, 1947). Bacteria play a major role in the degradation of
phenol in the ecosystem: in soil (Hickman & Novak, 1989), sediments (Shimp & Young,
1987) and water (Howard, 1989). Despite being toxic, phenol can be utilized by microbes
as carbon and energy sources (Gibson, 1968: Gibson et al., 1990; van Schie and Young,
2000). The number of bacteria capable of utilizing phenol is usually a small percentage of
the total population present in, for example, a soil sample (Hickman & Novak, 1989).
Many soil and litter inhabiting bacteria fungi can degrade aromatic compounds
(Gibson and Subramaniam, 1984). However, repeated exposure may result in acclimation
(the promotion of strain capable of utilizing phenol as food) (Young & Rivera, 1985;
Colvin & Rozich, 1986; Shimp & Pfaender, 1987; Wiggins & Alexander, 1988; Tibbles
and Baecker, 1989a). Several studies have shown that phenol can be degraded by a
variety of microorganisms such as bacteria, fungi, yeast and algae as previously shown in
Table 2.3. Many microbial strains have been found to be phenol-degrading under
mesophilic condition (Allsop et al., 1993).
In the last 20 years, studies have been performed on both aerobic (Oltmanns et al.,
1989) and anaerobic (Knoll and Winter, 1987) treatment of aromatic pollutants by using
pure microorganisms or pure culture. Aerobic processes of biological treatment are
generally preferred to degrade phenolic compounds (Fedorak et al., 1984) due to the low
costs associated with this option, and as well as to the possibility of their complete
mineralization (Collins and Daugulis, 1997b). Studies on phenol toxicity to bacteria in
phenol-contaminated sites have shown that bacteria can adapt to ambient phenol
concentrations, but increasing phenol concentrations appear to decrease the overall
biodegradation (Dean-Ross, 1989; Dean-Ross and Rahimi, 1995).
2.6.3.1 Phenol inhibitory levels for phenol degradation by microorganisms
Substrate inhibition is characteristic of toxic substrate metabolism (Santos and
Linardi, 2004). The toxicity of phenol at high concentrations level could inhibit the
29
related metabolism of degradation resulting in a lower efficiency by free cells (Chen et
al., 2002). The observed phenol inhibitory level reported by previous researchers is
shown in Table 2.5.
Table 2.5 : Phenol inhibitory levels for phenol degradation by microorganism
Microorganism
Observed phenolinhibitory level (mg L-1)
Reference
C. tropicalis, Trichosporoncutaneum and Dabaromycessubglobosus (mixed culture)
300
Chai et al., 2004
NA
300
Yoong et al., 2004
Ralstonia eutropha 335
ATCC 17697
282
Léonard et al., 1999
P. putida Q5
<25-120
Onsyko et al., 2002
Comamonas testosteroni P15
117
Yap et al., 1999
Comamonas testosteroni E23
235
Yap et al., 1999
P. putida CCRC 14365
80
Chung et al., 2003
P. putida ATCC 700007
50
Abuhamed et al., 2003
P. putida ATCC 49451
50
Wang and Loh, 1999
Halophilic bacteria CA00
50
Peyton et al., 2002
P. putida NRRL-ß-14875
40
Seker et al., 1997
Achromobacter sp. E1
40
Watanabe et al., 1996a
Alcaligenes sp. E2
30
Watanabe et al., 1996a
Alcaligenes R5
30
Watanabe et al., 1996a
P. putida DSM 548
25
Monteiro et al., 2000
P. putida MTCC 1194
25
Mahadevaswamy et al.,
2004
30
2.6.3.2 Phenol degradation lag period (TL)
It been reported that before a microbial cells can commence active metabolism of
a substrate, they have to adjust to their surrounding environment (Bailey and Ollis, 1986).
TL was always observed in the course of phenol degradation irrespective of difference of
microbial strain and conditions (free, immobilized or engineered) and is likely a function
of other variables such as pH, temperature and phenol concentration as a substrate,
inoculum size, electron acceptors, adaptation of bioparticles, and physiological state of
cells (Tibbles and Baecker, 1989a; Tarighian et al., 2001; Baek et al., 2001; González et
al., 2001a; Prieto et al., 2002).
The TL observed during the course of phenol degradation previously reported is
shown in Table 2.6. All data presented were from biodegradation studies when phenol
was used as sole carbon source.
31
Table 2.6: Observed phenol biodegradation lag period (TL)
TL (h)
1.5-2
IPC(mg L-1)
13-40
Microorganism
P. putida MTCC 1194
(immobilized cells)
Reference
Mahadevaswamy et al., 2004
2
500
T. cutaneum R57
Alexieva et al., 2004
2
500
Arthrobacter sp.
Kar et al., 1996
3.7
500
P. putida wild strain
Soda et al., 1998
2.9
500
P. putida BH (GEM)
Soda et al., 1998
Hao et al., 2002
2-3
200-300
Acinetobacter sp.
2-5
85-450
P. putida BH CCRC143659
P. putida F1 ATCC –
700007
3-17
3.5-8
50-600
Chung et al., 2003
Abuhamed et al., 2003,
2004
10-100
P. putida DSM 548
Monteiro et al., 2000
4
650
Ralstonia eutropha
Léonard et al., 1999
5
94
Bacillus sp.A2
Mutzel et al., 1996
5
249-1000
P. putida MTCC 1194
(immobilized)
Bandhyopadhyay et al.,
2001
6-40
50-320
Halophilic bacteria
Peyton et al., 2002
5-20
100-800
P. putida ATCC 49451
Wang and Loh, 1999
Nocardia sp.
Tibbles and Baecker, 1989a
P. putida ATCC 17484
Tarighian et al., 2001
10
10.5
0.29-0.40
300
15
0.29-0.40
Acinetobacter sp.
Tibbles and Baecker, 1989a
15
0.29-0.40
Arthrobacter sp.
Tibbles and Baecker, 1989a
15
200
P. putida Q5
Kotturi et al., 1991
25
1200
P. putida ATCC 11172
Loh and Liu, 2001
26
0.29-0.40
Micrococcus sp.
Tibbles and Baecker, 1989a
32
Table 2.6: Observed phenol degradation lag period (TL) TL(h)
4-35
4-20
8-20
IPC(mg L-1)
400-1000
400-1000
200
Microorganism
R. erythropolis UPV-1
(free cells)
R. erythropolis UPV-1
(immobilized cells)
continue
Reference
Prieto et al., 2002
Prieto et al., 2002
R. erythropolis UPV-1
(at different physiological
state of cells used)
Prieto et al., 2002
8-10
500
Acinetobacter sp. W17
Abd-El-Haleem et al., 2002
20
500
Alcaligenes strain P5
(different electronacceptor)
Baek et al., 2001
24
500
70
500
Acinetobacter sp. W-17
(immobilized cells)
Acinetobacter sp. W-17
(free cells)
Beshay et al., 2002
Beshay et al., 2002
70
500
Acinetobacter sp. W-17
(free cells)
60
1000
P. putida ATCC 17484
(from two adaptation)
González et al., 2001a
P. putida ATCC 17484
(from one adaptation)
González et al., 2001a
C. tropicalis NCYC 1503
(immobilized cells)
Chen et al., 2002
C. tropicalis NCYC 1503
(free cells)
Chen et al., 2002
P. putida EKII
Hinteregger et al., 1992
70
3-90
3-130
1->140
1000
200-1500
200-1500
200-1200
Abd-El-Haleem et al., 2003
33
2.6.3.3 Intermediates of phenol biodegradation and metabolic pathway
Phenol is converted by bacteria under aerobic conditions to carbon dioxide
(Southworth et al., 1985; Ursin, 1985; Aelion et al., 1987; Aquino et al., 1988), and
under anaerobic conditions to carbon dioxide (Bak & Widdell, 1986; Tschech & Fuchs,
1987) or methane (Healy & Young, 1979; Ehrlich et al., 1982; Young & Rivera, 1985;
Fedorak & Hrudey, 1986; Fedorak et al., 1986). The intermediates in the biodegradation
of phenol are benzoate, catechol, cis,cis-muconate, ȕ-ketoadipate, succinate and acetate
(Paris et al., 1982; Krug et al., 1985; Fedorak et al., 1986; Knoll & Winter, 1987).
Phenol degradation by microbial pure and mixed cultures have been actively studied
(Ahmed, 1995; Collins and Daugulis, 1997a; Schroder et al., 1997; Chang et al., 1998;
Ruiz-Ordaz et al., 1998). Most of the cultures tested are capable of degrading phenol at
low concentrations (Chang et al., 1998). Most studies on phenol degradation have been
carried out with bacteria mainly from the Pseudomonas genus (Ehrhardt and Rehm,
1989; Fava et al., 1995; Ahmed, 1995).
Phenol may be degraded in its free form as well as after adsorption onto soil or
sediment, although the presence of sorbent reduces the rate of biodegradation (Shimp &
Young, 1987; Knezovich et al., 1988). When phenol is the only carbon source, it can be
degraded in a biofilm with first-order kinetics at concentrations below 20 ug L-1 at 10oC.
The first-order rate constants are 3 to 30 times higher than those of easily degraded
organic compounds at 100-1000 fold higher concentrations (Arvin et al., 1991). Howard
(1989) reported that phenol degradation rates suggest rapid aerobic degradation in
sewage (typically > 90% with an 8 h retention time), soil (typically complete
biodegradation in 2-5 days), fresh water (typically biodegradation in < 1 day), and sea
water (typically 50% in 9 days). Anaerobic biodegradation is slower (Baker & Mayfield,
1980).
In bacteria, aromatic compounds are converted to few substrates: catechol,
protocatechuate and more rarely gentisate (Löcher, 1991). Representative aromatic
compounds that are converted via catechol are shown in Figure 2.2.
34
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me ta
Catechol
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Figure 2.2 Microbial metabolism of some aromatic compounds via catechol (Adapted
from Löcher, 1991).
As mentioned earlier, bacteria play a major role in the degradation of phenol in
soil, sediment and water. The number of bacteria capable of utilizing phenol is only a
small percentage of the total population present in, for example, a soil sample (Hickman
& Novak, 1989). However, a repeated exposure to phenol may result in acclimation as
suggested by a number of researchers (Young & Rivera, 1985; Colvin & Rozich, 1986;
Shimp & Pfaender, 1987; Wiggins & Alexander, 1988; Tibbles & Baecker, 1989a).
Phenol may be degraded in its free form as well as after adsorption onto soil or sediment,
although the presence of sorbent reduces the rate of biodegradation (Shimp & Young;
1987; Knezovich et al., 1988).
Phenol may be converted by bacteria under aerobic conditions to carbon dioxide
(Aquino et al., 1988) and under anaerobic conditions to carbon dioxide (Tschech &
35
Fuchs, 1987) or methane (Fedorak et al., 1986). The aerobic and anaerobic degradation
of phenol has been studied extensively using various microorganisms (Bak and Widdel,
1986; Karlsson et al., 1999; Ruiz-Ordaz et al., 2001; Mendonça et al., 2004; Yan et al.,
2005).
Under aerobic condition, oxygen is used as electron acceptor for the transfer of
electrons.The transfer of electrons between the electron-donor and electron-acceptor,
substrates is essential for creating and maintaining biomass. For instance, in the
biodegradation of phenol, phenol is the primary substrate and must be made available in
order to have biomass active in the biodegradation process. According to Rittmann and
Sàez (2003) once active biomass is present, any biotransformation reaction can occur,
provided the microorganisms possess enzymes for catalyzing the reaction. These
enzymes that are involved in the aerobic metabolism of aromatic compounds usually
define the range of substrates that can be transformed by certain metabolic pathway
(Pieper and Reineke, 2000).
The first step in aerobic metabolism is phenol hydroxylation to catechol by phenol
hydroxylase (EC 1.14.13.7) a NADPH-dependant flavoprotein (Neujahr and Gaal, 1973;
Enroth et al., 1998). It incorporates one oxygen atom of molecular oxygen into the
aromatic ring to form catechol. Phenol hydroxylases, strictly dependent on the presence
of NADPH, have been described in extracts of T. cutaenum (Neujahr and Gaal, 1973) and
C. tropicalis (Neujahr et al., 1974). The second step is catalyzed by catechol 1,2dioxygenase (EC 1.13.11.1; ortho fission) or catechol 2,3-dioxygenase ( EC 1.13.11.2;
meta fission). After several subsequent steps, the products are incorporated into the
tricarboxylic acid cycle (TCA) or Krebs cycle (Shingler, 1996). It has been established
that the aerobic degradation of phenolic compounds is metabolized by different strains
through either the ortho-or the meta-cleavage pathway (Bayly and Barbour, 1984;
Ahamad and Kunhi, 1996; Shingler, 1996).
36
In absence of O2
phenol hydroxlase
reaction stop. No
formation of catechol
OH
PHENOL
O2 + NADPH + H+
Phenol hydroxylase
NADP + H2O
OH
OH
o-fission
catechol 1,2-dioxygenase
Catechol
O2
COOCOOcis,cis-muconic acid
O
COO-
2H+
m-fission
catechol 2,3-dioxygenase
O2
2H+
In absence of O2,
C1,2D or C2,3D
reaction stop. No
formation of ccMA or
2-HMSA
OH
COOCHO
2-hydroxymuconic semialdehyde
COOß-ketoadipate
COOH
Acetyl˜CoA + succinate
CO2
acetaldehyde + pyruvate
Figure 2.3 The main pathways of phenol degradation under aerobic condition (ortho- and
meta fission of the benzene ring) (adapted from Krug et al., 1985) {Reaction: Phenol +
O2 + NADPH +H+
catechol + O2
NADP+ + H2O + catechol (Mörtberg and Neujahr, 1987),
ccMA + 2H+ (Ngai et al., 1990).
A number of researchers (Shindo et al., 1995; Collins and Daugulis, 1997b; Fan et
al., 1987; Livingstone and Chase, 1990; Yang and See, 1991) suggested that there are
many possible biotechnological applications of aromatic-degrading organisms and their
constituent enzymes have been investigated including the use in bioreactor systems for
removal of toxic waste products or treatment of contaminated wastes. Other applications
include the production of valuable biotransformation products such as picolinic acids
37
from catechol (Asano et al., 1994), cis,cis-muconic acids from benzoic acid, benzene,
toluene or catechol (Yoshikawa et al., 1990; Bang and Choi, 1995; Choi et al., 1997) and
also as a reporter gene in diagnostic systems, for example, catechol 2,3-dioxygenase gene
as suggested by Shindo et al., (1995).
Intermediates and products produced in aerobic degradation of phenol by
microorganisms are listed in Table 2.7.
Table 2.7: Intermediates and products of phenol degradation by microorganism
Microorganism
A. Bacteria
Intermediates
Products
AlcaligeneseutrophusJMP 134
catechol
cis,cis-muconate
2-HMSA
ß-ketoadipate,
succinate
formate,
Acetyl CoA
Müller and Babel, 1996
catechol
2-HMSA
Acetyl CoA
pyruvate
Ali et al., 1998
catechol
2-HMSA
Acetyl CoA
pyruvate
Bacillus sp.
P. putida
Reference
Mörsen & Rehmn, 1990
B. Fungi
A. fumigatus
(ATCC 28282)
C. Yeast
C. tropicalis
catechol,
hydroquinone
catechol
cis,cis-muconate
3-oxoadipate,
1,2,4-trihydroxybenzene,
maleylacetate
Jones et al., 1995
ȕ-ketoadipate,
succinate,
Acetyl CoA
Krug et al., 1985
Páca Jr. 2003
E. algae
Ochromonasdanica
catechol
2-HMSA
Acetyl CoA
Pyruvate
Semple and Cain, 1995
38
The cleavages of phenol by different microorganisms are listed in Table 2.8.
Table 2.8: Phenol metabolism pathway of microorganism
Microorganism
A. Bacteria
Pathway
Reference
Acinetobacter calcoaceticus
A. radioresistens
ortho
ortho
Paller et al., 1995
Pessione et al., 1999
Pessione & Giunta, 1997
Alcaligenes eutrophus
A. eutrophus JMP 134
A. faecalis
meta
meta & ortho
ortho
Leonard & Lindley, 1998
Müller & Babel, 1996
Bastos et al., 2000b
Bacillus strain Cro3.2
B. stearothermophilus
B. thermoleovorans A2
B. thermoglucosidasius A7
Comamonas testosteroni
meta
meta & ortho
meta
meta
meta
Ali et al., 1998
Adams & Robinson, 1988
Milo et al., 1999
Duffner et al., 2000.
Yap et al., 1999
Ochrobatrum tritici
Pseudomonas sp.
P. sp. CF600
P. cepacia AC1100
meta
meta & ortho
meta
meta
meta
P. pickettii
P. putida
meta
meta
P. putida BH
P. putida NCIB 10015
meta
meta
P. putida P35X
P. vesicularis
meta
meta
Ralstonia
R. eutropha
Staphylococcus sciuri
meta & ortho
meta
ortho
El-Sayed et al., 2003
de Liphay et al., 1999
Kang & Park, 1997.
Powlowski & Shingler, 1994
Ghadi & Sangodkar,1994
Nelson et al., 1987
Kukor & Olsen, 1991
Hill & Robinson, 1975
Yang & Humphrey, 1975
Morsen & Rehm, 1990
Takeo et al., 1995
Dagley & Gibson, 1965
Bayly & Dagley, 1969
Sala-Trepat et al., 1972
Ng et al., 1994.
Mrozik and àabuĪek, 2002
de Lipthay et al., 1999
Leonard et al., 1999
Mrozik and àabuĪek, 2002
39
Table 2.8: Phenol metabolism pathway of microorganism -
continue
Microorganism
B. Fungi
Pathway
Reference
Aspergillus fumigatus
ATCC 28282
A. (LA2, LA3,AE5)
ortho & para
Jones et al., 1995
ortho
Santos and Linardi, 2004
Penicillium (AF2, AF4, FIB9)
ortho
Santos and Linardi, 2004
Graphium (LE9, LA1,LA5, (LE11, FIB4, LE6, AE2)
ortho
Santos and Linardi, 2004
Scedosporium apiospermum
ortho
Clauȕen and Schmidt, 1998
Aureobasidium
ortho
Santos and Linardi, 2001
Candida tropicalis
ortho
Bastos et al., 2000a
Rhodotorula
R. glutinis
ortho
ortho
R. rubra
ortho
Trichosporon sp. LE3
ortho
Santos and Linardi, 2001
Katayama-Hirayama et al.,
1994
Katayama-Hirayama et al.,
1991
Santos and Linardi, 2001
Fusarium (FE16, FE11)
ortho
Santos and Linardi, 2004
meta
Semple and Cain, 1995,
Semple, 1997
C. Yeast
D. Algae
Ochromonas danica
As shown in Table 2.8, most bacteria and algae degrade phenol via meta pathway
with exception of A. euthrophus JMP 134 (Müller and Babel, 1996), B.
stearothermophilus (Adams & Robinson, 1988), Pseudomonas sp. and Ralstonia sp. (de
Liphay et al., 1999) found to degrade phenol via both meta and ortho pathway. Fungi and
yeast degrade phenol via ortho-cleavage pathway except A. fumigatus (Jones et al., 1995)
40
found to degrade phenol via ortho and para pathway. Some bacterial strains have the
capability to degrade substances with multiple pathways (Haigler et al., 1990).
2.6.4 Anaerobic biodegradation of phenol
Besides the fact that phenols are employed in industrial processes, phenol
compounds are commonly found in nature since they are also produced during
decomposition of organic materials, and produced by plants. Plants are known to play an
important role in the defend response of plants to various stress factors such as pathogen
and insects (Heller et al., 1990; Nicholson and Hammerschmidt, 1992). Increased in
environmental levels may also result from forest fire (Hubble et al., 1981).
Regardless of its origin (natural or anthropogenic) or due to the compounds that
may produced phenol during degradation some will enter anaerobic environment, such as
sediments and landfill. Phenolic compounds are among the common contaminants in
landfill leachate (Sawhney and Kozloski, 1984; Lesage et al., 1990; Christenssen et al.,
1994). Landfills were shown to be a habitat of anaerobic microbial populations capable of
degrading toluene, phenol and p-cresol as reported by Wang and Barlaz, (1998). In
landfills, anaerobic conditions are developed during refuse decomposition and CO2 is the
major electron sink (Barlaz, 1996). Therefore methanogenic processes control
biodegradation in the landfill ecosystem.
Anaerobic growth on phenol has been observed for various bacteria (Bak and
Widdel, 1986; Tschech and Fuchs, 1987, 1989; Sharak-Genthner et al., 1991; Gallert and
Winter, 1992; Zhang and Wiegel, 1992; Li et al., 1996; van Schie and Young, 1998;
Shinoda et al., 2000). In all cases studied, phenol appeared to be carboxylated to 4hydroxybenzoate and growth on phenol was dependent on the presence of CO2 (Tschech
and Fuchs, 1987). Consortia of fermenting bacteria convert phenol to benzoate and
decarboxylate 4-hydroxybenzoate to phenol (Gallert and Winter, 1992; Zhang and
Wiegel, 1992).
41
Benzoate is a key intermediate for the degradation of many aromatic compounds,
including phenol and chlorophenol (Knoll and Winter, 1989; Kobayashi et al., 1989).
Although degradation of benzoate is a multi-step pathway, it is directly converted to
acetate and hydrogen by bacteria such as Syntrophus buswellii (Tarvin and Bushwell,
1934; Mountfort et al., 1984). Many of the phenolic compounds subjected to anaerobic
degradation give rise to unsubstituted phenol as an intermediate (Young and Rivera,
1985; Zhang et al., 1989; Londry and Fedorak, 1991).
Phenol carboxylation to 4-hydroxybenzoate is a paradigm for a new type of
biological carboxylation reaction (Breinig et al., 2000). In all cases of anaerobic growth
by bacteria, phenol appeared to be carboxylated to 4-hydroxybenzoate and growth on
phenol was dependent on the presence of CO2 (Tschech and Fuchs, 1987; van Schie and
Young, 1998).
Denitrifying aromatic compound-degrading bacteria that have been isolated so far
are Thauera (Anders et al., 1995), Azoarcus (Zhou et al., 1995) species which are
members of the ß subclass of the class Proteobacteria (Heider and Fuchs, 1997), and
Magnetospirillum sp., a member of the Į subclass of the class Proteobacteria (Shinoda et
al., 2000). According to Breinig et al., (2000) phenol carboxylation to 4-hydroxybenzoate
is a paradigm for a new type of biological carboxylation reaction. The process has been
studied in the denitrifying bacterium Thauera aromatica (Lack and Fuchs, 1992, 1994;
Aresta et al., 1998; Breinig et al., 2000) (Figure 2.4).
Phenol carboxylation proceeds in two steps and involved in the formation of
phenylphosphate as the first intermediate (Phenol + X˜˜ P Æ phenylphosphate + X) (Lack
and Fuchs, 1992, 1994). Phenylphosphate is pospulated to be the substrate of second
enzyme, E2 (phenylphosphate carboxylase) (Lack and Fuchs, 1992). It requires Mn2+ and
catalyzes the carboxylation of phenylphosphate to 4-hydroxybenzoate (E2-phenolate +
CO2 ÅÆ E2 + 4-hydroxybenzoate.
42
E2, phenylphosphate carboxylase is
strongly inhibited by O2
PHENOL
X
CO2
Pi
OH
COO-
_
X-P
E1 -P
4-hydroxybenzoate
O
OE2
O=P-OE1
X-P, phosphoryl donor,
unknown so far
OH
Phenylphosphate
SCoA
O
Fd
C
Benzoyl coenzyme
A (CoA)
HSCoA
ATP
E3
AMP+PPi
SCoA
O
OH
E2
red
Fd
C
E5
2 ADP
+ 2 Pi
SCoA
O
red
C
E4
2 ATP
2 H2O
2 H-
H2O
+
2H
OH
4-hydroxybenzoylCoA
3 Acetyl- CoA + CO2
Figure 2.4 Postulated pathway of anaerobic phenol degradation in the denitrifying
bacterium T. aromatica. E1, phenylphosphate synthase; E2, phenylphosphate
carboxylase (the mechanism by which the phenolate anion is bound to the enzyme E2 is
unknown so far); E3, 4-hydroxybenzoate-CoA ligase; E4, 4-hydroxybenzoyl-CoA
reductase (dehydroxylating, ferredoxin dependent); E5, benzoyl-CoA reductase
(dearomatizing, ferredoxin dependent). Fd, ferredoxin. X-P, phosphoryl donor, unknown
so far (adapted from Breinig et al., 2000).
According to Schühle and Fuchs, (2004) phenylphosphate carboxylase is a
member of a new family of carboxylases /decarboxylases that act on phenolic compounds
using CO2 as substrate, do not contain biotin or thiamine diphosphate, require K+ and
divalent metal cation (Mg 2+ or Mn 2+) for activity, and are strongly inhibited by oxygen
(Figure 4.4).
43
The most studied pathway of phenol transformation under methanogenic
conditions is via the formation of benzyl-CoA. Gallert and Winter (1992, 1994) presented
evidence for the presence of enzymes performing carboxylation, decarboxylation and
dehydroxylation reactions during phenol transformation. These authors suggested that
phenol degrades to benzoate via 4-hydroxybenzoate, 4-hydroxybenzoyl-CoA and
benzoyl-CoA. The first and successful isolation of a bacterium degrading phenol to
benzoate and also 4-hydroxybenzoate to phenol and benzoate when growing on proteose
peptone without any external electron acceptor CO2 was reported by Li et al., (1996). In
another study, Knoll and Winter (1989) reported that phenol transformation to benzoate
took place in a mixed methanogenic culture fed with phenol as the only energy and
carbon source.
The influence of the methanogenic inhibitor bromoethane sulfonic acid (BES) and
a H2/CO2 atmosphere on anaerobic has been investigated by several researchers. Béchard
et al., (1990) reported on a methanogenic culture forming benzoate and acetate as
intermediary product during phenol degradation. The rate of phenol transformation in
their culture was found to be unaffected by the presence of BES. The study by Karlsson
et al., (1999) reported that during the degradation of 10 mM phenol in the presence of
BES under mesophilic conditions, part of the phenol was reductively transformed to
benzoate while the rest is oxidized to forming acetate as end product (Figure 2.5).
44
[2 CoA]
2 benzoate
3 CO2
3 phenol
3 4-OH-benzoate
3 4-OH-benzylCoA
3 benzoylCoA
6[H]
[1 CoA]
1 acetate
6 CO2
2 acetate
2 ATP+2 CoA
3 acetylCoA
2 ADP + 2P1
Figure 2.5 Phenol degradation pathway, phenol transformation to benzoate and acetate in
the presence of BES. The phenol is initially reduced, forming benzoyl-CoA from 4hydroxybenzoyl-CoA. In the reduction step six [H] are consumed. One third of the 4hydroxybenzoyl-CoA is then oxidized to acetate during the formation of six [H]. The
remaining 4-hydroxybenzoyl-CoA is converted to benzoate (Karlsson et al., 1999).
Fang et al., (1996) studied the feasibility of anaerobic treatment of wastewater
containing phenol as the sole substrate. They found out that wastewater containing 1260
mg L-1 of phenol could be treated anaerobically, however, the process required a lengthy
start-up and was easily disturbed by changes of temperature and phenol concentration. It
has been reported that among the high-rate processes developed, the up-flow anaerobic
sludge blanket (UASB) reactor is probably most successful commercially (Lettinga et al.,
1980; Lettinga and Hulshoff Pol, 1991; Fang and Chui, 1993).
45
2.7
Phenol biodegradation methods
Current technology for the biodegradation of toxic compounds involves the use of
microorganisms in batch and continuous processes, using either suspended or
immobilized cultures. In all of these instances, phenol toxicity is still a concern (Collins
and Daugulis, 1997a). The toxicity exerted by high concentrations causing loss of
cytoplasmic membrane integrity (Heipieper et al., 1991, 1992; Keweloh et al., 1990).
This finally resulted in disruption of energy transduction, disturbance of membrane
barrier function, inhibition of membrane protein function, and subsequent cell death (Yap
et al., 1999).
2.7.1
Batch fermentation
2.7.1.1 Definition
Batch fermentation is an example of a closed system, containing an initial limited
amount of nutrient (Standbury and Whitaker, 1984). Cultivation begins at the initial
limiting substrate concentration, So, and inoculum size, Xo, The biomass reaches its
maximum, Xmax, when the limiting substrate is depleted, S = 0 and then declines even in
the absence of exogenous elimination (Panikov, 1995). At the time t = 0, the sterile
nutrient solution in the fermenter is inoculated with microorganisms and incubation is
allowed to proceed under optimal physiological conditions. Control systems for batch
fermentation are normally associated with pH, dissolved oxygen tension and temperature.
The growth of cell consisted of number of phases namely, lag phase, exponential
growth phase and a stationary phase. After inoculation, there is a period of where no
growth took place and this phase is referred to as the lag phase and considered as a time
for adaptation (Scragg, 1992).
In batch fermentation, if growth is subjected to substrate inhibition, fermentation
has to be started with low initial substrate concentration.
46
2.7.1.2 Advantages and disadvantages
Most bioprocesses are based on batch reactors (Shuler and Kargi, 2002). The
principal advantages of batch cultures are; low contamination risk, the ability to run
different successive phase in the same vessel, and close control of the genetic stability of
microorganism (Scragg, 1992; Panikov, 1995). Conventional batch fermentations have
been used to degrade phenol, but are limited by the low initial concentrations required to
prevent complete inhibition of microbial activity (Andrews, 1968).
Batch cultures can suffer great variability from one run to another. Problems such
as substrate inhibition, low cell concentration, glucose effect, catabolite repression,
auxotropic mutants and high viscosity of the culture broth which are common in batch
processes (Shuler and Kargi, 2002). Batch culture under conditions of mineral limitation,
glucose repression may be protracted and even irreversible (Panikov, 1995).
2.7.2
Fed-batch Fermentation
2.7.2.1 Definition
In the broad sense, “fed-batch” is defined as a technique in microbial processes
where one or more nutrients are supplied to the bioreactor during cultivation and in which
the product remains in the containment until the end of the run or process (Yoshida et al.,
1973; Besli et al., 1995). It is may be regarded as a modification of batch operation
(Yamane and Shimizu, 1984). In fed-batch operation, nutrient is fed intermittently or
continuously with a low flow rate and contrary to the continuous system, effluent is not
removed until the maximum liquid fermenter is reached (Bali and Sengül, 2002) and thus
washout does not occur. Unlike a chemostat, the volume of a fed-batch is not constant as
feed is added. Highly concentrated organic and toxic compounds, in this way, are diluted
in a large volume of reactor and therefore inhibitory/toxic effects are reduced. As a result,
biodegradation of these compounds takes place at a higher rate (Kargi, 1996).
47
2.7.2.2 Advantages and disadvantages
Continuous-culture fermentations are unstable and unable to achieve high
removal efficiencies at high inlet phenol concentrations (Pawlowsky et al., 1973).
Immobilized cell reactors can achieve high conversions with better process stability, but
they require high levels of aeration and agitation and can, therefore be costly to operate
(Molin and Nilsson, 1985).
Fed-batch culture is usually used to overcome substrate inhibitory or catabolite
repression by intermittent feeding of the substrate. It the substrate is inhibitory,
intermittent addition of the substrate improves the productivity of the fermentation by
maintaining the substrate concentration low (Shuler and Kargi, 2002). Continuous and
fed-batch culture techniques often provide better yield and productivities in the
production of microbial metabolizes than batch cultures (Kumar et al., 1991). As the
microbial population in fed-batch culture remains diluted all the time, so there are no
negative cell to cell interactions or accumulation of non-dialysable metabolic products,
and the viability of all organisms was about 90% or higher (Panikov, 1995).
However, fed-batch reactor provides a number of advantages over continuous
reactors: 1) Because cells are not removed during fermentation, fed-batch fermenters are
well suited for the production of compounds produced during very slow or zero growth,
2) Unlike a continuous fermenter, the feed does not need to contain all the nutrients
needed to sustain growth. The feed may contain only a nitrogen source or a metabolic
precursor. With chemostat, the feed must contain all the nutrients required to support
growth, or otherwise cell washout will occur, 3) Contamination and /or mutation will also
not have the dramatic effect on fed-batch fermenter as a contaminant will not be able to
completely take over the fermenter (unless the contamination occurred during the early
stages of fermentation), 4) A fed-batch reactor can be operated in a variety of ways. For
example, the reactor can and often must be operated in the following sequence: Batch =>
Fed-batch => Batch. The feed can also be manipulated to maximize product formation.
For example, during fermentation, the feed composition and feed flow rate can be
48
adjusted to match the physiological state of the cells. For practical reasons, therefore,
some continuous operations have been replaced by fed-batch processes (Schügerl, 1987).
The application of a fed-batch approach to treat such effluents provides a possible
strategy comparable to the sequencing batch reactors proposed by Hughes and Cooper
(1996), and avoids direct release into the environment of the treated effluent. Fed-batch
fermentation has been shown to be an effective treatment in the biodegradation of phenol
using various strategy; control feed strategy using Ralstonia eutropha (Léonard et al.,
1999), two-phase partitioning bioreactors (TPPB) using Pseudomonas putida (Collins
and Daugulis, 1996, 1997a,b), Fed-batch system with oxistat control using Candida
tropicalis Ct2 (Komarkova et al., 2003). Bali and Sengül (2002) found that fed-batch
operation is a promising method for the treatment of high strength and/toxic wastewaters.
One disadvantage of fed-batch, it requires previous analysis of the
microorganisms, its requirements and the understanding of its physiology with the
productivity. Another drawback of fed-batch fermentation is that the high growth rate and
cell density are achieved at the expense of addition of pure O2 to degrade salicylate using
Pseudomonas cepacia with on-line control system (Tocaj et al., 1993).
2.8
Literature Review Summary
Phenol is an aromatic hydrocarbon commonly found in many industrial effluents
including those from oil refinery, petrochemical, coal coking and coal gasification
industries. Their concentration in these effluents ranged from 0.01 mg L-1–10,000 mg L-1.
Phenol has been also detected in ground water due to leaching through soil from spill or
landfill.
As an aromatic hydrocarbon, it is not readily biodegradable compared to the
simple and branched alkanes. Phenol has been known to be toxic and suspected
carcinogens. Its presence in the environment posed a health hazards to human, animals
49
and microorganisms. Therefore, its release into the environment is regulated by
government authority in many countries. Due to its unhealthy effects on human, there has
been a strong interest on their degradation using microorganisms isolated from soils,
water and sediments.
Despite of its toxicity, eukaryotes and prokaryotes could degrade phenol by using
it as an energy and carbon sources either in mixed or as a single substrate. Bacteria and
yeast play a major role in biodegradation of phenol in soil, sediments and water. Most
studies on phenol degradation have been carried out using bacteria, mainly from
Pseudomonas genus. Among the yeast strains, Candida tropicalis have been found able
to degrade phenol and its derivatives and aliphatic compounds.
Biodegradation has been found as a possible alternative to physical and chemical
methods for treatment of phenol because of its relatively low in cost and most
importantly a complete mineralization of phenol could be achieved. Biodegradation of
phenol can be carried out either under aerobic or anaerobic conditions. Aerobic
degradation is much preferred to anaerobic degradation due to the most important fact
that aerobic degradation could lead to complete mineralization producing water and CO2,
which are environmentally acceptable and less expensive.
Most studies on phenol biodegradation under aerobic condition have been carried
either in mixed culture or as sole carbon source out using batch or fed-batch fermentation
techniques. Both techniques have their own advantages and disadvantages. However,
phenol biodegradation under aerobic condition is still much preferred as compared to
anaerobic degradation.
In anaerobic growth by bacteria, phenol appeared to be carboxylated to 4hydroxybenzoate and growth is dependent on the presence of CO2. This carboxylation
would produced phenylphosphate as the first intermediate catalyzed by phenylphosphate
synthase. This first intermediate is then converted to 4-hydroxybenzoate catalyzed by
phenylphosphate carboxylase. The end products of phenol degradation in prokaryote cells
50
were acetyl-CoA and CO2.
The phenol degradation under aerobic condition in eukaryotes such as Candida
tropicalis is catalyzed mainly by two key enzymes namely phenol hydroxylase (PH) and
catechol 1,2-dioxygenase (C1,2D) via the ortho ring cleavage pathway (also known as ßketoadipate pathway) of catechol to cis,cis-muconic acid (ccMA). The end products of
phenol degradation in eukaryotic cells were succinic acid and acetyl CoA. On the other
hand, phenol degradation in prokaryotes such as Pseudomonas sp. is catalyzed by phenol
hydroxylase (PH) and catechol 2,3-dioxygenase (C2,3D) of catechol to 2hydroxymuconic semialdehyde (2-HMSA) via the meta ring cleavage pathway. The
activity of these enzymes could be affected by temperature, pH and concentration of
initial phenol concentration. The end products of phenol degradation in prokaryote cells
were pyruvate and acetyl CoA. The end products from both pathways are subsequently
incorporated into the TCA or Krebs cycle.
51
CHAPTER 3
GENERAL MATERIALS AND METHODS
3.1
Media preparation
3.1.1
Ramsay medium (RM) agar
For the isolation of heterotrophic microorganisms in the environmental samples,
Ramsay medium (RM) agar described by Ramsay et al., (1983) was used for the isolation
of microbes from the petrochemical wastes. The composition of RM is shown in Table
3.1. The medium was prepared by resuspending all ingredients except MgSO4.7H2O and
glucose in 1000 mL of distilled water and then autoclaved for 15 minutes at 121oC at 15
pounds per sq. in (psi) steam pressure. Filter sterilized MgSO4.7H2O and glucose were
added after autoclaving. The initial pH value of the medium was adjusted between 6.5 –
6.8.
Approximately 20 mL of autoclaved media was poured into pre-sterilized
disposable Petri dishes. The media was left to cool at room temperature before storing in
an incubator at 30oC for 24 hours to ensure they are free of contamination prior to
analysis.
52
Table 3.1: Composition of Ramsay Mediuma (RM)
Component
Composition (g L-1)
NH4NO3
2.0
KH2PO4
0.5
K2HPO4
1.0
MgSO4.7H2O
0.5
CaCl2.2H2O
0.01
KCl
0.1
Yeast extract
0.06
Glucose
20.0
Agar (2%)
20.0
a
Ramsay et al., (1983)
3.2
Sample collection
Ten petrochemical wastes and oil samples were collected from the Titan
Petrochemical (M) Sdn Bhd. (TPSB), Pasir Gudang, Johor and Exxon Mobil Oil
Refinery (EMOR) Port Dickson, Negeri Sembilan. The samples were transported in
sterile containers and stored at 4oC. The types of samples are shown in Table 3.2 and
sampling locations as shown in Figure 3.1 and 3.2.
53
Table 3.2: Oil and petrochemical wastes samples collected
No.
Sample
Sample Sampling location
Code
Site
1.
Influent of Floatation tank (A)
FTA
S1
TPSB
2.
Aeration basin wastewater
AB
S2
TPSB
3.
Activated sludge
AS
S3
TPSB
4.
Soil from sludge farm
SSF
S4
EMOR
5.
Soil-matrix from sludge farm
SMSF
S5
EMOR
6.
Effluent of biological treatment lagoon
ETL
S6
EMOR
7.
Bioscum from treatment lagoon
BTL
S7
EMOR
8.
Bottom sludge from treatment lagoon
STL
S8
EMOR
9.
Crude oil
CO
-
EMOR
10.
Waxy oil
WO
-
EMOR
NEUTRAL
TANK 1
RAIN WATER
EQUIPMENT OILY DRAIN
STEAM GEN. BLOWDOWN
SPENT CAUSTIC
CAUSTIC PUMP
SEAL WATER
CONDENSATE FILTER EFFLUENT
COOLING TOWER BLOWDOWN
TANKAGE AREA RAINWATER
POLUUTED RAIN WATER
QUENCH EXCHANGER WATER
OILY WATER FROM PE PLANT
COOLING TOWER BLOW DOWN
DEMIN UNIT WASTE BOILER
BLOWDOWN
OIL
SEPARATOR
OIL PIT
COAGULATION
FeCL3
NaOH
H2SO4
OIL
SEPARATOR
S1
AIR
FLOTATION
POLYMER
CHLORINE
CHAMBER
Key: S1-S3 Sampling
Points
S3
SEDIMEN
TATION
TO OPEN
DITCH
Discharge quality,2001
pH = av=7.3; range=5.9-9
BOD=av=23.7; range=5-63
COD=av=114.3; range=51210
OBSERVATION
POND
TO SEA
WATER
(OPEN DITCH)
SLOP OIL
TANK
S2
ACTIVATED
SLUDGE
TREATMENT
H3PO4
Coagulation
Figure 3.1 Wastewater treatment system and sampling points at Titan
Petrochemical Sdn Bhd. (TPSB)Pasir Gudang, Johor
EQUIPMENT
OILY PIT
BUFFER
PIT
NEUTRAL
TANK 1
H2SO4
RAINWATER
BUFFER PIT
54
B.
A.
10%TPH
S4
S5
reused
Load
4-5m
gal/day Biological Treatment Lagoon
•Pond Dimension=1,300’ x 350’
•Retention time= 10 days
Eichorrnia crassipes
S7
Key: S4-S8 =sampling points
sea
4%TPH
after 6
months
Treated
Soil
Disposal
stream
Effluent
•1-2 ppm oil content
S8 •pH=7.6
•Temp=30oC
•BOD=30-40 ppm
S6
•50-80 tons oil sludge/yr
•O:P:N = 100:10:1
•Tillage once in 2 weeks
•No lining provided
•No seeding
•Treatment zone required
(1ft depth x dimension x 10%)
•pH 4 >adjusted to 5-6 with CaCo3
Sludge Farm
Treatment
Figure 3.2 Waste treatment system and sampling points at Exxon Mobil oil
refinery (EMOR) Port Dickson, N.Sembilan
Wastewater from
Refinery Processes
•Decommissioned tank after 10-15 yrs
•Biological treatment lagoon after maintenance
•Crude oil
Oil Sludge
Sources
55
56
3.3
Bacterial culture preservation
Microorganisms require preservation methods in order to ensure optimal viability,
purity and stability for individual strains.
3.3.1 Short-term preservation
The well-defined colonies on the basis of colony morphological characteristics of
all pure bacterial isolates were transferred into RM slant and preserved at 4oC in
refrigerator for future experimental use.
Working pure culture of RAS-Cr1, RETL-Cr1 and RETL-Cr3 were transferred
periodically (every 2 weeks) onto RM agar and preserved at 4oC in refrigerator for ongoing experiments.
3.3.2 Long-term preservation
Pure culture of RAS-Cr1, RETL-Cr1 and RETL-Cr3 were transferred into
cryovial containing cryopreservation beads stock and preserved at -70oC for future
experimental use.
3.4
Phylogenetic analysis of phenol-degrading RETL-Cr1
3.4.1 DNA Extraction
The yeast C. tropicalis RETL-Cr1 was maintained on RM agar. Yeast lysate was
prepared from a 1.5 mL of 24-h culture in RM broth which was previously incubated at
37oC with agitation at 200 rpm. Yeast cells were pelleted by centrifugation at 12,000 rpm
57
for 2 min and resuspended in 293 µL of 50 mM EDTA. To the lysate, 7.5 µL of lyticase
(20mg/mL) was added followed by incubation at 37oC for 60 min.
The extract was then centrifuged at 12,000 rpm for 2 min and resuspended in 300
µL of nucleic lysis solution provided in the Wizard Genomic DNA purification Kit
(Promega Corp., Madison, Wis.) and purification was performed accordingly.
3.4.2
Electrophoresis
PCR products were electrophoresed through a 1.5% agarose in 1X TBE buffer as
the running buffer. The gel was electrophoresed at 4.8 V/cm for 2 hours. A 100-bp DNA
ladder (Promega Corp., Madison, Wis.) was used as marker to estimate the size of DNA
bands. The gels was stained with ethidium bromide-TBE solution for 20 min and then
photographed using UV Transilluminator.
3.4.3
Sequencing and analysis
Amplified DNA were first purified using Wizard Genomic DNA purification Kit
(Promega Corp., Madison, Wis.) before being sent for sequencing to First Base Sdn. Bhd
The sequences obtained were aligned using Clustal W, version 1.82 (Thompson et al.,
1994). The two different sets of nucleotide sequences obtained were checked against
related sequences derived from the GenBank database via the program BLASTN
(Altschul et al., 1990).
58
3.5
Sample Analysis
3.5.1
Determination of biomass concentration
With samples grown in batch culture, sampling was done periodically to
determine the density. Cell density was monitored spectrophotometrically by measuring
the absorbance at 600nm using the Jenway 6300 spectrophotometer, U.K.
The cell dry weight concentration was determined gravitmetrically. Five mL
aliquots were centrifuged for 15 minutes at 15,000 rpm at 10oC in a pre-weighted 30 mL
tubes. The samples were washed twice with distilled water and the pellets were dried at
105oC in an oven overnight, cooled in a dessicator and reweighting until a constant
weight was obtained. The difference between the first (empty) and the second weight was
used to determine the dry weight of biomass as g L-1.
Dry cell weight was then estimated using calibration curve constructed based on
the relationship between optical density at 600 nm and dry weight cell (Appendix A1)
3.5.2 Determination of specific growth rate
In a batch culture, the exponential increase in biomass after inoculation is
measured as a function of time and analyzed to obtained specific growth rate (µ), for that
substrate concentration (Yoong and Edgehill, 1993;Yoong et al., 2004).
The specific growth rate was measured from the slope of the biomass (dry weight)
curve by delineating points between the log growth phase, represented by the
equation below.
µ = (ln Xt - ln Xo)/t
where Xo
= biomass concentration (dry weight) at time 0
59
Xt
= biomass concentration (dry weight) at time t
t
= elapsed time between measurements
3.5.3 Determination of average phenol degradation rate
The average rate of phenol degradation was calculated by dividing the total
amount of phenol consumed with time required for total consumption of phenol (Kar et
al., 1996).
Average phenol degradation rate =
Amount of phenol consumed
Time required for complete consumption of phenol
Time required for total consumption of phenol was calculated by substracting the lag
period (TL) from the total time required for the same.
3.5.4 Determination of glucose
Glucose was concentration was measured using Sigma® kit 510 (Sigma®
Diagnostics, St. Louis, MO) according to manufacturer’s instruction.
Standard Curve was used to calculate residual glucose concentration using
Shimadzu Spectrophotometer Model based on Sigma® procedure 510. Standard curve is
shown in Appendix A2 and procedure for determination of glucose is shown in Appendix
A3.
60
3.5.5 Determination of phenol, catechol and cis,cis muconic acid
Phenol, catechol and cis,cis-muconic acid were determined by isocratic elution
high performance liquid chromatography (HPLC) (W600 2487) using a Waters Hypersil
C18 5µm (4.6 mm x 250 mm) column with UV detector at 280 nm. The mobile phases
used were acetic acid (1% v/v) in water and acetic acid (1% v/v) in acetonitrile, at a flow
rate of 1 mL min -1. Solvents were of HPLC grade. 1 mL sample was centrifuged at
15,000 rpm for 10 minutes in a microfuge (Hettich centrifuge, Germany). The
supernatant was filtered through a 0.25 µm nylon filter to remove cell debris. The filtrate
was cooled and stored at -20oC for subsequent analysis. Aliquots of 40 µL of filtered
samples were injected into the HPLC for phenol determination and analyzed in
duplicates. The HPLC-analytical parameters used in determination of phenol, catechol
and cis,cis-muconic acid is presented in Appendix A4.
61
CHAPTER 4
ISOLATION, SCREENING AND CHARACTERIZATION OF POTENTIAL
PHENOL-DEGRADERS FROM PETROCHEMICAL WASTES
4.1
Introduction
Microorganisms are ubiquitous in nature. A review of literature indicates that the
natural ecosystem harbor microorganisms that are capable of degrading oil in marine,
freshwater and soil ecosystems. In unpolluted ecosystems, hydrocarbon-degrading
organisms represent less than 0.1% of the culturable heterotrophic microbial community,
whereas, in contaminated environments they constitute up to 100% of the viable
microbial population (Atlas, 1981; Ridgeway et al., 1990).
The size of the aerobic hydrocarbon-degrading population in a microbial
community was directly correlated to the extent of hydrocarbon contamination of the
environment (Ridgeway et al., (1990). The involvement of microbes in the remediation
of phenolic compounds produced is widespread (Sutton et al., 1999; Tisler et al., 1999),
however, little is known of the diversity of organisms which fulfill the process. A number
of researchers (Ward et al., 1990; Wagner et al., 1993; Snaidr et al., 1997) have
suggested that microbial diversity in the environment is much greater than that have been
isolated in the laboratory.
62
The microorganisms in natural ecosystem can be exploited for environmental,
industrial and commercial applications. Microbial isolation strategies usually involve a
period of enrichment in liquid culture followed by separation of organisms in or on solid
media where they are allowed to grow as colonies (Hardman et al., 1993). Rapid and
reliable identification and classification of microorganisms are important in
environmental and industrial microbiology. Microorganisms such as bacteria and yeast
are sources of antibiotics, enzymes, and other bioactive compounds for medicine and
biotechnology (Short, 1997; Oh and Kim, 1998; Picataggio et al., 1991).
According to Cornelissen and Sijm (1996) biodegradation of substrate provide
microorganisms with energy and building materials that are used for growth of new cells,
maintenance of cells and catabolism of less degradable substances; energy is also lost in
the form of heat. Cornelissen and Sijm (1996) suggested that in bacterial cellular
processes without co-metabolism, 47-83% of the calculated amount of available energy
was consumed by growth and 20-35% for cell maintenance and heat loss. On the other
hand, with co-metabolism, 7-13% of the actual amount of energy generated are used on
these energy consuming processes; and in some cases the amount of energy consumed by
co-metabolism is equal to or larger than the amount of energy consumed by growth.
Bacteria, yeast and fungi capability of utilizing phenolic compounds are found in
soil and water environments (Kumaran and Paruchuri, 1997). Phenol degraders have been
isolated both from contaminated and uncontaminated environments (Bastos et al., 2002b;
Vojta et al., 2002; Santos and Linardi, 2004). However, this present study is the first
attempt to isolate potential phenol-degraders from wastewater treatment plants of
petrochemical industries. Microorganisms from contaminated environments are cultured
under laboratory conditions for a number of reasons. These include, isolation and
characterization of microorganisms that are able to degrade specific pollutant, for
production of large-scale inocula for bioaugmentation to accelerate remediation of
contaminated environments (Watanabe, 2001). An understanding of microorganisms as
pure cultures or consortia may assist the development of bioremediation technology and
bioremediation monitoring systems (Head, 1998). Attempts to isolate these specific
63
pollutant-degraders and adapt them to biological wastewater treatment processes have
been made by many researchers (Takahashi et al., 1981; Masqué et al., 1987; Hinteregger
et al., 1992).
This chapter describes the isolation, characterization and identification of
microbial strains from wastes of two petrochemical industries in Malaysia that have the
potential to degrade phenol as sole carbon source.
4.2
Materials and Methods
4.2.1 Media preparation
The composition and preparation of Ramsay Medium used is presented in section
3.1.1.
4.2.2 Sample collection
The types of samples and sampling locations are presented in section 3.2.
4.2.3
Isolation of microorganisms
All cultures were isolated via plating after enrichment in batch (shake-serum
bottles). In enrichment method, for solid samples, 50 gram of sample was transferred into
100 mL of the enrichment medium (RM) whereas for liquid samples, 50 mL of samples
were inoculated into pre-sterilized serum bottle containing 100 mL of enrichment
medium. All cultures were incubated on a rotary shaker set at 200 rpm at 37oC for 24
hours.
64
The isolation of the microbial strains was done by spread plate technique as
prescribed in APHA 9215 (APHA, 1989). A serial dilution of the samples were prepared
before inoculating onto RM agar plates. This was to make observation and enumeration
of colonies much easier. Nine dilution bottles filled with 9.0 mL sterilized buffer water
were needed at the start of dilution procedures for each sample. One mL of sample was
transferred into a water blank. Then, 1 mL of the first dilution was transferred into
another dilution and so on until the dilution of 10-9 had been reached. For the inoculation
procedures, dilution 10-3 to 10-5 were chosen. Aliquots (0.1 mL) were spread evenly onto
the RM agar. Duplicates were done for each inoculation.
The effect of temperature and presence of oxygen were determined by incubating
the cultures at 37oC under aerobic and anaerobic conditions for 48h. Anaerobic condition
was obtained by incubating the culture in an anaerobic jar containing gas generating kit
(Oxoid). Anaerobic indicator was used to ensure anaerobic condition throughout
incubation period.
The detailed information and procedures on Heterotrophic Plate Count – Test
Method APHA 9215 are outlined in Appendix 5.
For isolation of a single colony, the well defined isolated colonies were streaked
onto fresh RM agar and incubated at 37oC for 48 h.
4.2.4 Screening for phenol-degrading microorganisms
4.2.4.1 Test for growth on RM agar containing 1 mM phenol
The three selected microbial isolates (RAS-Cr1, RETL-Cr1 and RETL-Cr3) were
tested for growth on RM agar containing 1 mM (94 mg L-1) phenol.
65
Bacterial suspension of each selected isolates were prepared in 10 mL presterilized distilled water by picking 1-2 colonies and resuspended in the distilled water.
Bacterial suspension (0.1 mL) was inoculated onto the RM agar containing 1 mM phenol.
The solid culture was grown in duplicate and incubated at 37oC for 24 hours. Cell count
was enumerated using the colony counter.
4.2.4.2 Test for phenol tolerance of selected isolates
The three selected microbial isolates were tested for phenol tolerance in RM broth
containing varying concentration of phenol. A 1.5 mL inoculum was transferred into a 30
mL universal bottle containing 15 mL of RM containing varying concentration of phenol
ranging from 1 mM to 10 mM. Duplicate cultures were incubated at 30oC with agitation
at 200 rpm for 24 hours. Growth of RETL-Cr1, RAS-Cr1 and RETL-Cr3 was determined
by spread plate technique. Cell count was enumerated by spread plate method by
inoculating 0.1 mL aliquots onto RM agar.
Isolate RETL-Cr1 which was found to be the best phenol degrader was further
tested for growth in RM containing varying concentration of phenol ranging from 1 mM
to 100 mM. A 1.5 mL inoculum was transferred into a 30 mL universal bottle containing
15 mL of RM supplemented with 1 mM glucose. Duplicate cultures were incubated at
30oC with agitation at 200 rpm for 96 hours. Growth of RETL-Cr1 was determined by
spread plate technique. Cell count was enumerated by spread plate method by inoculating
0.1 mL aliquots onto RM agar.
4.2.4.3 Test for survivality
A 10 mL inoculum was transferred into a 250 ml conical flask containing 90 ml
of RM in the absence or presence of 3 mM phenol. In the absence of phenol, 1 mM
glucose was used as control. All cultures were grown in duplicate and incubated at 37oC
66
with agitation at 200 rpm for 18 hours. One mL sample was withdrawn hourly to analyze
for total viable count.
Total viable count was enumerated by spread plate method using 0.1 mL of the
dilution 107 to 109 onto RM agar. The dilution series of 107 to109 were performed in
duplicates. The colony forming units (CFU) (30-300) on each plate were counted using a
colony counter. Concentrations are reported as CFU/mL.
4.2.5
Phenol degradation by selected isolates
A 10 mL inoculum was transferred into a 250 ml conical flask containing 90 mL
of Ramsay Medium supplemented with 3 mM phenol and 1 mM glucose. All cultures
were grown in duplicate and incubated aerobically at 37oC with agitation at 200 rpm for a
period of 18 hours. Three mL sample was withdrawn hourly to analyze for optical density
and residual concentration of phenol and glucose.
4.2.6 Morphological Characterization
4.2.6.1 Colony morphology
Colony morphology characteristics which include size, shape, and colour of the
isolates were examined from 24 hours culture on RM agar.
4.2.6.2 Cellular morphology
Selected isolates (RAS-Cr1, RETL-Cr1 and RETL-Cr3) from overnight culture
were Gram-stained and examined using a bright-field microscope (x1000) (Olympus C35 AD-4) to determine the morphology of the bacterial cells. Fresh cultures were stained
67
with malachite green to test for presence of spores and safranin for the vegetative portion
of cell (Benson, 1980).
The cellular morphology assessment, Gram stain and endospore staining were
done. The procedures for Gram stain and endospore staining can be found in Appendix 6.
4.2.7 Biochemical Tests
Biochemical characterization of the isolates include the test for fermentation
reaction (lactose) and enzyme activity (catalase, citrate, methyl red, oxidase, urease and
Vogues Proskauer) were carried out as described by MacFaddin, (1980). All the
biochemical test and basic procedures can be also found in Appendix 6.
4.2.8
Identification of selected isolates
4.2.8.1 Phylogenetic analysis of phenol-degrading RETL-Cr1
From the five isolates selected for their phenol-degrading capabilities, RETL-Cr1
was chosen for further characterization. Isolate RETL-Cr1 was identified by PCR
amplification of ribosomal-DNA using ITS1 and ITS4 as forward and reverse primers,
respectively. ITS1 and ITS4 are universal fungal-specific primers (White et al., 1990;
Park et al., 2000; Fujita et al., 2001).
(i)
PCR Procedure
DNA was extracted before PCR amplification. The DNA extraction procedures
are presented in section 3.4.1.
68
PCR amplification was done using the primer pairs of ITS1 (5’TCCGTAGGTGAACCTGCG-3’) and ITS4 (5’-TCCTCCGCTTATTGATATGC-3’) as
described by White et al., (1990). ITS1 and ITS4 primers were designed from a
conserved motif regions of 18S and 28S ribosomal DNA. The ITS1 - ITS4 primer pair
was used to amplify the intervening 5.8S rDNA and the adjacent ITS1 and ITS2 regions
(Fujita et al., 2001) (Figure 4.1).
ITS1 primer
18S rDNA
ITS3 primer
ITS1
5.8S rDNA
ITS2
28S rDNA
ITS4 primer
Figure 4.1 Schematic representation of the fungal ribosomal genes
containing the primer target areas used in this study (Fujita et al., 2001).
PCR amplification was performed according to the method of Fujita et al., (2001).
Four microlitres of sample was added to the PCR master mix, which consist of 10 µL of
10X PCR buffer, 8 µL of a deoxynucleoside triphosphate mixture (0.1 mM each dNTP),
1.6 µL of each primer (40 pmol of each primer ( ITS1, ITS4), and 0.8 µL (2.0 U) of Taq
DNA polymerase topped up to100 µL with distilled water. Amplification was performed
in a GeneAmp PCR system 9700 thermal cycle (Perkin-Elmer Corp., Emeryville, Calif.),
under the following PCR condition: an initial denaturation temperature of 94oC for 4 min;
30 cycles of denaturation at 94oC for 30 s, annealing at 55oC for 30 s, and extension at
72oC for 1 min; and followed by a 4-min final extension at 72oC.
(ii)
Electrophoresis, Sequencing and Analysis
The procedures are explained in detail in section 3.4.2 and 3.4.3.
69
The flowchart of the experimental design carried out in this study was
summarized in Figure 4.2 below:
Location 1: TPSB
1. Aeration Basin wastewater
(AB)
2. Activated Sludge (AS)
3. Influent Floatation Tank
Aeration ( FTA)
SAMPLES
Location 2:
EMOR
1. Crude oil (CO)
2. Waxy oil (WO)
3. Effluent Treatment Lagoon (ETL)
4. Soil Sludge Farm (SSF)
5. Soil-oil matrix (SMSF)
6. Bioscum (BTL)
CULTURE
Aerobic & Anaerobic at 37oC
Enrichment Medium
RM at 37oC for 24 hrs
(aerobic and anaerobic) (54 strains)
SCREENING AND CHARACTERIZATION OF
3 SELECTED STRAINS (RM 1mM PHENOL ADDED)
(RAS-Cr1, RETL-Cr1, RETL-Cr3)
PHENOL DEGRADATION STUDY OF
3 SELECTED STRAINS
(RAS-Cr1, RETL-Cr1, RETL-Cr3)
MOLECULAR IDENTIFICATION OF
MICROBE OF INTEREST (RETL-Cr1)
Figure 4.2 Experimental design of isolation, screening and characterization of
phenol-degrading microorganisms from petrochemical wastes.
70
4.2.9
Sample Analysis
4.2.9.1 Determination of Biomass Concentration
The procedures are presented in section 3.5.1
4.2.9.2 Determination of average phenol degradation rate
The procedures are presented in section 3.5.3
4.2.9.3 Determination of Glucose Concentration
The procedures are presented in section 3.5.4
4.2.9.4 Determination of Phenol Concentration
The procedures are presented in section 3.5.5
4.3
Results and Discussion
4.3.1
Isolation and screening for phenol-degrading microorganisms
Microbial isolation and screening is an important stage for evaluating the
potential biodegraders for different pollutants in the environment that can be used in
environmental biotechnology, which relies on the pollutant-degrading capacities of
naturally occurring microorganisms (Liu and Suflita 1993). Phenolic compounds are
widely distributed in the environment both from natural and industrial sources. Several
aerobic microorganisms that degrade phenol have been isolated (Yang and Humphrey,
71
1975; Folsom et al., 1990; Marcos et al., 1997). Bacteria, yeast and fungi capable of
utilizing phenolic compounds are found in contaminated and uncontaminated soil and
water environment (Kumaran and Paruchuri, 1997; Zinjarde and Pant, 2002; Bastos et al.,
2000a; Margesin et al., 2005).
The isolation of potential phenol-degraders microorganism was performed under
aerobic and anaerobic condition. Figure 4.3 shows the number of isolates obtained under
these two conditions.
Number of strains isolated
40
35
35
30
25
19
20
15
10
5
0
Aerobic
Anaerobic
Physiological condition
Figure 4.3 Number of strains isolated from petrochemical samples taken via plating
after enrichment in RM incubated at 37oC.
A total of 54 microbial isolates were obtained from the petrochemical samples via
plating on RM agar after culture enrichment in RM broth incubated at 37oC under aerobic
and anaerobic condition. In this present study, enrichment method was not for selective
isolation of microorganisms but rather for enhancement of cell population growth.
Therefore no phenol was added for the selection. As shown in Figure 4.3 microbial
isolates obtained from the petrochemical samples indicates that more aerobic organisms
can be isolated as compared to anaerobic organisms.
72
In this study, phenol biodegradation under aerobic condition was investigated as it
is still much preferred to anaerobic degradation because of the following possible
reasons. Firstly, the actual degradation pathway, enzymes involved, the participation of
electron carrying proteins and electrochemical gradients during ATP-formation and
reversed electron transport are not well understood and requires extensive studies. This
was insinuated by Karlsson et al., (1999) and to date very little knowledge has been
acquired regarding these aspects of phenol degradation. Even Breinig et al., (2000)
reported that the enzymes involved in anerobic phenol degradation have not been studied
yet, although carboxylation of the aromatic ring is widespread among anaerobic
microorganisms. However degradation pathways under aerobic condition and the key
enzymes involved has been well documented as previously shown in Figure 2.3 (section
2.6.3.3). On the practical aspects, Fang et al., (1996) reported that several problems
could arise in phenol degradation under anaerobic condition such as; it required a lengthy
start-up, meaning that it requires a certain amount of adjustment period before
degradation occurs. He also suggested that phenol degradation under anaerobic condition
can be easily disturbed by the changes of temperature and phenol concentration. To make
situation worse, reasonably high tendency of filter clogging could happen as put forward
by Levin and Gealt, (1993). Hence, aerobic degradation is much preferred to anaerobic
degradation due to the most important fact that aerobic degradation could lead to
complete mineralization producing water and CO2, which are environmentally acceptable
and less expensive.
Of the 35 strains isolated aerobically, 8 strains were selected to be grown in RM
agar containing 1 mM phenol for primary selection on phenol (Table 4.1). Preliminary
studies have shown that these isolates were capable of utilizing various carbon sources
such as crude oil, glucose, diesel, glycerol and kerosene (results not shown). Thus, it is of
our interest to further analyze them for growth on phenol as they have the tendency to
grow on the said carbon sources as sole carbon source. As shown in Table 4.1, isolates
RAS-Cr1, RETL-Cr1 and RETL-Cr3 were the most prominent utilizers of phenol being
able to grow profusely on RM agar containing 1 mM phenol. This result is an extremely
good indicator that these isolates were able to degrade phenol as sole carbon and energy
73
source. Hence, these three potential isolates were chosen for further screening.
Table 4.1 Aerobic growth comparison of selected isolates on RM agar containing
1 mM phenol at 37oC
N0.
1.
Strain
RFTA-O2
Source
Influent of Floatation Tank A
Growth
+
2.
RAS-Cr1
Activated sludge
++++
3.
RSSF-Cr1
Soil from sludge farm
+
4.
RETL-Cr1
Effluent of biological treatment lagoon
++++
5.
RETL-Cr2
Effluent of biological treatment lagoon
+
6.
RETL-Cr3
Effluent of biological treatment lagoon
++++
7.
RETL-Y1
Effluent of biological treatment lagoon
-
8.
SMSF(+g)-Cr4
Soil-matrix from sludge farm
+
*
*
*
Key: - = no growth; + [weak growth (<30 colonies)], ++ [moderate growth (31 -100
colonies), +++ [dense growth (101 -300 colonies)], ++++ [extremely dense growth (>300
colonies)]
Of the 8 strains tested for growth on phenol, three isolates designated RAS-Cr1,
RETL-Cr1 and RETL-Cr3 (as marked in*) were selected for phenol tolerance test using
Ramsay broth supplemented with different concentrations of phenol. As shown in Figure
4.6, lower concentration of phenol of up to 4 mM (376 mg L-1) was tolerated by the 3
strains while RETL-Crl could still tolerate phenol up to 6 mM. Minimal growth for all
three was observed at the highest concentration of 10 mM (941 mg L-1) in RM broth
suggesting possible toxicity of phenol at such concentration. This maximum IPC level of
10 mM was chosen because the preliminary study done prior to this where these isolates
were screened on agar plate were able to tolerate phenol up to a maximum of 8 mM but
were unable to grow in RM agar containing 10 mM phenol (results not shown). Thus, a
cut off phenol concentration of 10 mM was chosen bearing in mind that increase in
phenol concentration could affect the growth and tolerance of these strains. This
behaviour is characteristic of toxic substrate metabolism as suggested by Hill and
74
Robinson, (1975), who reported that, as concentration of toxic substance increases, the
more detrimental it becomes to the organism.
Among these 3 strains, RETL-Cr1 has better tolerance towards phenol as
compared to the others (Figure 4.4). The tolerance level test of phenol by RETL-Cr1 is
presented in Figure 4.5.
8
CFU/mL (Log10)
7
6
5
4
3
2
1
0
1mM
2mM
4mM
6mM
8mM
10mM
Phenol conc. (mM)
Figure 4.4 Growth comparison of selected isolates; RETL-Cr1(Ŷ), RETL-Cr3 (Ɣ) and
RAS-Cr1 (Ÿ) grown aerobically on RM broth containing varying initial phenol
concentration as a sole carbon source incubated at 37oC after 24 h.
75
8
CFU/mL (Log10)
7
6
5
4
3
2
1
100
90
80
70
60
55
50
45
40
35
30
25
20
15
10
5
1
0
0
Phenol conc. (mM)
Figure 4.5 Test for phenol tolerance limit of isolate RETL-Cr1 in RM containing 1 mM
glucose incubated at 30oC, pH 6.5 after 96 h.
The former experiments were done without the inclusion of glucose as the initial
carbon source before the utilization of phenol as its primary carbon source.The next step
to investigate the possibility of using glucose as the cometabolite for phenol. Glucose has
been known as a cometabolite for the degradation of various cyclic hydrocarbon
xenobiotics. The term comtabolism is used to indicate any situation in which another
substrate enhances the biodegradation of a target compound. Glucose was added at a very
low concentration of 1 mM as opposed to the 2% (w/v) under normal circumstances. It is
interesting to note that RETL-Cr1 has considerably high tolerance towards phenol in the
range of 1 mM – 60 mM (94 mg L-1 - 5,647 mg L-1) in the presence of glucose suggesting
that tolerance level is very high for this isolate (Figure 4.7). The above experiment was
done in the presence of 1 mM glucose provided as the initial energy source. Strain RETLCr1 grew well on phenol concentration up to 30 mM (2,823 mg L-1) phenol. There was
no growth observed at concentrations beyond 70 mM up to 100 mM (6,588 – 9,411 mg L1
) possibly due to the toxic nature of phenol. Comparison on phenol tolerance limit of
RETL-Cr1 as shown in Figure 4.5 and Figure 4.6 shows that addition of 1 mM glucose
has improved the tolerance limit towards phenol. Glucose as a modulator could have
acted as co-substrate or as inducer (Rittman and Sáez, 1993). The interest in
76
biodegradation of pollutants in the presence of simple alternate carbon/energy sources is
to enhance biodegradation of pollutants through intentional addition of simple substrate
as suggested by Wang et al., (1996). Previously, it has been shown that addition of
glucose either enhance or inhibit degradation of phenol and its derivatives (Schmidt et
al., 1987; Hess et al., 1990; Kar et al., 1996; Wang et al., 1996).
The ability of RETL-Cr1 to grow in medium containing very high phenol
concentration was not surprising, given that phenols are found in wastewater discharges
from oil refinery (Pfeffer, 1979) and RETL-Cr1 may have adapted to chronic exposure of
this compound. An assumption can be made here that the sample obtained from the
wastewater of the Exxon Mobil petroleum refinery could contain considerable amounts
of phenol which may have allowed prior adaptation of the strain to phenol. These three
selected strains capable of growth in RM broth containing phenol were obtained from
activated sludge (RAS-Cr1) of a petrochemical industrial wastewater treatment plant and
biological treatment lagoon effluent (RETL-Cr1 and RETL-Cr3) of an oil refinery. These
results are expected as phenol and its derivatives are some of the major hazardous
compounds found in an oil refinery and other industrial wastewater (API, 1969; Pfeffer et
al., 1979; Watanabe et al., 1996a; Godjevargova et al., 2003).
The three strains were then tested for their capability to degrade phenol without
glucose. As shown in Figure 4.6 isolates designated RAS-Cr1, RETL-Cr1 and RETL-Cr3
were the most prominent degraders of phenol of which RETL-Cr1 was able to grow
prominently on RM agar in the presence of 3 mM (282 mg L-1) phenol with a percentage
survival of 97%, with no addition of glucose. As can be seen in Figure 4.6, RETL-Cr1
was the best isolate that was able to grow very densely in RM agar in the presence of 3
mM (282 mg L-1) phenol. All the other isolates were also able to grow on RM agar but at
a much lower density as compared to RETL-Cr1.
77
100
14
90
80
10
70
60
8
50
6
40
30
4
Survivality (%)
CFU/mL (Log)
12
20
2
10
0
0
RAS-Cr1
RETL-Cr1
RETL-Cr3
Selected isolates
Figure 4.6 Growth comparison of selected isolates grown aerobically in RM
broth containing 3mM phenol (Ŷ). Percentage of survival (Ɣ) of selected isolates at 37oC,
pH 6.5.
4.3.2
Morphological and physiological characterization of selected strains
Colony morphology criteria of these bacterial isolates were observed and
summarized in Table 4.2.
All isolates grown on solid media were observed as creamy and white gelatinous
colonies. Most were round in shape with either smooth or irregular edges.
Table 4.3 summarizes the results for biochemical tests, spore test, Gram stain
reactions and morphological description of the isolate strains. All the isolates were
catalase, urease and methyl red positive and able to ferment lactose. All the isolates
except for RAS-Cr1 were motile. Isolates RAS-Cr1 and RETL-Cr3 were Gram-negative
rods, whereas RETL-Cr1 was oval with budding. The length of cells of these selected
isolates ranged from 0.3-10 Pm. None of these three isolates was spore-forming.
78
When compared to the other strains, RETL-Crl showed budding characteristic and
was much bigger in size reaching up to 10Pm not typical of most eubacteria. Alcoholly
smell was emitted, typical of yeasts. For the colony morphology, the colonies were
raised and umbonate as viewed from the side. Therefore based on the colony
morphology and size, RETL-Cr1 could possibly be a yeast strain. The next step
phylogenetic characterization was then carried out on the best phenol degrader i.e. RETLCr1 based on its rDNA.
TPSB
EMOR
EMOR
RAS-Cr1
RETL-Cr1
RETL-Cr3
ETL
ETL
AS
cream
cream
cream
filiform
round
round
wavy
smooth
smooth
hilly
convex
drop-like
spreading
small colony
slimy
Table 4.2: Colony morphology of selected isolates grown on RM agar at 37oC after 24 hours incubation isolated from the two
sampling locations.
Strain
Location
Source
Colony characteristics
Remark
Colour
Configuration
Margin
Elevation
79
80
Table 4.3: Biochemical tests, cellular morphology, and Gram stain reaction of selected
strains
Test
Selected isolates
RAS-Cr1
RETL-Cr1
RETL-Cr3
FermentationReaction
Lactose
+ve(*g)
+ve
+ve
EnzymeActivity
Catalase
+ve
+ve
+ve
Citrate
+ve
-ve
-ve
Metyl red
+ve
+ve
+ve
Oxidase
-ve
-ve
-ve
Urease
+ve
+ve
+ve
V. Proskauer
+ve
+ve
+ve
TSI
A/A,G
A/A
A/A
Motility
+ve
+ve
-ve
Spore test
-ve
-ve
-ve
reaction
-ve
NA
-ve
Morphology
rod
oval (budding)
Length (um)
0.4
10
0.4
+ve
(pink)
+ve
(pink)
-ve
Gram stainrod
Growth on :MacConkey
agar
Blood agar
+ve(Ȗ)
+ve(Ȗ)
+ve (ß)
(*g)= gas production; + ve =positive result; -ve = negative result, NA = not applicable
81
4.3.3
Biodegradation of phenol by selected strains
It has been reported that in almost all ecosystems, the availability of carbon and
energy sources is extremely restricted (Morita, 1988, 1993). These carbons are available
in low concentration of a few micrograms per litre or lower. These carbon compounds
originate mainly from the hydrolysis of particulate organic matter, excretion products of
higher organisms and also from xenobiotic chemicals released into the environment
(Schmidt and Alexander, 1985; Münster and Chróst, 1990; Kirchman, 1993; Münster,
1993).
Biodegradation of phenolic pollutants in aquatic and terrestrial environments was
found not only restricted to the activity of a few adaptable microorganisms, but occurs
widely within bacteria (Bayly and Wigmore, 1973; Kohler et al., 1992), fungi (Jones et
al., 1993, 1995), yeast (Middlehoven, 1993). Among the prokaryotes, Pseudomonas
putida is a widely used microorganism for biodegradation of phenols as previously
shown in Table 2.3 (section 2.6.1) of Chapter 2. On the other hand, among the yeast
strains, Trichosporon sp. (Godjevargova et al., 2000), Rhodotorula sp. (KatayamaHirayama, 1991,1994), and Candida sp. (Bastos et al., 2000a; Komarkova et al., 2003)
were reported to degrade phenol.
The phenol degradation experiments revealed that among the selected strains,
RETL-Cr1 was the best phenol-utilizer with 100 % efficiency of degradation as shown in
Figure 4.7. In contrast to phenol degradation by isolate RETL-Cr1, phenol degradation
was not obvious in RM broth for RAS-Cr1 and RETL-Cr3.
Phenol removal efficiency (%)
82
120
100
100
80
60
40
20
16
14
RETL-Cr3
RAS-Cr1
0
RETL-Cr1
Selected isolates
Figure 4.7 Phenol removal efficiency by selected isolates in RM incubated at 37oC, pH
6.5.
To further investigate the potential of RETL-Cr1 in phenol degradation, RETLCr1 was allowed to degrade phenol at an IPC of 3 mM in the presence of 1mM glucose.
This is to understand the function of glucose during the process of phenol degradation
where the concentration of phenol was monitored along with the depletion of glucose.
Meanwhile cell density was measured simultaneously at OD600 where the whole
monitoring process was done over a period of 18 hours. It is noticeable that from figure
4.8, rapid depletion of glucose was observed within 1 hour of incubation. The utilization
of phenol by isolate RETL-Cr1 was almost concurrently with glucose. Most of the
phenol, however, was utilized during the exponential phase of growth by RETL-Cr1.
83
0.8
0.7
250
0.6
200
0.5
0.4
150
0.3
100
OD600
Phenol and glucose conc. (mg
L-1)
300
0.2
50
0.1
0
0
0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18
Time (h)
Figure 4.8 Degradation of phenol (Ŷ) against time and glucose utilization(Ɣ) by growth
pattern of RETL-Cr1 (Ƒ ) in RM containing 3 mM phenol at 37oC, pH 6.5.
This figure also illustrates that complete degradation shows that 100% removal of
phenol could be achieved after 15 hours of incubation in shake flask culture. Since
phenol and glucose were simultaneously used up with a complete depletion of glucose
after during the first hour suggesting that it is used as an initial energy source. The strain
reached its maximum growth within 17 hours and eventually declined. The growth
indicates that as phenol decreases, growth increases. In shake flask culture or otherwise
known as batch culture, strain RETL-Cr1 exhibited a degradation rate of 18.8 mg L-1 with
100% removal efficiency at specific growth rate (µ) = 0.313 h-1 after 17 hours incubation.
Both phenol and glucose were degraded simultaneously; glucose at a faster rate achieving
80% degradation after 2 hours. This could be referred to as simultaneous substrate
utilization pattern as suggested by Egli, (1995) where two substrates were utilized
together.
Following the investigation on RETL-Cr1, a comparison of the three potential
isolates namely RETL-Cr1, RAS-Cr1 and RETL-Cr3 was also accomplished. As
summarized in Table 4.4, the following rates of degradation were achieved. It is
interesting to note that RETL-Cr1 could degrade phenol (IPC 3mM) at a rate of 100%
84
within 18 hours, whereas RAS-Cr1 and RETL-Cr3 could only obtain a degradation rate
of less than 3 mg L-1 with 14 to 16% removal efficiency at specific growth rates (µ),
0.323 h-1 to 0.363 h-1. The difference in rate of degradation for all could be due to their
differences in microbial metabolism that attributed to genetic differences and their
responses to changes in their environment as suggested by Shuler and Kargi, (2002) and
Landis and Yu (2003). Based on these observations, isolates RAS-Cr1 and RETL-Cr3
were abandoned and focus was only placed on RETL-Cr1.
100
16
14
RETL-Cr1
RETL-Cr3
RAS-Cr1
2.80
2.85
18.8
2.77
2.67
10.36
0.323
0.359
0.313
18
18
17
243.03
253.517
0.225
49
19
97
Table 4.4: Growth kinetics & performance of phenol degradation at 3mM IPC by selected isolates at 37oC, pH 6.5
µ
Max. degradation Residual phenol
Survival
Isolate
Average phenol biodegradation
Xmax
Efficiency (100%) Rate (mg L-1 h-1)
(g L-1)
(h-1)
time (h)
(mg L-1)
(%)
85
86
4.3.4
Characterization and Identification of the phenol-degrading RETLCr1
The strain of choice that is RETL-Cr1 was characterized morphologically and
biochemically. Further identification of the strain by molecular typing was attempted
thereafter. Strain RETL-Cr1 formed small, round, cream-coloured colonies, with no
pigmentation on RM agar after 24 hours incubation (Figure 4.9).
Figure 4.9 Colony morphology of RETL-Cr1 on RM agar under stereo- microscope
(x12)
The cells were oval, measuring 10 µm long and 6 µm in diameter and 1 budding
was observed (Figure 4.10). Isolate RETL-Cr1 was motile, catalase, urease and methyl
red positive and able to ferment lactose and sucrose. Details of the physiological and
biochemical characteristics of strain RETL-Cr1 is summarized in Table 4.3.
87
budding
10µmx6µm
Figure 4.10 Gram morphology of RETL-Cr1 magnified x1000 under light microscopy.
PCR-based methods (Restriction Fragment Length Polymorphisms (RLFP), PCR
with species-specific probes, random amplification of polymorphic DNA analysis
(RAPD), and Multiplex PCR using internal transcribed spacer (ITS) 1 and 2 regions) for
the rapid detection and identification of yeast strains such as Candida species have been
described by (Morace et al., 1997; Shin et al., 1997, 1999; Stefan et al., 1997; Jackson et
al., 1999; Fujita and Hashimoto, 2000 and Fujita et al., 2001). The multiplex PCR using
internal transcribed spacer (ITS) 1 and 2 regions is sensitive, rapid and specific for yeast
organisms (Fujita et al., 2001).
The size of the PCR product of C. tropicalis RETL-Cr1 obtained in this study was
approximately 450 bp (Figure 4.11), just slightly smaller than those reported by Fujita et
al., (2001) and Guillamón et al., (1998). They reported PCR products of more than
500bp. According to Fujita et al., (2001) amplification of all fungi tested using ITS1 and
ITS4 primers yielded fragments 350 to 880 bp long (Fujita et al., 2001).
88
Lane
1 2
10000 bp
500 bp
400 bp
100 bp
Figure 4.11 The amplified DNA from C. tropicalis RETL-Cr1 ribosomal gene generated
using TS1 and TS4 primers is shown in Lane 2. Lane 1, molecular weight size reference
marker (100-bp ladder).
1
GAGTCTTCGTCTCAAGGACATTGATTCCATGGGT
35
CTTTTTTTAGTACTGTTACTTTGGCGGCAGGAGTAAATATCTTACCGCCAGAGGTCTTTA
95
TAACACTCAATTTAATTTTATTTATTATTCAAAGACGATTATATTTTATAAATAGTCAAA
155 ACTTGTCAACAACGGATCTCTTGGTTCTCGCATCNATGAAGAACGCAGCGAAATGCGAT
214 ACGTAATATGAATTGCAGATATTCGTGAATCAT CGAATCTTTGAACGCACATTGCGCCCT
274 TTGGTATTCCAAAGGGCATGCCTGTTTGAGCGTCNTTTCTCCCTCAAACCCCTGGGTTTG
334 GTGTTGAGCAATACGCTTAGGTTTGTTTGAAATATTTCCAATTGTGGACAACTATTTATG
394 TTATAGCGACTTAGGTTTATCCAAAACGCTACAACCATAAAGGAAGTCCACTGAATAAT
453 TTCATAACTTTTGACCTCAAATCAGGTAC
Figure 4.12 Complete sequence of the 5.8S rDNA (Italics) flanked by adjacent
ITS1 and ITS2 regions of C. tropicalis RETL-Cr1.
Based on a BLASTN search of GenBank, the complete sequences of ITS1-5.8S
rDNA-ITS2 regions and portions of I8S and 28S for the purified DNA products of
RETL-Cr1 (Figure 4.12) shared 98% similarity with Candida tropicalis ( score: 404 bits,
89
E value: e-110). The isolate RETL-Cr1 was redesignated Candida tropicalis RETL-Cr1.
The strain was deposited into the GenBank under the accession number AY725426.
According to Cavalca et al., (2004), the degradative ability of microorganisms
was spread over different genera, reflecting the importance of functional diversity in
polluted environments to decontaminate mixture of compounds. The ability to metabolize
aromatic compound, is well known for free living soil and water dwelling
microorganisms (Karasevitch, 1982). For instance, Candida has been found to be the
most abundant yeast genus present in soil (Mok et al., 1984) and oil-contaminated
seawater (Zinjarde and Pant, 2002). According to Ballestros et al., (1991), C. tropicalis is
known by its vigorous growth on various carbon sources and its phenol-degradative
capability is well documented (Shimizu et al., 1973; Neujahr et al., 1974; Kumaran,
1980; Krug and Straube, 1986; Kumaran and Parachuri, 1997;Chang et al., 1998; Bastos
et al., 2000a; Ruiz-Ordaz et al., 2001; Chen et al., 2002; Vojta et al.,2002, and
Komarkova et al., 2003). Candida tropicalis that were known capable of degrading
phenol have been isolated from both phenol contaminated and uncontaminated
environments (Vojta et al., 2002); activated sludge (Komarkova et al., 2003; Yan et al,
2005); phenol bearing industrial wastes (Kumaran, 1980) from pristine soils (Bastos et
al., 2000a). Studies by Bastos et al., (2000b) and Koutny et al., (2003) supported the idea
that natural uncontaminated environment contain sufficient genetic diversity to make
them valid choices for the isolation of microorganisms useful in bioremediation.
An investigation on the origin of the phenol-degrading Candida tropicalis by
other researchers previously reported is shown in Table 2.4 of Chapter 2 (Section 2.6.2).
4.4
Conclusions
Microbial isolation and screening is an important stage for evaluating the
biodegradation potential of the isolate towards a specific pollutant. In conclusion, a new
indigenous phenol-degrading yeast strain RETL-Cr1, was isolated from wastewater
90
treatment plant effluent of Exxon Mobil oil refinery in Malaysia. The most distinctive
feature of the strain RETL-Cr1 is that it has a very high tolerance limit towards phenol
reaching up to 60 mM (5,647 mg L -1) and was able to degrade phenol efficiently at initial
phenol concentration of 3 mM (282 mg L-1) in the presence of glucose. The degradation
rate achieved was 18.8 mg L-1 with 100% removal efficiency at µ = 0.313 h-1 after 17
hours incubation. Based on a BLASTN search of GenBank, the complete sequences of
ITS1-5.8S rDNA-ITS2 regions and portions of I8S and 28S for the purified DNA
products of RETL-Cr1 shared 98% similarity with Candida tropicalis. The isolate RETLCr1 was redesignated Candida tropicalis RETL-Cr1.
This preliminary screening for the ability of C. tropicalis RETL-Cr1 suggests
appreciable degradation potential for phenol which was the basis for further investigation.
The parameters that need consideration for optimization may include temperature, pH,
initial phenol concentration (IPC) and effect of glucose in mixed substrate on phenol
degradation. This yeast may be the suitable organism for use in future bioremediation
processes for industrial effluents or contaminated soils. This is the first report in Malaysia
of its kind of an indigenous phenol-degrading yeast.
91
CHAPTER 5
BIODEGRADATION OF PHENOL IN BATCH CULTURES OF YEAST Candida
tropicalis RETL-Cr1
5.1
Introduction
Many organisms, both bacteria and fungi could biodegrade a diverse range of
hydrocarbons, including aromatic and aliphatic molecular structure (Atlas, 1981; Leahy
and Colwell, 1990; Cerniglia, 1992). Aerobic catabolism of aromatic compounds has
been investigated for a variety of microorganisms and for different natural and xenobiotic
compounds (Gibson and Subramaniam, 1984; Haggblom and Valo, 1995; Wild et al.,
1997). Phenol degradation by pure and mixed microbial cultures had been studied
(Ahmed, 1995; Collins and Daugulis, 1997a; Schroder et al., 1997; Chang et al., 1998;
Ruiz-Ordaz et al., 1998).
Most of the earlier studies concerning degradation of phenol have been performed
using bacteria, mainly from Pseudomonas genus. Some investigations have also been
carried out using yeast strains to degrade phenol (Godjevargova et al., 2003). Yeasts are
widely distributed in nature and have extremely diverse metabolic capabilities that can
utilize a wide range of nutrients under a variable of environmental conditions (TornaiLehoczki et al., 2003). For example, Candida tropicalis are able to utilize disaccharides,
alkanes, alkane derivatives, fatty acids, phenols and crude oil (Kurihara et al., 1992;
92
Kawachi et al., 1997; Murzakov et al., 2003). Other industrial importance of Candida
tropicalis are for the production of xylitol (Yahashi et al., 1996; Azuma et al., 2000;
Walther et al., 2001; Lima et al., 2003) and production of microbial protein and fodder
yeast (Stanton and Dasilva, 1978).
Candida tropicalis, a diploid asexual organism (Picataggio et al., 1991) has the
capability of effective degradation of high concentration of phenol (Krug et al., 1985;
Chang et al., 1998; Ruiz-Ordaz et al., 1998, 2000, 2001; Chen et al., 2002). In this study,
a yeast strain designated Candida tropicalis RETL-Cr1 which also has the capacity to
degrade phenol is the organism of interest and focus has been placed on this strain for its
phenol degrading capability.
Microorganisms are able to grow and sustain themselves by getting nutrients,
electron and energy from their environments. Biodegradation of organic substrates
provide microorganisms with energy and building materials that are used for growth of
new cells, cell maintenance and co-metabolism of other less degradable substances
(Cornelissen and Sijm, 1993).
We have shown that RETL-Cr1 could possibly be a yeast strain of the genus
Candida. From the phylogenetic analysis done, it could be confirmed as a Candida
tropicalis and were designate this strain to be C. tropicalis RETL-Cr1. From the literature
it could be observed that both bacteria and fungi could biodegrade a diverse range of
hydrocarbons, including aromatic and aliphatic molecular structure (Atlas, 1981; Leahy
and Colwell, 1990; Cerniglia, 1992) especially through aerobic degradation. Phenol
degradation by pure and mixed microbial cultures has also been studied (Ahmed, 1995;
Collins and Daugulis, 1997a; Schroder et al., 1997; Chang et al., 1998; Ruiz-Ordaz et al.,
1998).
Candida tropicalis, a diploid asexual organism (Picataggio et al., 1991) has the
capability of effective degradation of high concentration of phenol (Krug et al., 1985;
Chang et al., 1998; Ruiz-Ordaz et al., 1998, 2000, 2001; Chen et al., 2002). In this
93
present study, a yeast strain designated Candida tropicalis RETL-Cr1 which also has the
capacity to degrade phenol is the main organism being studied here.
Phenol degradation can be achieved in either batch or continuous system and most
researchers have reported batch or continuous phenol biodegradation. Most bioprocesses
are based on batch reactors (Shuler and Kargi, 2002). The principal advantages of batch
cultures are; low contamination risk, the ability to run different successive phase in the
same vessel, and close control of the genetic stability of microorganism (Scragg, 1992;
Panikov, 1995). One disadvantage of fed-batch is that it requires previous analysis of the
microorganisms that is its requirements and the understanding of its physiology with the
productivity (for instance, its capability to degrade certain toxic chemicals) (Shuler and
Kargi, 2002).
This chapter describes the attempt to study the degradation of phenol under
various temperatures, initial pH values and in a suitable medium such as Ramsay medium
in the presence or absence of glucose using free cells of C. tropicalis RETL-Cr1. The
ability to degrade phenol in batch fermentation technique by free cells of C. tropicalis
RETL-Cr1 in the presence and absence of glucose at different temperature and pH was
compared. The purpose of the present study was to assess and compare the ability of an
indigenous C. tropicalis strain RETL-Cr1 to degrade phenol in batch fermentation
technique using both shake-flask and a bioreactor.
5.2
Materials and Methods
5.2.1
Culture media
In all the experiments, the medium described by Ramsay et al., (1983) containing
(g/L): 2.0g NH4NO3, 0.5g
KH2PO4, 1.0g K2HPO4, 0.5g MgSO4.7H2O, 0.01g
CaCl2.2H20, 0.1g KCl and 0.06g yeast extract was used. Phenol solution was filter
sterilized using a 0.2 µm membrane filtration before addition into the medium. Phenol
94
was added as a sole carbon source. The pH was adjusted to between 6.5-6.8.
5.2.2
Batch fermentation: Shake-flask culture
Ten mL of overnight culture was transferred into 250 ml conical flask containing
90 mL of the medium with varying phenol concentrations of 3 mM, 5 mM, 7 mM and 10
mM, added only after sterilization. Cultures in duplicates were incubated at 30oC with
shaking at 200 rpm. Samples were withdrawn at a regular intervals and analyzed for cell
growth, phenol, catechol and cis,cis-muconic acid concentrations.
5.2.1.1 Effect of temperature on phenol degradation
The manipulation of temperature studies were carried out at temperatures at 30oC,
37oC and 40oC for C. tropicalis RETL-Cr1 under batch cultivation (shake-flask) was
done. Preliminary study has shown that C. tropicalis RETL-Cr1 does not grow well
below 30oC and above 40oC. Incubation took place at agitation of 200 rpm for 18 h.
5.2.1.2 Effect of pH on phenol degradation
pH 4.5, 5.5, 6.5, 7.0 and 8.0 for C. tropicalis RETL-Cr1 under batch cultivation
(shake-flask) were applied. Incubation took place at an agitation speed of 200 rpm for 18
h.
5.2.1.3 Effect of glucose on phenol degradation
To study the effect of glucose on phenol degradation by C. tropicalis RETL-Cr1
the culture medium was supplemented with 1 mM glucose and one without the addition
of glucose. The initial pH value of the medium was adjusted between 6.5 – 6.8. The
cultures were incubated at 37oC and agitated at 200 rpm.
95
Duplicates were set up and samples were withdrawn at regular intervals and
analyzed for cell density and levels of residual glucose and phenol.
5.2.3
Batch fermentation: Bioreactor culture
Fermentation runs, repeated twice were conducted batchwise in a thoroughly
mixed reactor using sterilized media. RM broth (0.5 L) containing 3 mM phenol was
placed in the reactor and inoculated with 50 mL of overnight yeast cultures. Fermentation
was carried out in a Biostat B 2L model fermenter from B. Braun Biotech Int. GmbHIn
for 18 hours under the following conditions: incubation temperature at 30oC and impeller
speed of 200 rpm. pH was not controlled since the fermentation values of 6.5-6.8 were
suitable for effective growth of the yeast. However, pH was monitored throughout the
fermentation process, five mL samples were taken periodically throughout the operation
for determination of residual phenol, catechol, cis,cis-muconic acid and optical density.
5.2.4
Experimental Design
The flowchart of the experimental design carried out in this study is summarized
in Figure 5.1.
96
Batch
phenol degradation
Shake-flask
at 200 rpm
Bioreactor
at 30oC, pH 6.5,
IPC=3mM, 200 rpm; glucose
Optimization
Temperature
(30oC (+ & glucose), 37oC &
40oC; IPC=3mM
pH
(4.5, 5.5, 6.5, 7,
8), 30oC, IPC=
3mM, -glucose
IPC
(3mM, 5mM, 7mM
& 10mM) at 30oC,
pH 6.5, - glucose
Determination and quantification
of catechol and cis,cis-muconic acid
Comparison on the kinetics and
performance of phenol degradation by
C. tropicalis RETL-CR1
Figure 5.1 Experimental design of phenol degradation by C. tropicalis RETL-Cr1 in
batch culture.
5.2.5
Sample Analysis
5.2.5.1 Determination of biomass concentration
The experimental procedures were presented in section 3.5.1
97
5.2.5.2 Determination of average phenol degradation rate
The experimental procedures were presented in section 3.5.3
5.2.5.3 Determination of phenol, catechol and cis,cis-muconic acid
The experimental procedures were presented in section 3.5.5
5.3
Results and Discussion
5.3.1 Optimization of phenol degradation in shake-flask culture
5.3.1.1 The effect of temperature on phenol degradation in shake-flask culture
The next set of experiments were carried out at temperature ranging from 30oC to
40oC to study the effect of temperature on phenol degradation by C. tropicalis RETL-Cr1
in the absence of glucose. The reason for choosing this temperature range temperature is
because preliminary studies done prior to this has shown that temperature below 30oC or
above 40oC could not support growth of C. tropicalis RETL-Cr1, what more degrade
phenol (results not shown). Figure 5.2 illustrates the effect of temperature on the rate of
phenol degradation.
Phenol degradation rate (g L-1 h-1)
98
0.03
0.0257
0.025
0.0188
0.02
0.015
0.01
0.005
0.0009
0
30
37
40
o
Temperature ( C)
Figure 5.2 The effect of temperature on the average phenol degradation rate by C.
tropicalis RETL-Cr1 in the absence of glucose in RM medium containing 3 mM phenol,
pH 6.5 in shake-flask culture.
High phenol degradation efficiency of phenol could be possible at a temperature
range of between 30-37oC. Phenol degradation capability of C. tropicalis RETL-Cr1 was
optimum at 30oC under aerobic condition using phenol as sole carbon source. Under the
optimized conditions of 30oC and pH 6.5, in RM containing initial phenol concentration
of 3 mM phenol, C. tropicalis was able to degrade phenol effectively at a rate of 0.0257 g
L-1 h-1. Consequently, phenol degradation rate decreased with increasing temperature. A
shift to higher temperature of 40oC appears to affect the biodegradation capability. The
biodegradation rate was reduced by 29-fold. A study performed by Perron and Welander
(2004) reported that a combined process consisting of a fungal (Mortierrella sarnyensis
Mil’ko) and a bacterial step was shown to be efficient in the degradation of phenol and
cresols even at low as 4oC. Our results can be supported by the above findings where it
confirms that mesophilic temperature between 30o-37oC was suitable for phenol
degradation by either bacteria or fungus.
99
The influence of temperature on enzyme activity could be rationalized in the
following manner. The rate of enzyme reaction increases with temperature increase and
rate of movement of molecules is slower at lower temperature than at higher temperature;
so there is not enough energy to spark a chemical reaction at a lower temperature. Thus,
temperature below 30oC may not be enough to start a reaction. However, as temperature
approaches 37oC, the enzymatic reaction began to decline. This is terribly obvious at
40oC, where rate of degradation was negligible. It could be speculated that decrease in
activity was as a result of protein denaturation (Skopes, 1994) which in turn result in the
loss of its three-dimensional structure. Protein depend on three dimensional structure to
maintain its activity, therefore the unfolding of protein due to denaturation may have
broken bonds in the protein structure crucial for maintaining its shape. Hence, it could be
deduced that at 40oC, the protein has lost its three dimensional structure, and inactivate
the protein.
Kinetic studies on the effect of temperature were also done to elaborate the
importance of temperature on growth, productivity and rates of degradation. Table 5.1
summarizes the results of the effect of temperature on degradation of phenol by C.
tropicalis RETL-Cr1 in RM medium with 3 mM initial phenol concentration in the
absence of glucose maintained at pH 6.5 in batch culture (shake-flask).
100
Table 5.1: Effect of temperature on phenol degradation by C .tropicalis RETL-Cr1 at
three different temperatures, pH 6.5 (shake-flask) after 18h incubation.
Kinetics parameters/ Performance
30
Temperature (oC)
37
40
Xmax (gdw L-1)
9.765
9.23
0.80
µ (h-1)
0.3718
0.3226
(-)
Yx/s (g g-1)
29
30
31
Ypc/s (g g-1)
0.088
0.022
0.05
0.003
0.0003
0.0015
Catechol productivity (g L-1 h-1)
0.003
0.0005
0.0001
Catmax (g L-1)
0.0204
0.0023
0.0008
t (Catmax) (h)
7
5
8
YpccMA/s (g g-1)
0.039
0.030
0
YpccMA/x (g g-1)
0.0013
0.001
0
ccMA productivity (g L-1 h-1)
0.0006
0.0005
0
ccMAmax (g L-1)
0.011
0.0088
0
t (ccMAmax) (h)
18
18
0
Phenol biodegradation rate
(g L-1 h -1)
0.0257
0.0188
0.0009
Phenol removal efficiency (%)
100
100
6.2
Incubation time (IT) (h)
17
18
18
TL (lag time) (h)
3
3
3
Biodegradation time (BT) (h)
14
15
15
Ypc/x
(g g-1)
As shown in Table 5.1, at an optimum temperature of 30oC was conducive for
phenol degradation giving 100% efficiency. It is interesting to point out that although a
specific growth rate (µ) of 0.3718 h-1 and with a cellular yield of 29 g cell dry weight per
g phenol utilized was achieved. The rate of biodegradation rate achieved at 30oC was
0.0257 g L-1 h -1. This is twice as high as that achieved at 37oC. Although a rate of
degradation of 100% was achieved at 37oC, µ was slightly lower and biodegradation rate
101
was comparatively lower. As indicated here, at 40oC, no cell growth was observed and
biodegradation rate was negligible (where the biodegradation rate of 0.0009 (g L-1 h -1 was
achieved). Thus, a shift to higher temperature also appears to affect the biomass
concentration. The decline of microbial activity beyond the optimum temperature could
be due to the effects on enzyme denaturation as suggested by Shuler and Kargi, (2002)
and Suthersen, (1999). This involves the denaturation of the catalytic site that lead to
inactivation of the enzyme.
We can possibly hyphothesize that aerobic degradation of phenol by C. tropicalis
RETL-Cr1 could take the ortho pathway. Based on this, it is predicted that two
intermediary products could be formed; namely catechol and cis,cis-muconic acid.
According to Alexieva et al., (2004), the efficiency of phenol degradation depends on the
properties of two key enzymes; monoxygenase phenol hydroxylase (PH) (EC 1.14.13.7)
(Neujahr and Gaal, 1973). The first step in aerobic degradation of phenol is the
conversion of phenol to catechol by monoxygenase phenol hydroxylase (Neujahr and
Gaal, 1973). Catechol is the central intermediate in the phenol degradation pathway.
Catechol is then further degraded to cis,cis-muconic acid (ccMA) which is catalysed by
catechol 1,2 dioxygenase (C1,2D). According to Spanning and Neujahr (1991) C1,2D is
induced simultaneously with phenol hydroxylase (PH).
As shown in Table 5.1 during phenol degradation by C. tropicalis RETL-Cr1,
phenol was used mainly for biomass production instead of the intermediate products,
catechol and ccMA. This is clearly shown by the kinetic parameter yield as follows: with
a cellular yield g cell dry weight per g phenol utilized (Yx/s) = 29-31 g g -1, product yield
(catechol) per g phenol utilized (Ypc/s)= 0.01 – 0.05 g g-1 and product yield (cis,cismuconic acid) per g phenol utilized (YpccMA/s) = 0 - 0.039 g g-1.
Phenol biodegradation rate, catechol and ccMA productivity were higher at 30oC
and decreased with increasing temperature. The maximum concentration (Catmax =
0.0204 g L-1), high productivity of catechol (0.003 g L-1 h-1), ccMAmax (0.011 g L-1) and
ccMA productivity (0.0006 g L-1 h-1) were achieved at 30oC as compared to 37oC and
102
40oC. The production of these intermediates could indicate that the two major enzymes
primarily PH and C1,2D could be affected by the temperature. It could be postulated that
the optimum temperature for both PH and C1,2D in C. tropicalis RETL-Cr1 could be
around 30oC, judging by the Catmax (g L-1) and ccMAmax (g L-1) at this temperature.
Hence at this optimum temperature of 30oC, both enzymes could work optimally thus a
shorter degradation time was achieved. Our result is in good agreement with that reported
by Santos and Linardi (2004) where a comparatively higher PH and C1,2D activities
were also observed at high phenol degradation rate by two fungal sp. namely Graphium
and Fusarium sp. incubated at optimum temperature of 30oC.
If these two enzymes are actually affected by temperature variation, it is possible
to postulate that at an optimum temperature, C1,2D catalyzed the conversion of catechol
to ccMA effectively leaving no accumulation of catechol in the medium that may inhibit
PH which is responsible for the degradation of phenol to catechol. This suggests that
30oC could be the optimum temperature for both PH and C1,2D in C. tropicalis RETLCr1 where this phenomenon is similar to that reported of Trichosporon cutaneum
(Mörtberg and Neujahr, 1987). The following illustration (Figure 5.3) demonstrates the
possibility of such optimum activity for PH and C1,2D at 30oC.
103
Temperature range of
30oC optimum for PH
activity
Temperature range of 30oC
optimum for C1,2D
activity
G
L
F
+
+
D
2
2
F
L
Q
R
2 & 2&
F
X
&
& + P
V
L
F
V
L
F
+
+
2
O
R
K
F
H
W
D
F
O2+NADPH + H+
Phenol
NADP
No inhibition on PH by
catechol
+
+
O2
C1,2D
2H+
& &
+
+
2
+
2
PH
H2O
No accumulation of
catechol
Figure 5.3 Hypothetical illustration of PH and C1,2D optimum activity during phenol
degradation by C. tropicalis RETL-Cr1 at optimum temperature. PH = phenol
hydroxylase, C1,2D = catechol 1,2-dioxygenase, [Reaction: phenol + O2 + NADPH + H+
NADP+ + H2O + catechol (Mörtberg and Neujahr, 1987); catechol + O2
ccMA +
2H+ (Ngai et al., 1990)].
The optimum physiological temperature of 30oC for phenol degradation has been
quoted for a number of mesophilic bacteria, fungi and yeast. Some examples are; bacteria
such as Bacillus sp. (Bushwell, 1975), Pseudomonas sp. (Allsop et al., 1993; Zilli et al.,
1996; Reardon et al., 2000), Fungi such as Graphium and Fusarium sp. (Santos and
Linardi, 2004), and yeast strain such as Candida tropicalis (Shimizu et al. 1973; Yan et
al., 2005) and Trichosporon cutaneum (Yang and Humphrey, 1975).
At 40oC, phenol hydroxylase (PH) activity in C. tropicalis RETL-Cr1 seemed to
be affected as indicated by a lower Catmax (0.0008 g L-1) and there was no ccMA detected
to suggest that CI,2D could have been denatured. This can be supported by findings
reported by Fernandez et al., (2005). In this study by Fernandez et al., (2005) the stability
of the enzyme in C. tropicalis RETL-Cr1 become extremely unstable as the temperature
increased from 37oC to 50oC. This could be also due to conformational changes in the
104
protein structure or denaturation of the catalytic site that led the inactivation of the
enzyme as discussed before. These results can be also supported by Rochkind et al.,
(1986) who suggested that an increase in temperature just a few degrees above the
optimum can slow growth dramatically by the inactivation of the enzyme systems, and
continued exposure to high temperature may denature membrane lipids, resulting in cell
death (Gaudy and Gaudy, 1988).
5.3.1.2 The effect of glucose on phenol degradation
Microorganisms acquire nutrients, electron and energy from their environments to
support growth. Biodegradation of organic substrates provide microorganisms with
energy and building materials that are used for growth of new cells, cell maintenance and
co-metabolism of other less degradable substances (Cornelissen and Sijm, 1993). The
basic process in microbial metabolism is best illustrated using a model by Rittmann and
Sàez, (1993) presented in Figure 5.4. This model could possibly be applicable to illustrate
the effect of glucose on the electron and energy flows in C. tropicalis RETL-Cr1.
D
IC
2e
DOX
A
-
2e
ICH2
2e
NUTRIENTs
-
-
ADP
+
Pi
ATP
A red
Biomass
Synthesis
Biomass
Maintenance
Figure 5.4 Typical electron and energy flows in a bacterial cell (Rittmann and Sàez,
1993). D = primary electron-donor substrate, DOX = oxidized electron-donor substrate, A =
primary electron-acceptor substrate, Ared = reduced electron-acceptor substrate, ICH2 = reduced
internal cosubstrate, ATP = adenosine triphosphate, ADP = adenosine diphosphate, and Pi =
inorganic phosphate.
105
In nature microorganisms grow mostly in a medium supplemented with additional
substrates (Harder and Dijkhuizen, 1982). Hence, growth could be manipulated by
addition of two or more nutrients simultaneously (Rutgers et al., 1990; Egli, 1991). In
general, microbial degradation of a compound in a mixture can be strongly influenced by
other compounds present in the mixture (Egli, 1995). If a microbial population is grown
on mixed substrates present in the medium, the microbes consume only one, or both the
substrates. Consequently, several utilization patterns can be observed. In a mixed
substrates, individual substrates can have a synergistic, antagonistic, or no effect on one
another, resulting in a growth rate that is higher, lower, or the same than if the substrates
were present individually (Meyer et al., 1984; Saéz and Rittmann, 1993; Egli, 1995).
The following experiments were initiated to investigate and conduct a kinetic
analysis on phenol degradation in the presence or absence of glucose by C. tropicalis
RETL-Cr1. Phenol and glucose were selected for two reasons. First, phenol is a toxic
compound representing wastes of industrial origin. Second, glucose is non-toxic, a
common substrate which can represent wastes of urban or agricultural origin.
Table 5.2 shows that C. tropicalis RETL-Cr1 was found to degrade phenol
completely in media either in the presence or absence of glucose. However, in the
presence of glucose, a lower specific growth rate and degradation rate were achieved.
The degradation rate of phenol in the presence of glucose was reduced by 1.4-fold.
106
Table 5.2 Effect of glucose on phenol degradation by C. tropicalis RETL-Cr1 at
30oC, pH 6.5
Kinetic parameters/Performance
Ramsay medium(RM)
With glucose
Without glucose
Xmax (gdw L-1)
11.8
9.765
µ (h-1)
0.3065
0.3718
Yx/s (g g-1)
36
29
Ypc/s (g g-1)
0.58
0.088
0.0067
0.003
Catechol productivity (g L-1 h-1)
0.0096
0.003
Catmax (g L-1)
0.067
0.0204
t (Catmax) (h)
7
7
0.064
0.039
0.0018
0.0013
0.0011
0.0006
ccMAmax (g L )
0.018
0.011
t (ccMAmax) (h)
16
18
Phenol biodegradation rate
(g L-1 h -1)
0.0188
0.0257
Phenol degradation efficiency (%)
100
100
Ypc/x
(g g-1)
-1
YpccMA/s (g g )
-1
YpccMA/x (g g )
-1
-1
ccMA productivity (g L h )
-1
These substrates utilization could be due to uncompetitive cross inhibition as
suggested by Wang et al., (1996). According to Shuler and Kargi (2002), noncompetitive are not substrate analogs whereby an inhibitor will bind on site other than the
active site and reduce enzyme affinity to the substrate. Previous studies have shown
similar negative effects of glucose on phenol degradation by different microbial species.
For examples; C. tropicalis (Bastos et al., (2000a), subsurface soil microorganisms
(Swindoll et al., 1988) activated sludge culture (Rozich and Colvin, 1986), Arthrobacter
species (Kar et al., 1996). In this study by Kar et al., (1996) phenol degradation was
completely inhibited when glucose concentration was at 2 g L-1.
107
According to Rittmann and Sàez, (1993) inhibition can affect biodegradation of a
secondary substrate directly or indirectly. In the direct case, the inhibitor affects the
enzymes that are responsible for degrading such substrate. In the indirect case, the
inhibitor retards the electron and energy flows of primary-substrate utilization and will
slow the degradation of the secondary substrate by decreasing the amount of active
biomass and/or altering the availability of internal co-substrates, such as ICH2 (reduced
internal cosubstrate).
It is known that the structure of glucose and phenol are not similar and degraded
by different enzymes. However, according to Wang et al., (1996) glucose has an affinity
for the enzyme involved in the biodegradation of phenol. This binding does not result in
product formation but reduces the affinity of the enzyme for phenol and hence the phenol
degradation rate is reduced.
Further investigation on the negative effects of glucose on phenol degradation
also had been investigated by Kar et al., (1996) and found that glucose inhibited one of
the enzymes which participate in phenol degradation metabolism, inhibits phenol
transportation inside the cell, and cell prefers easily degradable glucose. According to
Spånning and Neujahr (1991) phenol hydroxylase and catechol 1,2-dioxygenase (C1,2D)
was very low in glucose grown cells of a yeast Trichosporon cutaneum. Chang et al.,
(1995) reported that phenol hydroxylase (PH) of C. tropicalis M4 was greatly suppressed
when the cells was grown on medium containing 0.2% glucose and there was no phenol
hydroxylase activity detected when glucose concentration was at 2%. Furthermore, study
by Bastos et al., (2000a) also found that glucose and acetate at 5 mM repressed
degradation of 7.5 mM phenol by C. tropicalis. The results from these studies suggested
that glucose might have blocked either partially or totally the synthesis of phenol
hydroxylase and C1,2D of these yeasts. This repression of catabolism of phenol by
glucose is referred as catabolite repression as described by Ampe et al., (1998).
However, in this present study C. tropicalis RETL-Cr1 was still able to degrade
phenol efficiently in the presence of glucose suggesting there was no or low direct
108
inhibition on the key enzymes (PH and C1,2D) of C. tropicalis RETL-Cr1. The
successful degradation of phenol in the presence of glucose could probably be due to
relatively low concentration of glucose (1mM = 198 mg L-1) as compared to that reported
by Chang et al., (1995) and Bastos et al., (2000a). Thus direct inhibition of glucose on
PH and C1,2D was absent or minimum. From all these studies, it can be concluded that
the extent of suppression on synthesis of phenol hydroxylase depends on the
concentration of glucose. Higher concentration of glucose will probably lead to higher
repression on the synthesis of phenol hydroxylase and catechol 1,2-dioxygenase.
Consequently phenol biodegradation rate decreases as the concentration of glucose
increases.
Therefore, our study indicates that glucose could have caused indirect inhibition
by retarding the primary electron and energy flows of primary-substrate utilization in C.
tropicalis RETL-Cr1 thus reduced the degradation rate of phenol as illustrated by Figure
5.5.
109
Glucose as noncompetitive inhibitor
slow donor oxidation
Decouplers reduce energy gain
from electron transfer
Glucose slow acceptor
reduction by
A
uncompetitive inhibition
IC
D
2e
-
2e
DOX
ICH2
2e
NUTRIENTS
-
-
ADP
+
ATP
Pi
A red
Biomass
Synthesis
Biomass
Maintenance
Glucose increase
biomass yield (Yx/s)
Figure 5.5 Hypothetical illustration on how glucose may affect the primary flows of
electrons and energy during phenol degradation by C. tropicalis RETL-Cr1 (adapted
from Rittmann and Sàez, 1993) – modified; D = primary electron-donor substrate, DOX =
oxidized electron-donor substrate, A = primary electron-acceptor substrate, Ared = reduced
electron-acceptor substrate, ICH2 = reduced internal cosubstrate, ATP = adenosine triphosphate,
ADP = adenosine diphosphate, and Pi = inorganic phosphate
When an investigation was done to understand the effect of glucose on phenol
utilization and cell density, the following trend was observed. Phenol and glucose was
simultaneously degraded by C. tropicalis RETL-Cr1 when grown in the presence of both
substrates. This result is in good agreement with the results obtained by Wang et al.,
(1996) when a mixture of phenol and glucose was degraded by microorganism of
bacterial origin, i.e. P. putida ATCC 17514 at 28oC, pH 7.2. The typical biodegradation
profile of phenol and utilization of glucose by C. tropicalis RETL-Cr1 at 30oC, pH 6.5
over time is shown in Figure 5.6.
300
12
250
10
200
8
150
6
100
4
50
2
0
0
Biomass conc ( gdw L-1)
Phenol and glucose conc. (mg
L-1)
110
0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18
Time (h)
Figure 5.6 Degradation of phenol by C. tropicalis RETL-Cr1 represented by (Ŷ) and
utilization of glucose (Ɣ) (1mM), growth pattern of C. tropicalis RETL-Cr1 (Ƒ) in
medium containing 3 mM initial phenol concentration at 30oC, pH 6.5.
Figure 5.6 also indicates that there is a sharp decline in glucose utilization in the
first hour of incubation and was depleted completely after 11 hours. This again suggests
that the 1 mM glucose was used as an initial energy source. The decline of phenol
concentration was much more rapid after 9 hours of incubation and reached zero after 15
hours of incubation. Therefore the yeast utilizes glucose at the same rate it utilizes phenol
at 30oC, pH 6.5.
Similar substrate utilization pattern has been also observed in batch cultures for
Pseudomonas putida grown in phenol and glucose (Tarighian et al., 2003). This growth
pattern has been referred to as the simultaneously substrate utilization pattern as
described by Egli (1995). This simultaneous utilization substrate occurs when the
enzymes associated with two substrates “coexist” (Narang, 1998) and has been observed
for both bacteria and yeasts, irrespective of whether growth occurs under aerobic,
anaerobic, mesophilic or thermophilic conditions.
111
On the other hand, it is important to note that sequential utilization of substrate is
a problem in a waste treatment system whereby a mixture of nutrients were available
(some of which are pollutants). Many microorganisms may choose to consume the
preferred substrate and leave the pollutant behind (Goldstein et al., 1985). Since C.
tropicalis RETL-Cr1 had simultaneous substrate utilization pattern, it could serve as a
potential candidate for pollution control in industrial wastewater treatment and
bioremediation (bioaugmentation) that may able to degrade more than one organic
pollutants at the same time.
5.3.1.3 The effect of pH on phenol degradation
For optimum microbial activity in the environment, the preferred range of pH is
between pH6 to 8 (McLelland, 1996). Therefore, it is not surprising to find that most
microorganisms have evolved with pH tolerances within this range (Suthersen, 1999).
Most heterotrophic bacteria and fungi favour a pH near neutrality with fungi being more
tolerant to acidic conditions (Atlas, 1988). Nevertheless there are strains which can thrive
outside this limit. It has been shown that indigenous microorganisms can adapt to lower
pH environments (Weidemeier et al., 1994). The next series of experiments were carried
out to study the effect of pH on phenol degradation by C. tropicalis RETL-Cr1.
Figure 5.7 shows the effect of pH on phenol degradation capability of C.
tropicalis RETL-Cr1 in RM containing 3 mM phenol at 30oC.
0.03
-1
-1
Phenol degradation rate (g L h )
112
0.025
0.025
0.0185
0.02
0.018
0.015
0.01
0.0062
0.0053
0.005
0
4.5
5.5
6.5
7
8
pH
Figure 5.7 The effect of pH on phenol degradation rate of C. tropicalis RETL-Cr1 in
RM containing 3 mM initial phenol concentration at 30oC.
C. tropicalis RETL-Cr1 was observed to have a wide pH range for phenol
degradation from 5.5 up to 7 achieving a phenol degradation rate of between 0.018 g L-1
h-1 and 0.0257 g L-1 h-1. The maximum degradation rate of phenol was 0.0257 g L-1 h-1 at
pH 6.5. However, when the pH values were lower and higher than 6.5, biodegradation
rate was affected significantly. The optimum growth conditions for C. tropicalis RETLCr1 was comparable to Trichosporon cutaneum R57 at 30oC, pH 6.0 (Godjevargova et
al., 2000), and Candida tropicalis at 30oC, pH 6.5-7.2 (Chen et al., 2002; Yan et al.,
2005).
The optimum pH range of C. tropicalis RETL-Cr1 is much narrower compared to
C. tropicalis strain isolated from pristine forest soil at pH range of 3-9 that reported by
Bastos et al., (2000a). The reason for this difference could be our Candida species was
isolated from a different source with a different pH environment. This optimum pH of C.
tropicalis RETL-Cr1 was lower as compared to phenol-degrading microorganisms of
bacterial origin such as Pseudomonas sp., Arthrobacter sp. Bacillus cereus, Citrobacter
113
freundii, Micrococcus agilis and Pseudomonas putida biovar B, Nocardiodes sp.and
Alcaligenes faecalis performed at pH range of 7.0 to 10 (Sarnaik and Kanekar, 1995;
Kanekar et al., 1999; Bastos et al. 2000b). Again this shows that yeast in general prefers
a much lower range of pH as compared to bacteria.
Table 5.3 summarizes the effect of initial pH on degradation of phenol by C.
tropicalis RETL-Cr1 in RM broth containing 3 mM initial phenol concentrations
incubated at 30oC.
114
Table 5.3: Effect of pH on phenol degradation by C. tropicalis RETL-Cr1 at 30oC after
18 hours incubation (RM broth with 3 mM IPC).
Kinetic parameters/Performance
pH
4.5
5.5
6.5
7
8
6.5
13.2
9.765
12.5
4.44
0.2367
0.3141
0.3718 0.3195 0.151
51
29
0.006
0.088
0.072
0.026
0.0003 0.0001
0.003
0.005
0.001
Catechol productivity (g L-1 h-1)
0.00007 0.0001
0.003
0.0007 0.0002
Catmax (g L-1)
0.0013 0.0013
0.0204 0.0046 0.0023
Xmax (gdw L-1)
µ (h-1)
Yx/s (g g-1)
68
Ypc/s (g g-1)
0.017
Ypc/x
(g g-1)
t (Catmax) (h)
18
17
YpccMA/s (g g-1)
0
0.015
YpccMA/x (g g-1)
0
ccMA productivity (g L-1 h-1)
39
38
7
15
0.039
0.036
0
0.0003
0.0013
0.001
0
0
0.0002
0.0006
0.0006
0
ccMAmax (g L-1)
0
0.004
0.011
0.010
0
t (ccMAmax) (h)
0
17
17
0
7
18
Phenol biodegradation rate
(g L-1 h -1)
0.0053 0.0185
Phenol removal efficiency (%)
26.7
98.5
100
98.3
30.5
Incubation time (IT) (h)
18
18
17
18
18
TL (lag time) (h)
4
3
3
3
4
Biodegradation time (BT) (h)
14
15
14
15
14
0.0257 0.0181 0.0062
As shown in Table 5.3, pH6.5 is the most conducive pH for phenol degradation.
during phenol degradation by C. tropicalis RETL-Cr1, again phenol was used mainly for
biomass production instead of the intermediates, catechol and ccMA. This is clearly
shown by the kinetic parameter yields which are as follows: cellular yield g cell dry
weight per g phenol utilized (Yx/s) = 29-68 g g -1, product yield (catechol) per g phenol
115
utilized (Ypc/s) = 0.006 – 0.0072 g g-1 and and product yield (cis,cis-muconic acid) per g
phenol utilized (YpccMA/s) = 0 - 0.039 g g-1.
Careful observation showed that tolerance to acidic conditions fell sharply at pH
below pH 5.5 and similarly at higher alkaline conditions above pH 7.0. At pH 6.5, C.
tropicalis RETL-Cr1 was able to degrade phenol efficiently (100%) at a rate of 0.0257 g
L-1 h-1 with mean generation time of µ 0.3718 h-1, achieving a biomass concentration of
9.765 gdw L-1. However, at a much lower pH (4.5) or higher pH (8.0) the biodegradation
efficiency was only between 27 to 31% with biodegradation rate less than 0.007 g L-1 h-1.
This goes to show that pH 6.5 was the optimum initial pH for maximum phenol
degradation. These results indicate that the variation in pH of the medium may have
changed the ionic form of the active site and changed the activity of the enzyme and then
the reaction rate as suggested by Shuler and Kargi (2002).
The results obtained suggest that the optimum pH for both phenol hydroxylase
(PH) and C1,2D of C. tropicalis RETL-Cr1 could be probably at pH 6.5. This is not
surprising because yeast in general prefers a much lower range of pH as compared to
bacteria. This optimum temperature for PH activity from C. tropicalis RETL-Cr1 was
similar to that of Trichosporon cutaneum reported by Mörtberg and Neujahr, (1987), but
was lower than optimum pH range of 7.6 to 8.0 for C. tropicalis H15 (Krug and Straube,
1986), 7.5 for A. radioresistens (Divari et al., (2003). On the other hand, C1,2D of C.
tropicalis RETL-Cr1 was comparable to optimum pH of 7.5 to 9.6 for C. tropicalis H15
(Krug and Straube, 1986). The reason for these differences could be that C. tropicalis
was isolated from a different source with a different pH environment.
According to Shuler and Kargi, (2002), enzyme activity is only active over a
certain pH range. The pH affects the microorganism’s ability in terms of its cellular
functions, cell membrane transport, and transport protein of microbial cells (Weidemeier
et al., (1994). In addition, the solubility of a compound at different pH values will also be
involved in determining the rate of degradation (Singleton,1994).
116
As shown in Table 5.3, low catechol productivity was achieved and no ccMA was
detected at pH 4.5 and a pH as high as 8.0. This is probably due to the reduction of PH
activity and complete inhibition of C1,2D synthesis of C. tropicalis RETL-Cr1. Figure
5.8 is a hypothetical illustration on how low and high pH affects PH and C1,2D activity
during phenol degradation by C. tropicalis RETL-Cr1.
Lower pH (4.5) and higher
pH (8.0) stop C1,2D
activity
Lower pH (4.5) and
higher pH (8.0) reduce
PH activity
No formation of ccMA
G
L
F
+
+
D
2
2
F
L
Q
R
2 & 2&
F
X
&
& + P
V
L
F
V
L
F
+
+
2
O
R
K
F
H
W
D
F
O2+NADPH+H+
Phenol
+
NADP +
C1,2D
O2
H2O
2H+
& &
+
+
2
+
2
PH
Catechol accumulate
and slow PH activity
Figure 5.8 Hypothetical illustration on how low and high pH may affect PH and C1,2D
activity during phenol degradation by C. tropicalis RETL-Cr1. PH = phenol hydroxylase,
C1,2D = catechol 1,2-dioxygenase, NADP = Nicotinamide adenine dinucleotide
phosphate [Reaction: phenol + O2 + NADPH +H+
(Mörtberg and Neujahr, 1987), catechol + O2
NADP+ + H2O + catechol
ccMA + 2H+(Ngai et al., 1990)]
5.3.1.4 The effect of initial phenol concentration (IPC)
It is known that concentration of a particular chemical is another important factor
to determine biodegradation efficiency. Concentration of a substrate below the threshold
concentrations are not degraded because it is too low to support growth and maintenance
(Boethling and Alexander, 1979). On the other hand, higher concentrations may exhibit
117
toxic effects that reduced biodegradation rates (Alexander, 1985; Cornelissen and Sijm,
1996).
The next series of experiments were carried out to determine the effect of
increased of initial phenol concentration (IPC) from 3 mM to 10 mM (282-941 mg L-1)
on the degree of biodegradation. This was done under batch fermentation in shakeflasks.The initial phenol concentration affects on phenol degradation capability of C.
tropicalis RETL-Cr1 are shown in Table 5.4.
As indicated in Table 5.4, a considerably high phenol degradation could be
achieved at 3 mM initial phenol concentration with an efficiency of 100% and recorded a
degradation rate of 0.0257 g L-1 h-1 at µ = 0.3718 h-1 after incubation of only14 hours.
Initial phenol concentration of 5 mM (0.470 g L-1) was also conducive for phenol
degradation rate but required a slightly longer time. The results obtained here are
comparable to that by Chen et al., (2002) using free cells of C. tropicalis. In this study,
Chen et al., (2002) reported that at 100 mg L-1 which is approximately 1.06 mM, the
phenol removal efficiency was 85% after a long 30 hour incubation at 5 mM initial
phenol concentration. For complete degradation of 5 mM initial phenol concentration, it
required 72 hours of incubation in their case.
118
Table 5.4: The effect of initial phenol concentration (IPC) on phenol degradation by
C. tropicalis RETL-Cr1 at 30oC, pH 6.5 in shake-flask.
Kinetic parameters/Performance
Initial phenol concentration (mM)
3
5
7
10
Xmax (gdw L-1)
9.765
13.098
13.914
4.592
µ (h -1)
0.3718
0.4116
0.3212
0.1557
Y x/s (g g-1)
29
27.4
39.0
24.0
Ypc/s (g g-1)
0.088
0.021
0.013
0.008
0.003
0.0067
0.0003
0.0006
0.003
0.0005
0.00024 0.0008
Ypc/x
(g g-1)
Catechol productivity (g L-1 h-1)
Cat max (g L-1)
0.0204
t (Catmax) (h)
7
0.00698 0.00429 0.00206
14
18
3
YpccMA/s (g g-1)
0.039
0.066
0.017
0.027
YpccMA/x (g g-1)
0.0013
0.0026
0.0004
0.0001
ccMA productivity (g L-1 h-1)
0.0006
0.0015
0.0003
0.00003
ccMAmax (g L-1)
0.011
0.0274
0.00548 0.00045
18
18
t (ccMAmax)
(h)
Rate of phenol degradation (g L-1 h-1)
18
0.0293
18
0.0257
0.0322
Phenol degradation efficiency (%)
100
98
Optimum time for optimum degradation (h)
14
18
18
18
Lag time (TL) (h)
3
5
7
8
56
0.0165
21
Table 5.4 also shows that when the initial phenol concentration was further
increased, a lower values of the specific growth rates were achieved; a phenomenon due
to substrate inhibition as suggested by many researchers (Wang et al., 1996; Ruiz-Ordaz
et al., 1998; Bandyopadhyay et al., 1998; Erhan et al., 2002; Hao et al., 2002). The
specific growth rate µ tends to increase with substrate (Monod-type relationship), but µ
also tends to decrease due to inhibitory effects of substrate (phenol) as its concentration is
119
increased which is consistent with Haldane inhibition kinetics (Yoong et al., 1997). This
means that the C. tropicalis RETL-Cr1 cells could probably be under the inhibitory
influence exhibited by phenol.
It is also seen that the phenol degradation rate increased initially with phenol
concentration and decreased in the degradation rate after phenol concentration reached 5
mM) (470 mg L-1). This means that phenol inhibits the C. tropicalis RETL-Cr1 at
concentration level higher than 5 mM of initial phenol concentration. Free cells of C.
tropicalis RETL-Cr1 had poor degradation efficiency when IPC was at 7 mM-10 mM
(659-941 mg L-1) of only between 21-56% removal efficiency were achieved at the end
of degradation (18h). Free cells of C. tropicalis RETL-Cr1 had lower degradation rate
when IPC (So) was increased to 7 mM and 10 mM (659-941 mg L-1).
Besides being a substrate, phenol could also acts an inhibitor (Neujahr and
Kjellén, 1978; Mörtberg and Neujahr, 1987). Self-inhibition, in which high concentration
of a substrate inhibit its own degradation has been reported by Godrej and Sherrard,
(1988) and Sàez and Rittmann, (1991). These authors suggested that self inhibitory
substrates probably hinder energy and electron flows at several locations and do not only
inhibit their own enzyme-catalyzed transformation.
It is also seen that during phenol degradation by C. tropicalis RETL-Cr1, phenol
instead of the intermediates catechol and ccMA was utilized mainly for biomass
production. This is clearly shown by the kinetic parameter yields which are as follows:
cellular yield g cell dry weight per g phenol utilized (Yx/s) = 24-39 g g -1, product yield
(catechol) per g phenol utilized (Ypc/s) = 0.0008 – 0.003 g g-1 and product yield (cis,cismuconic acid) per g phenol utilized (YpccMA/s) = 0.017 - 0.066 g g-1.
Once again, under growing cell conditions, catechol productivity was highest at 3
mM initial phenol concentration but decreased with increasing phenol concentration.
ccMA productivity increases up to 5 mM initial phenol concentration but decreased with
further increased in phenol concentrations. This characteristic was again probably due to
120
inhibition exerted by phenol.
In the present study, phenol as self-inhibitory substrate may probably have
exerted both direct inhibition which affects the key enzymes during degradation by C.
tropicalis RETL-Cr1 as illustrated in Figure 5.9 and indirect inhibition which affects the
electron and energy flows as illustrated in Figure 5.10.
Phenol at IPC above 5mM
reduce PH activity
Phenol at IPC above 5mM
reduce C1,2D activity
+ H2O
G
L
F
+
+
D
2
2
F
L
Q
R
2 & 2&
F
X
&
& + P
V
L
F
V
L
F
NADP
+
+
+
2
O
R
K
F
H
W
D
F
O2+ NADPH + H+
Phenol
C1,2D
O2
2H+
& &
+
+
2
+
2
PH
Figure 5.9 Hypothetical illustration on how high phenol concentration may affect PH and
C1,2D activity during phenol degradation by C. tropicalis RETL-Cr1. PH = phenol
hydroxylase, C1,2D = catechol 1,2-dioxygenase, NADPH = Nicotinamide adenine
dinucleotide phosphate [Reaction: phenol + O2 + NADPH +H+
catechol (Mörtberg and Neujahr, 1987)), catechol + O2
1990).
NADP+ + H2O +
ccMA + 2H+ (Ngai et al.,
121
Decouplers stop or reduce
energy gain from electron
transfer
Phenol as self- inhibitor at
IPC above 5 mM slow or
stop donor oxidation
Phenol as selfinhibitor at IPC above
5 mM slow or stop
A
acceptor reduction
IC
D
2e
-
2e
ICH2
DOX
2e
NUTRIENTS
-
-
ADP
+
Pi
ATP
A red
Biomass
Synthesis
Biomass
Maintenance
Figure 5.10 Hypothetical illustration on how high phenol concentration may affect the
primary flows of electrons and energy during phenol degradation by C. tropicalis RETLCr1 (adapted from Rittmann and Sàez, 1993) – modified; D = primary electron-donor
substrate, DOX = oxidized electron-donor substrate, A = primary electron-acceptor substrate, Ared
= reduced electron-acceptor substrate, ICH2 = reduced internal cosubstrate, ATP = adenosine
triphosphate, ADP = adenosine diphosphate, and Pi = inorganic phosphate.
The inhibitory effect of phenol on C. tropicalis RETL-Cr1 observed at
concentration of 5 mM (470 mg L-1) is in good agreement to that of Bandyopadhyay et
al., (1998). According to Bandyopadhyay et al., (1998) inhibition effects of phenol as
substrate have become predominant above concentration of 500 mg L-1 (5 mM) when
microorganism of bacterial origin, Pseudomonas putida MTCC 1194 was used.
Interestingly, similar results were reported by Kotturi et al., (1991); Sàez and Rittmann
(1993); Dikshitulu et al., (1993). They reported that biodegradation of phenol as an
individual compound has been observed to follow Monod’s kinetics at initial
concentration below 4.2 mM (400 mg L-1) but became inhibitory above this
concentration. However, the observed phenol inhibitory level of 5 mM (470 mg L-1 )
exhibited by C. tropicalis RETL-Cr1 was 2-fold higher than that reported by Léonard et
122
al., 1999; Chai et al.,(2004) and Yoong et al., (2004), and 19-fold higher than that
reported by Monteiro et al., (2000); Mahadevaswamy et al., (2004). The observed phenol
inhibitory levels observed in other microorganisms reported by previous researchers are
shown in Table 2.5 (Section 2.6.3.1) of chapter 2.
It can be hypothesized that substrate inhibition happened in C. tropicalis RETLCr1 during phenol degradation. Different toxic organic compounds have been reported to
have deleterious effects on numerous sites within cells. Depending on their physical and
chemical properties, and different types have shown to increase membrane fluidity (Isken
and de Bont, 1998), decrease ATP synthesis (Loffhagen et al., 1995), and modify or
denature biomolecules (Tamarit et al., 1998) in bacteria. The toxicity of phenol at a high
concentration level could inhibit the related metabolism of biodegradation of phenol
resulting in lower removal efficiency by free cells of C. tropicalis as suggested by Chen
et al., (2002).
Substrate inhibition is a characteristic of phenol metabolism being a toxic
substrate as suggested by Santos and Linardi, (2004) for different microorganisms at
different concentration levels. For instance, Chung et al., (2003) reported that
Pseudomonas putida cannot tolerate the toxicity of phenol at high concentrations of
between 8.5 –10.6 mM (800-1000 mg L-1). The toxicity of phenol at high concentrations
level above 1,500 mg L-1 could inhibit the related metabolism of degradation resulting in
a lower efficiency of free cells of C. tropicalis to degrade phenol (Chen et al., 2002).
Besides results from laboratory data, phenol toxicity studies in phenol-contaminated sites
also have shown that increasing phenol concentrations appeared to decrease the overall
phenol biodegradation of bacteria (Dean-Ross, 1989).
The toxicity of aromatic compounds is frequently attributed to the disruption of
membrane structure by hydrophobic interactions with the lipid bilayer structure caused
by the lipophilic nature of such compounds (Sikkema et al., 1994). Phenol toxicity is
always associated with loss of cytoplasmic membrane integrity causing in disruption of
energy transduction, disturbance of membrane barrier function, inhibition of membrane
123
protein function, and subsequent cell death (Keweloh et al., 1990; Heipieper et al., 1991,
1992). However, according to Leonard and Lindley (1999) phenol hydroxylase (PH) is
the major site for phenol inhibition and this enzyme is sensitive to hydrophobic stress.
Leonard and Lindley (1999) further suggested that subcellular location of PH would be
the cell membrane, thereby avoiding penetration of phenol into the cytosol. In another
study, Ettayebi et al., (2003) reported that cell fusion was pronounced in free cells of
Candida tropicalis. In this study by Ettayebi et al., (2003) found that free cells were
aggregated to lower contact points with toxic phenols. These results by Ettayebi et al.,
(2003) are in agreement with the investigation made by Zache and Rehm, (1989).
Besides, substrate inhibition caused by phenol, accumulation of metabolic
intermediates are also responsible for lower phenol degradation efficiency as suggested
by many researchers : Wang et al., (1979), Kleþka and Gibson, (1981), Bartels et al.,
(1984), Allsop et al., (1993); Wang and Loh (1999) and Komarkova et al., (2003). This
happens when the aromatic ring is cleaved, the intermediate could become more reactive
and compete with phenol in the series of enzymatic reactions, resulting in strong
inhibition on phenol consumption (Wang and Loh, 1999).
Divari et al., (2003) suggested that accumulation of end product such as catechol
might inactivate the enzyme, phenol hydroxylase and therefore the presence of
downstream enzyme, C1,2D is necessary to shift the equilibrium towards the
detoxification reactions. It is known that catechol is considered more toxic than phenol.
Phenol inhibition has been observed in the presence of catechol after reaching
concentration of 460 mg L-1 (Erhan et al., 2002) In this study the maximum concentration
of catechol detected was only 20.4 mg L-1 which is significantly lower to that reported by
Erhan et al., (2002), and furthermore catechol was depleted completely at the end of the
incubation period. Therefore in this case, catechol might not be responsible for inhibition
on C. tropicalis RETL-Cr1. The intermediates responsible could be ccMA and other
unidentified open-ring products. In our case ccMA was seemed to accumulate in the
medium at the end of each biodegradation process.
124
Catabolism of phenol was confirmed with the detection of two intermediates; the
first one being catechol and the second intermediate cis,cis-muconic acid (ccMA) which
is the breakdown of catechol. As shown in Figure 5.11, the two intermediates catechol
and cis,cis-muconic acid were analyzed after 18 hours with the starting phenol
concentrations varying from 3-10 mM. This correlation of phenol degradation and the
emergence of two substrates are studied here. It was observed that catechol maximum
concentration (Catmax) of 20 mg L-1 (0.18 mM) was the highest when the lowest initial
phenol concentration (IPC) of 282 mg L-1 (3 mM) was used. Catmax decreased with
increasing IPC. Meanwhile, cis,cis-muconic acid maximum concentration (ccMAmax) of
27.4 mg L-1 (0.19 mM) was the highest when 5 mM IPC was used. ccMAmax was
increased initially up to 5mM IPC but decreased with further increases in IPC. This
means that at much higher IPC however, negligible units of both catechol and ccMA
were produced which may indicate that substrate inhibition might have taken place here
as discussed before. Similarly the phenol removal efficiency decreased with increasing
IPC.
catechol and cis,cis-muconic
acid conc. (mg L-1)
25
80
20
60
15
40
10
20
5
0
0
3
5
7
phenol removal effciency (%)
100
30
10
Initial phenol concentration(mM)
Figure 5.11 Concentration of intermediates; catechol (Ŷ) and cis,cis-muconic acid (Ƒ)
and phenol removal efficiency (ż) at various IPC by C. tropicalis RETL-Cr1.
125
Since 5 mM IPC appeared to be relatively non-inhibitory to phenol degradation
this concentration was chosen for the next experiment. Figure 5.12 illustrates the rate of
phenol degradation and production of intermediates i.e., catechol and cis,cis-muconic
acid at IPC of 5 mM (470 mg L-1) under batch fermentation in shake-flask.
phenol conc. (mg L-1)
400
25
350
20
300
250
15
200
10
150
100
5
50
0
catechol and ccMA conc.
(mg L-1)
30
450
0
0
1
2
3
4
5
6
7
8
9 10 11 12 13 14 15 16 17 18
Time (h)
Figure 5.12 Degradation of phenol (Ŷ) and production of intermediates; catechol(Ɣ) and
cis,cis-muconic acid (ccMA) (Ÿ) by C. tropicalis RETL-Cr1 against time at IPC of 5
mM in Ramsay medium at 30oC, pH 6.5 in shake-flask.
As shown in Figure 5.12, conversion of catechol and cis,cis-muconic acid were
not simultaneous. Catechol was formed during the early stages of phenol degradation
(after 2 hours) suggesting phenol hydroxylase activity reached maximum levels at the
beginning of the exponential phase as suggested by Fialovà et al., (2004). On the other
hand, cis,cis-muconic acid was formed during the later stage of the biodegradation
process (after 6 hours).
The production of catechol reached its maximum at a particular time and
subsequently disappeared near the time when phenol was further converted to ccMA.
Catechol was produced after two hours, maximized at 7 mg L-1 after 14 hours and
subsequently depleted at the end of cultivation. Interestingly, a similar trend of catechol
production was also observed in immobilized P. putida that was able to degrade phenol at
126
concentration between 225 mg L-1 to 450 mg L-1 at optimum temperature at 30oC, pH 6.8
(Chung et al., 2003). A conclusion that can be made from these two experiments is that
temperature and pH plays an important role in determining the behavior of catechol
irrespective of different microbial species and treatment.
On the other hand, cis,cis-muconic acid was only formed after 6 hours of the
degradation process and increased exponentially as phenol concentration decreased
exponentially and maximized at 27.4 mg L-1 at the end of the cultivation and seemed to
accumulate in the medium.
5.3.2
Comparison of phenol degradation in shake-flask and bioreactor
The ability to degrade phenol in batch culture (shake-flask and bioreactor) by free
cells of C. tropicalis RETL-Cr1 containing 3 mM initial phenol concentration was
compared. IPC of 3 mM was chosen because at this concentration, 100% phenol removal
efficiency was achieved. The comparative profiles of shake-flask (100 mL volume in 250
mL flask, with agitation speed of 200 rpm) and bioreactor of batch fermentation (500 mL
in 2L bioreactor with agitation speed of 200 rpm, airflow of 1L/min) are shown in Table
5.5.
Complete degradation of phenol at 3 mM was achieved after 14 hours incubation
using batch cultures (shake-flask with an agitation speed of 200 rpm) at a rate of 0.0257
g L-1 h-1 with specific growth rate (µ) = 0.3718 h-1. The time required to biodegrade
phenol completely supplied to the bioreactor was shortened to 10 hours. Hence, the
degradation rate was increased to 0.0395 g L-1 h-1 at µ = 0.5381 h-1 with bioreactor. There
was a 35% increase in phenol degradation efficiency or 1.5-fold increased in
biodegradation rate. This performance was comparable to that reported by Kang and Park
(1997) when phenol was degraded at a faster rate in a bioreactor than in flasks culture
using mixed Pseudomonas sp. and was increased by 1.8-fold.
127
Table 5.5: Comparison of phenol degradation performance in shake-flask and
bioreactor with an IPC of 3 mM of C. tropicalis at 30oC, pH 6.5.
Kinetic parameters/ Performance
Phenol biodegradation
Shake-flask
Bioreactor
Xmax (gdw L-1)
9.765
9.322
µ (h-1)
0.3718
0.5381
Rate of phenol degradation (g L-1 h-1)
0.0257
0.0395
a) catechol (g L-1)
0.0204
0.0043
b) ccMA (g L-1)
0.011
0.00983
Yx/s (g g-1)
29
28
Ypc/s (g g-1)
0.088
0.020
0.003
0.0006
t(catmax) (h)
7
8
Catechol productivity (g L-1 h-1)
0.003
0.0005
YpccMA/s (g g-1)
0.039
0.036
YpccMA/x (g g-1)
0.0013
0.0013
t(ccMAmax) (h)
18
18
ccMA productivity (g L-1 h-1)
0.0006
0.00055
Biodegradation time (h)
14
10
Rate of degradation % of phenol
degradation improvement
100
135
Degradation improvement to
Shake-flask culture (x times)
-
1.5
Production of byproducts
Ypc/x
(g g-1)
The increase in rate of degradation may be due to the improved growth conditions
as provided by proper mixing and aeration in the bioreactor. It is known that sufficient
supply of oxygen is a critical factor in aerobic bioprocesses. Aerobic biodegradation
occurs in the presence of molecular oxygen which is the electron acceptor. The Biostat®
B. 2L used in the phenol degradation fermentation provides both agitation and aeration.
128
According to Armenante, (1993) agitation in the bioreactor serves two purposes. The first
one is to achieve homogeneous mixing and to disperse dissolved nutrients and biomass in
the reactor provided by the impeller. Secondly, to disperse the air by breaking up the
bubbles produced by the spargers to increase gas-liquid interfacial area and promote
oxygen transfer. According to Ju and Sundarajan, (1995), the retention of the intercepted
cells of yeast by the bubbles was expected to be the primary factors responsible for cell
accumulation in the gas-liquid interface so homogenous mixing is required here.
Bartholomew et al., (1950) proposed that oxygen may be transferred directly from
the gas bubble to respiring microorganisms accumulated at the gas/liquid interface.
Leonard and Lindley, (1999) observed that approximately 40% of the oxygen
consumption in cells of Ralstonia eutropha catabolizing phenol attributed to respiratory
consumption and the remainder being directly associated with phenol hydoxylase and
catechol 2,3-dioxygenase activities. The oxygen respiration is measured in terms of
oxygen consumption per unit of phenol utilized. It has been reported that oxygen
consumption by Trichosporon cutaneum cells was 2 µmol O2/µmol phenol (Gaal and
Neujahr, 1979).
5.3.3 Time course of phenol degradation by C. tropicalis RETL-Cr1 under
optimum condition
Figure 5.13 shows phenol degradation against time and biomass concentration of
C. tropicalis RETL-Cr1 when grown on phenol at IPC of 3 mM, with optimum condition
at 30oC, pH 6.5 and in the absence of glucose under batch fermentation in bioreactor.
300
12
250
10
200
8
150
6
100
4
50
2
0
0
Biomass conc. (gdw L-1)
Phenol conc. (mg L-1)
129
0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18
Time (h)
Figure 5.13 Degradation of phenol (Ŷ) by C. tropicalis RETL-Cr1 against time in RM
broth with initial concentration of phenol of 3 mM in the absence of glucose at 30oC, pH
6.5. Biomass concentration (¨) of C. tropicalis RETL-Cr1.
As shown in Figure 5.13, C. tropicalis RETL-Cr1 also degrade 3 mM phenol
efficiently in RM broth without addition of glucose at optimum condition (30oC, pH 6.5).
The decline in phenol concentration was more rapid during growth and complete
depletion was observed after 14 hours of incubation. A lag period (TL) of 3 hours was
observed in the time course of phenol degradation. The rate of phenol degradation
coincides with the increase in growth of C. tropicalis RETL-Cr1. This may suggest that
phenol degradation can still take place even if there is no additional carbon source added.
Hence, phenol may be used as sole carbon source in place of glucose. Growth profile of
C. tropicalis RETL-Cr1 in the absence of glucose showed no apparent lag phase which
may indicate that C. tropicalis RETL-Cr1 may adapt to phenol almost immediately.
When phenol was completely exhausted, growth declined. C. tropicalis RETL-Cr1 was
able to remove phenol completely at a u value of 0.3718 h-1 after 10 hours incubation in a
bioreactor culture.
130
5.4
Conclusions
From the results obtained in this study allow the following conclusions can be
made.
C. tropicalis RETL-Cr1 degrade phenol very efficiently in either medium either
(in the presence or absence of glucose) at an initial phenol concentration of 3 mM.
Phenol degradation rate and biomass concentration were affected by both temperature
and pH. This yeast was able to degrade phenol at temperature range between 30oC to
37oC and at wide range of pH values from 5.5 to 7.0. The optimal condition for growth
and phenol degradation was at 30oC, pH 6.5 in medium without glucose supplement.
Under batch condition, phenol was utilized by C. tropicalis RETL-Cr1 mainly for
biomass production instead of the intermediates, catechol and ccMA. This is clearly
shown by the kinetic parameter yields at optimum conditions as follows: Yx/s = 29 g g -1,
Ypc/s = 0.088 g g-1 and YpccMA/s = 0.039 g g-1. It can be clearly seen that the batch cultures
(shake-flask) was characterized by high biomass yield (Y x/s) but with low product yields
i.e. (catechol) yields (Ypc/s and Ypc/x) and ccMA (YccMA/s and YccMA/x). This relatively
high biomass yield could be due to the high metabolic efficiency of C. tropicalis RETLCr1 and also to high carbon/oxygen ration in phenol as suggested by Hao et al., (2002).
It was observed that the specific growth rates of C. tropicalis RETL-Cr1
decreased at higher concentrations of phenol suggesting that substrate inhibition may
play some role on the rate of phenol degradation. The effect of inhibition was observed at
phenol concentration above 5 mM (470 mg L-1).
Phenol was degraded at a faster rate in a bioreactor than in shake-flask culture due
to improved growth conditions offering proper mixing and aeration. The highest removal
efficiency in shake-flask cultures was 100% with degradation rate of 0.0257 g L-1 h-1 at µ
= 0.3718 h-1 after 14 h cultivation in medium containing 3 mM phenol. The degradation
rate of phenol was improved by 1.5-fold in a bioreactor where the incubation time was
131
shortened to 10 hours.
The catabolism of phenol was confirmed with the detection of intermediary
products, catechol and cis,cis-muconic acid. Higher catmax, catechol productivity,
ccMAmax and ccMA productivity were achieved at temperature range from 30oC to 37oC
and pH from 5.5 to 7.0 suggesting that at these physiological conditions were optimum
for phenol hydroxylase (PH) and catechol 1,2-dioxygenase (C1,2D) activities.
Temperature of above 37oC may have caused denaturation of C1,2D of C. tropicalis
RETL-Cr1 and hence catechol was not converted to ccMA. The catechol may have
accumulated in the medium and exerted inhibition on PH which is responsible for phenol
conversion to catechol.
In this optimization of phenol degradation by C. tropicalis RETL-Cr1, the effect
of different process variables like pH, temperature and presence or absence of glucose on
phenol biodegradation has been determined and optimum process conditions was
established for further investigations.
Due to the possibility of substrate inhibition, an attempt to overcome this problem
was made by improving the degradation capability of C. tropicalis RETL-Cr1 using fedbatch system which will be discussed in Chapter 6.
132
CHAPTER 6
IMPROVEMENT OF PHENOL BIODEGRADATION BY FED-BATCH
CULTURE OF Candida tropicalis RETL-Cr1
6.1
Introduction
Conventional batch fermentations of phenol degradation are limited by the low
initial phenol concentrations required to prevent complete inhibition of microbial activity.
Problems such as substrate inhibition, low cell concentration, glucose effect, catabolite
repression, and high viscosity of the culture broth are common in batch processes
(Andrews, 1968; Shuler and Kargi, 2002). Therefore, for practical reasons, certain
continuous operations have been replaced by fed-batch processes (Schügerl, 1987). In the
broad sense, “fed-batch” is defined as a technique in microbial processes where one or
more limiting nutrients are supplied to the bioreactor (in some cases, all nutrients are
gradually fed into the bioreactor) during cultivation and in which the products remain in
the containment until the end of the run (Besli et al., 1995). Fed-batch culture is a
modification of batch operation which is between batch and continuous fermentation
(Yamane and Shimizu, 1984).
Phenol biodegradation operated under this technique involves feeding of
substrates and nutrients into the bioreactor either intermittently (fix mode) or
continuously. This intermittent feeding technique is to achieve an increase degradation
133
rate by maintaining constant substrate concentration and specific growth rate and at the
same time maintaining low levels of phenol in the bioreactor. This condition able to
minimize deleterious effects of substrate inhibition and catabolite repression (d’Anjou
and Daugulis, 2000).
In this study, fed-batch fermentation using free cell of C. tropicalis RETL-Cr1
with an intermittent feeding strategy was attempted to remedy the problem of inhibition
encountered during batch culture as discussed in Chapter 5, and also to determine the
capability of C. tropicalis RETL-Cr1 to degrade phenol at high concentration conditions
using fed-batch system. This fed-batch fermentation technique is a progressive addition
of limiting substrate (phenol) from a lower to a very high concentration. This condition is
to build up tolerance towards phenol in C. tropicalis RETL-Cr1. During intermittent
feeding in fed-batch culture the degradation rate of phenol will improves substrate
concentration in order to prevent substrate inhibitory or catabolite repression.
6.2
Materials and Methods
6.2.1
Batch and Fed-batch Experimental Design
The experimental set-up of the fermenter for batch system was carried out under
aerobic condition in a bench-scale bioreactor (Biostat B 2L Model B. Braun) with a 0.5 L
working volume. The vessel have internal concave bottom with outer thermostat jacket.
The agitation of culture broth was achieved by using of two 0.05 m diameter of 6 bladed
rushton turbine impellers. The top plate of this vessel is made of stainless steel and
consisted of seven ports for pH and oxygen electrodes, inoculation, sampling,
temperature opening sensor, air sparger, acid, alkali and antifoam inlets.
During the fermentation process the profile of culture pH was measured by using
Ingold pH probe. This probe was calibrated using pH buffer 7.0 and 4.0 prior to
sterilization at 121oC for 15 minutes. The initial pH of all media was adjusted to 6.5. The
134
culture pH was not controlled but recorded continuously during the fermentation process.
Silicone oil (3% v/v) was used as an antifoam. The temperature of the fermenter was
controlled at 30oC. In all experiments, agitation speed was set at 200 rpm and the
dissolved oxygen tension in the culture broth was controlled by controlling the air flow
rate. The airflow was set at 1L/min. The calibration of oxygen probe was conducted after
sterilization by sparging the bioreactor with nitrogen free oxygen for 10 minutes.
The fed-batch culture was set up as shown in Figure 6.1. Fed-batch experiments
were carried out with a 1.5 L working volume under aerobic condition (Figure 6.1). The
condition and procedures of the fermentation process were conducted similar to as
previously described in a batch culture experiment.
135
Gas outlet
Filter
Peristaltic pump
Feed: Ramsay Medium
+ Phenol
(100 mL/feed)
Anti-foam
Sampling Port
Impellers
Fermenter
Temperature Controller
Figure 6.1 Fermenter Set-Up for Fed-batch Culture
The experiments were started batch-wise for up to 10 h. Ramsay Medium (RM)
(0.5 L) containing 3 mM phenol was placed in the reactor and inoculated with 50 mL
overnight yeast cultures. Fed-batch feeding was initiated at t = 10 h. Each feed of fresh
medium added were 100 mL containing varying concentration of phenol from 4 mM up
to 820 mM as a sole carbon source. Five mL samples were taken periodically throughout
the operation for determination of residual phenol, catechol, cis,cis-muconic acid and
optical density.
136
6.2.2
Sample Analysis
6.2.2.1 Determination of biomass concentration
The experimental procedures are presented in section 3.5.1
6.2.2.2 Determination of average phenol degradation rate
The experimental procedures are presented in section 3.5.3
6.2.2.3 Determination of phenol, catechol and cis,cis muconic acid
The experimental procedures are presented in section 3.5.5
6.2.3
Microscopy observation
The yeast cultures were subjected to periodical observations through microscopic
observation and spread plate to ensure they were not contaminated with undesirable
species. None of the yeast cultures were contaminated due to aseptic technique and the
antiseptic nature of phenol.
6.3
Results and Discussion
During batch culture it was observed that the specific growth rates of C. tropicalis
RETL-Cr1 was affected significantly at higher concentrations of phenol suggesting that
substrate inhibition may play some role on the rate of phenol degradation. The effect of
inhibition was observed at initial phenol concentration (IPC) above 5 mM (470 mg L-1)
which has been discussed in Chapter 5. To overcome the growth inhibitory substrate
problems observed during phenol biodegradation by C. tropicalis RETL-Cr1 in batch
137
cultures, fed-batch approach was attempted as suggested by Reardon et al., (2000) for
better efficiency of phenol biodegradation.
6.3.1 Batch fermentation
Phenol biodegradation by C. tropicalis RETL-Cr1 was initiated with batch mode
for 10 h at 30oC, initial pH 6.5 in Ramsay medium containing (RM) 282 mg L-1 (3 mM)
phenol as substrate (Figure 6.2). The batch culture was converted to fed-batch when X =
Xmax and S = 0. In this experiment the fed-batch culture was initiated at t = 10 h.
Fed-batch initiated at t10
250
10
200
8
150
6
-1
Phenol conc. (g L )
12
100
4
S= 0
50
2
0
0
0
1 2 3
4 5 6
7 8
Biomass (gdw L-1), Cat &
ccMA (mg L-1) Conc.
X = X max
300
9 10 11 12 13 14 15 16 17 18
Time (h)
Figure 6.2 Time course of phenol (Ŷ) degradation in batch culture by C. tropicalis in RM
at 30oC, initial pH 6.5. Symbols: (ż) Catmax (ż) ccMAmax (Ÿ) and (Ƒ) Biomass
concentration of C. tropicalis RETL-Cr1.
Figure 6.2 shows the profile of phenol degradation and production of catechol and
ccMA during batch mode by C. tropicalis RETL-Cr1. Catechol was produced after 3
hours and catmax (4.3 mg L-1) was achieved after 8 hours incubation and depleted near
zero an hour later. On the other hand, ccMA was produced after 7 hours and ccMAmax
138
(8.07 mg L-1) was achieved after 9 hours incubation.
6.3.2
Fed-batch fermentation
The use of fed-batch was to improve capability of C. tropicalis RETL-Cr1 to
degrade phenol at high level concentration. The feed was initiated with a low
concentration of phenol as the limiting substrate at 4 mM (376 mg L-1) to allow better
adaptation of the yeast cells to phenol. In this system the growth rate and substrate
concentration were maintained at suitable levels throughout the cultivation process. The
fed-batch fermentation had permitted extension of the operating time with high cell
concentrations of C. tropicalis RETL-Cr1 achieved. Linear increase of degradation rate
was observed and reached a maximum biomass concentration of 61 g L-1 at the end of
biodegradation process.
Table 6.1 shows the kinetic parameters and the overall performance of phenol
degradation in fed-batch system by free cells of C. tropicalis RETL-Cr1. As shown in
Table 6.1, phenol degradation efficiency achieved was between 96 % -100%. The highest
phenol degradation rate of 2.39 g L-1 h-1 was achieved when the feed concentration was at
820 mM (77.2 g L-1) but with a high residual phenol concentration of 2.97 g L-1.
Therefore, in terms of degradation efficiency, C. tropicalis RETL-Cr1 was able to
degrade phenol more effectively when the feed concentration was 28.2 g L-1 (300 mM) as
there was 91-fold fall of residual phenol as compared to the feed concentration at 820
mM. The phenol degradation rate of 1.76 g L-1 h-1 achieved at 300 mM feed was still
reasonably high.
The fed-batch system had successfully avoided the effect of substrate inhibition
encountered during phenol degradation using batch system. Phenol as the substrate did
not cause an inhibitory response as anticipated. Fed-batch culture has been used in
circumstances where component of a medium need to be maintained at low level to
prevent toxic effect to the organism (Stanbury and Whitaker, 1984). The system has
139
allowed the addition of a very high phenol concentration up to 820 mM (77.2 g L-1) into
the medium without affecting phenol degradation efficiency of C. tropicalis RETL-Cr1.
Table 6.1 Kinetic parameters/Performance of fed-batch fermentation of phenol
degradation by C. tropicalis RETL-Cr1.
Parameters/Performance
Conc. (g L-1)
0.376
0.941
Intermittent Feed
1.882
4.701
28.2
77.17
Conc. (mM)
4
10
20
50
300
820
Operational time (h)
10
27
37
61
105
680
Opt.degradation time (h)
8
2
2
3
16
31
Xmax (g L-1)
9.90
12.8
13.5
30.3
59.3
61.9
Residual phenol (g L-1)
0.0
0.00078
Rate of phenol –
degradation (g L-1 h-1)
0.05
0.47
0.93
1.56
1.76
2.39
Degradation efficiency (%)
100
100
98.4
99.7
99.9
96.2
Yx/s (g g-1)
1.54
3.70
3.90
4.48
1.77
0.71
0.03166 0.01475 0.03279
2.9668
Catechol concentration(g L-1)
0.00080 0.00084 0.00078 0.00073 0.0
0.0
Ypc/s (g g-1)
2.1x10-3 8.9x10-4 4.1x10-4 1.6x10-4
0.0
0.0
Ypc/x (g g-1)
1.4x10-3 2.4x10-4 1.9x10-4 3.5x10-5
0.0
0.0
Catechol productivity
(g L-1 h-1)
5.3x10-5 3.1x10-5 2.1x10-5 1.2x10-5
0.0
0.0
Cis,cis-muconic acidconcentration (g L-1)
0.0
0.0
0.0
0.00092
0.00282 0.00927
YpccMA/s (g g-1)
0.0
0.0
0.0
2.0x10-4
1.0x10-4
1.3x10-4
YpccMA/x (g g-1)
0.0
0.0
0.0
4.4x10-5
5.6x10-5
1.8x10-4
ccMA productivity(g L-1 h-1)
0.0
0.0
0.0
1.5x10-5
2.6x10-5 1.4x10-5
140
Table 6.1 also shows that catechol was fully utilized by C. tropicalis RETL-Cr1
and the onset of accumulation of ccMA was observed after feed solution containing 4.7 g
L-1 phenol was added and increased until the end of the fermentation period. This
increase in accumulation and level-off on the biomass concentration could be due to
ccMA inhibitory effect either direct or indirectly on C. tropicalis RETL-Cr1. However,
the degradation rate was still increasing with an increase in residual phenol concentration.
This behavior could be an indication of tolerance of the cells of C. tropicalis RETL-Cr1
towards higher phenol concentration. This tolerance could have build-up during the
addition of constant intermittent feeding from a low concentration of phenol to a gradual
increased of higher phenol concentration. During this acclimation time, physiological
changes in the metabolic system of cells took place in response to exposure to a new
environment (Bali and Sengül, 2002) which involved changes in regulation and
production of enzymes, cell sizes and composition and in genetic characteristics as
suggested by Alexander, (1994) and Moustafa El-Sayed (2003).
Phenol degradation rate can be correlated to both growth and phenol
concentration. Therefore, in order to improve phenol degradation rate in fed-batch, it
required a constant residual phenol concentration in the reactor for growth without
exceeding the critical concentration which would cause growth inhibition as suggested by
Léonard et al., (1999). According to Cruickshank et al., (2000), more frequent feedings
for optimal fed-batch feeding strategies could lead to a larger amount of phenol
consumed. Similar strategies have been carried out in this study. High phenol degradation
rate achieved could be specifically due to frequent addition of fixed volume of fresh feed
RM (100 mL) that has been able to maintain the concentration of phenol below inhibitory
levels of C. tropicalis RETL-Cr1 (470 mg L-1) thus substrate inhibition during
fermentation process was avoided. Similar observation was reported by Vrionis et al.,
(2002) when the aqueous phase phenol concentration was maintained in between 400-460
mg L-1 which was below the inhibitory level of (500 mg L-1 ) during fed-batch
fermentation of P. putida on phenol.
141
In another report, Pamment et al., (1978) suggested that the acclimation time
involved the translation of new genetic information resulting in a shift in the
concentrations of ribonuclease and protein molecules inside the cells. As an example,
adaptation of Pseudomonas putida to phenol prior degradation has reported an increase in
phenol degradation rate by 2-fold (Gonzalez et al., 2001b). Similarly, free cells of C.
tropicalis RETL-Cr1 was adapted to lower concentration of phenol at 3 mM (282 mg L-1)
during batch fermentation prior to fed-batch with a gradual increase of higher phenol
concentration from 4 mM to 820 mM (0.376 g L-1 - 77.2 g L-1). There was a 273-fold
increase of phenol concentration in the fed-batch as compared to batch system. Hence,
gradual increase in phenol in fed-batch could help C. tropicalis RETL-Cr1 adapt in order
to degrade phenol at very high concentration.
Figure 6.3 shows a typical time course of phenol degradation capability by C.
tropicalis RETL-Cr1 in fed-batch fermentation. The results achieved certainly proved the
ability of C. tropicalis RETL-Cr1 to degrade phenol at high concentration using fedbatch system. The maximum degradation rate in the fed-batch culture was up to 2.3 g L-1
h-1 at the end of the fermentation process.
0.00003
0.000025
2
0.00002
1.5
Start feeding
0.000015
1
0.00001
0.5
0.000005
0
Cat & ccMA productivity (g L-1 h-1)
Phenol degradation rate (g L-1 h-1)
2.5
0
10
18
22
24
27
29
37
61
76
105 179 496 680
Time (h)
Figure 6.3 Time course of phenol degradation in fed-batch fermentation by C. tropicalis
RETL-Cr1 in RM at 30oC, initial pH 6.5. Symbols: (Ŷ) phenol degradation rate, catechol
productivity (Ɣ) and (Ÿ) cis,cis-muconic acid productivity.
142
As clearly shown in Figure 6.3, fed-batch was characterized by high phenol
degradation rate but with low catechol and cis,cis-muconic productivity as compared to
batch system. The low product yield was probably due to the utilization of catechol as a
product of phenol degradation which occurred simultaneously with phenol utilization and
thus catechol was detected only at low concentration. On the other hand, ccMA product
yield from catechol was observed to be just as low as it is difficult to utilize catechol
probably due to its toxicity as suggested by Maxwell et al., (1986), Gomi and Horiguchi,
(1986) and Yoshikawa et al., (1990). For comparison, ccMA productivity by C. tropicalis
RETL-Cr1 both from batch and fed-batch were lower by 1330-fold as compared to that
reported by Bang and Choi (1995).
As shown in Table 6.1, there was an increase of cell biomass yield (Yx/s) until
addition of solution feed containing 4.7 g L-1 (50 mM) phenol and decreased after further
increase of addition of phenol beyond 50 mM. The cell biomass yield of phenol
degradation was found to vary between 0.71 g g -1 to 4.48 g g-1 during the course of
phenol degradation in fed-batch system. This variation in cell mass yield could probably
be due to the accumulation of intermediate and their inhibition effect on phenol
consumption as suggested by Wang et al., (1996) and Allsop et al., (1993). In this present
study, the intermediate involved probably ccMA as can be clearly seen in Figure 6.3. The
accumulation of ccMA in the fermentation broth can be explained as follows.
During phenol degradation by C. tropicalis RETL-Cr1 in fed-batch system, pH
was not controlled. However, it was monitored throughout the fermentation process. It
was observed that the pH value dropped from 6.5 to 3.9 at the end of the fermentation
period. The phenomenon of pH reduction could probably due to the increase of chloride
ions and production of organic acids as suggested by Bali and Sengül, (2002) and
Okerentugba and Ezeronye, (2003). pH value as low as 3.9 could have a negative effect
on the conversion of ccMA to muconolactone which is catalyzed by muconate
cycloisomerase (ccMA lactonase) (EC 5.5.1.1).
143
As previously shown in Figure 5.8 (section 5.3.1.3) even at pH 4.5 the synthesis
of C1,2D siezed and consequently there was no formation of ccMA. It could be
hypothesized that there was an inactivation of ccMALe resulting in the intermediate
product, ccMA not being converted to muconolactone and thus got accumulated in the
medium. The hypothetical direct effects of low pH on PH, C1,2D and ccMALe in C.
tropicalis RETL-Cr1 is shown in Figure 6.4, whereas the indirect effect of cumulative
accumulation of ccMA is illustrated in Figure 6.5.
Lower pH (3.9) partially
inhibits PH activity
Lower pH (3.9) stop
C1,2D activity
G
L
F
+
+
D
2
2
F
L
Q
R
2 & 2&
F
X
&
& + P
V
L
F
V
L
F
+
+
2
NADP+ + H2O
O
R
K
F
H
W
D
F
O2+NADPH+H+
C1,2D
O2
Phenol
2H+
& &
+
+
2
+
2
PH
ccMA accumulate in the
medium
Lower pH (3.9) stop
ccMALe activity
ccMALe
No formation of
muconolactone
o
COOC=O
Muconolactone
Figure 6.4 Hypothetical illustration on how low pH (3.9) may affect PH, C1,2D and
ccMA lactonizing enzyme (ccMALe) activity at the end of phenol degradation process
by C. tropicalis RETL-Cr1 in fed-batch fermentation. [Metabolic sequences for phenol
catabolism: PH = phenol hydroxylase, C1,2D = catechol 1,2-dioxygenase (Gaal and
Neujahr, 1979), and Reaction: phenol + O2 + NADPH +H+
(Mörtberg and Neujahr, 1987), catechol + O2
NADP+ + H2O + catechol
ccMA + 2H+ (Ngai et al., 1990)],
NADP = Nicotinamide adenine dinucleotide phosphate.
144
ccMA as noncompetitive inhibitor
slow donor oxidation
D
Decouplers reduce energy gain
from electron transfer
ccMA slow acceptor
reduction by
A uncompetitive inhibition
IC
2e
-
2e
DOX
ICH2
2e
NUTRIENTS
-
-
ADP
+
Pi
A
ATP
red
Biomass
Synthesis
Biomass
Maintennance
Figure 6.5 Hypothetical illustration on how ccMA may affect the primary flow of
electrons and energy during phenol degradation by C. tropicalis RETL-Cr1 (adapted
from Rittmann and Sáez, 1993) – modified; D = primary electron-donor substrate, DOX =
oxidized electron-donor substrate, A = primary electron-acceptor substrate, Ared =
reduced electron-acceptor substrate, ICH2 = reduced internal cosubstrate, ATP =
adenosine triphosphate, ADP = adenosine diphosphate, and Pi = inorganic phosphate;
ccMA = cis,cis muconic acid.
The overall comparison of the performance of phenol degradation in batch and
fed-batch system by free cells of C. tropicalis RETL-Cr1 is presented in Table 6.2.
Phenol degradation rate of 2.3 g L-1 h-1 in fed-batch system was 85 times and 61 times
higher as compared to batch (shake-flask) and bioreactor, respectively. The high phenol
degradation rate is also higher by 15-fold as compared to C. tropicalis Ct2 using fedbatch system reported by Komarkova et al. 2003.
Improvement
to batch (x times)
(bioreactor)
(shake-flask)
ccMA conc.
(g L-1)
YpccMA/s (g g-1)
YpccMA/x (g g-1)
ccMA productivity
(g L-1 h-1)
Catechol conc.(g L-1)
Ypc/s (g g-1)
Ypc/x (g g-1)
Catechol productivity
(g L-1 h-1)
Yx/s (g g-1)
Xmax (g L-1)
Efficiency (%)
Degradation rate(g L-1 h -1)
Conc.(g L-1)
-
-
0.00983
0.036
0.0013
0.00055
0.0005
0.003
0.0107
0.039
0.0013
0.0006
0.0043
0.020
0.0006
0.0204
0.088
0.003
28
0.0395
0.0257
29
9.32
100
9.765
100
0.76
1.0
11.8
17
0.0
0.0
0.0
0.0
3.1x10-5
5.3x10-5
0.0
0.0
0.0
0.0
0.00084
8.9x10-4
2.4x10-4
0.0008
2.1x10-3
1.4x10-3
3.70
0.47
0.05
1.54
12.8
100
0.941
9.9
100
0.376
Fermentation system
Batch
Shake-flask
Bioreactor
282.3
282.3
Parameters/Performance
RETL-Cr1
23.5
33
0.0
0.0
0.0
0.0
2.1x10-5
0.00078
4.1x10-4
1.9x10-4
3.90
0.93
13.5
98.4
39.5
56
0.00092
2.0x10-4
4.4x10-5
1.5x10-5
1.2x10-5
0.00073
1.6x10-4
3.5x10-5
4.48
1.56
30.3
99.7
44.6
63
0.00282
1.0x10-4
5.6x10-5
2.6x10-5
0.0
0.0
0.0
0.0
1.77
1.76
59.3
99.9
Fed-batch
Intermittent Feed
1.882
4.701
28.2
60.5
85
0.00927
1.3x10-4
1.8x10-4
1.4x10-5
0.0
0.0
0.0
0.0
0.71
2.39
61.9
96.2
77.17
Table 6.2: Kinetic parameters/Performance of phenol biodegradation in batch and fed-batch fermentation by C. tropicalis
145
146
Phenol degradation by C. tropicalis RETL-Cr1 in fed batch was also
characterized by phenol being utilized mainly for biomass production. The maximum cell
concentration (Xmax) in fed-batch culture increased up to 61.9 g L-1. This value was
higher by 7-fold than the maximum cell concentration in batch system (bioreactor). This
is because the intermittent feeding technique was able to maintain a constant substrate
concentration and specific growth rate and at the same time maintaining low levels of
phenol in the bioreactor. This condition was able to prevent deleterious effects of
substrate inhibition.
The relationship between growth, substrate consumption and product formation is
expressed in yield coefficients (Yx/s). It can be clearly seen that the fed-batch was
characterized by high phenol degradation rate and biomass yield (Y x/s) but with low
product yields i.e. (catechol) yields (Ypc/s and Ypc/x) and ccMA (YccMA/s and YccMA/x).
High biomass yield could be due to the high metabolic efficiency of C. tropicalis RETLCr1 and also to high carbon/oxygen ration in phenol as suggested by Hao et al., (2002).
The reasons of the low product yield (Yp/s) achieved has been explained previously in this
chapter.
6.4
Conclusions
C. tropicalis RETL-Cr1 was capable of achieving a very high phenol degradation
rate of 2.39 g L-1 h-1, which is 85 times improvement of degradation rate compared to the
shake-flask batch system and 61 times fold as compared to the bioreactor batch system.
The observed biomass yields on phenol (Yx/s) both in the fed-batch and batch systems are
considerably high. However, the biomass yield in the fed-batch system was lower by 6fold than the batch system. Similarly, the product yield (catechol) (Ypc/s, Ypc/x) and the
maximum productivity of cis,cis-muconic acid were also lower due to reasons that have
been discussed. These high biomass yields obtained could probably be due to high
metabolic activity of C. tropicalis RETL-Cr1 and high carbon/oxygen ratio in
147
phenol as suggested by previous researchers.
The results achieved certainly proved the ability of C. tropicalis RETL-Cr1 to
degrade phenol at high concentration using fed-batch system. Its biodegradation
capability can be regarded extremely significant. Thus, it has great potential in the
application for degradation of phenol in industrial waste streams e.g. in industrial effluent
treatment and decontamination of polluted sites.
148
CHAPTER 7
PHENOL METABOLIC PATHWAY OF Candida tropicalis RETL-Cr1
7.1
Introduction
A number of studies on phenol degradation with prokaryotic microorganisms
have been carried out (Hinteregger et al. 1992; Collins and Daugulis, 1997a; Leonard and
Lindley, 1999). Only some members of yeast genera Rhodotorula, Trichosporon, and
Candida that can metabolized phenolic compounds as a sole carbon and energy source (
Katayama-Hirayama et al.,1994; Alexieva et al., 2002; Chen et al., 2002; Santos and
Linardi, 2004).
Aerobic organisms degrade phenol to catechol followed by oxidative cleavage of
the ring. This oxidative ring cleavage of catechol can occur in one of two ways. The
ortho cleavage to produce cis,cis-muconic acid (ccMA) or the meta cleavage to produce
2-hydroxymuconic semialdehyde (2-HMSA). The production and accumulation of these
intermediates during phenol degradation has been commonly observed (Li and
Humphrey, 1989; Mörsen and Rehm, 1990; Allsop et al., 1993). The identification of
products formed during the biodegradation process of phenol is essential for a better
understanding of the degradation mechanism.
The final objective of the present study was to determine the intermediates
produced during phenol degradation by C. tropicalis RETL-Cr1 and to postulate possible
149
phenol metabolism pathway.
7.2 Materials and Methods
7.2.1
Meta-cleavage dioxygenase assays
To determine whether meta-cleavage of phenol was involved, spray plate method
and a test tube assay as described by Kim and Zylstra (1995) were performed. A spray
plate method was used to screen for colonies showing meta-cleavage dioxygenase
activity on plates. An ether solution of catechol (0.1% w/v) was sprayed on to colonies of
C. tropicalis RETL-Cr1 and observed for yellow colour formation as a result of a metacleavage of catechol by meta-cleavage dioxygenase.
A test tube assay was also employed for detection of low levels of meta-cleavage
dioxygenase activity. One loopful of cells grown overnight on plates was suspended in 1
mL of 50 mM phosphate buffer (pH 7.5 and pH 6.5). 20µL Catechol (20 mM stock
solution in methanol) was added, and the formation of a yellow colour was monitored
visually over time.
7.2.2 Determination of cis,cis-muconic acid
Cis,cis-muconic acid as indicator of ortho-cleavage of phenol was determined
using HPLC. The HPLC-analytical parameters and procedures used in determination of
cis,cis-muconic acid is presented in section 3.5.5 of Chapter 3.
7.2.3 Experimental Design
The flowchart of the experimental design carried in this study is presented in
Figure 7.1.
150
Methods
Test 3
HPLC –Chromatography
to detect cis,cis-muconic
acid (ccMA)
ortho-pathway
Enzymatic assays
Test 1
Spray method
Test 2
Test-tube method
To detect meta cleavage
dioxygenase activity
meta-pathway
Figure 7.1 Experimental design to postulate phenol metabolic pathway of C. tropicalis
RETL-Cr1.
7.3
Results and Discussion
7.3.1
Determination of intermediates of C. tropicalis RETL-Cr1
The first step taken to determine the possible phenol metabolic pathway of C.
tropicalis RETL-Cr1 was to perform the enzymatic assays to detect catechol 2,3
dioxygenase activity for meta-pathway. The enzymatic assays performed did not indicate
presence of the activity for this enzyme. In meta-cleavage pathway, catechol is converted
to 2-hydroxymuconic semialdehyde (2-HMSA) catalyzed by catechol 2,3-dioxygenase
151
(C2,3D). Colonies of C. tropicalis RETL-Cr1 cultured on plates or suspended in
phosphate buffer did not develop a yellow colour when catechol was sprayed or added
suggests that there was no significant formation of 2-hydroxymuconic semialdehyde
indicative of the meta-pathway (Bushwell, 1975; Kim and Zylstra (1995). It was then
assumed that C. tropicalis RETL-Cr1 probably metabolized phenol via the orthopathway. Then the next step was to perform the HPLC chromatography to detect the
presence of cis,cis-muconic acid, the intermediate indicator for ortho-pathway.
During degradation of phenol using free cells of C. tropicalis RETL-Cr1, HPLC
analysis of the samples taken from batch system containing varying initial phenol
concentration (IPC) from 282- to 940 mg L-1 (3 mM-10 mM) and fed-batch containing
phenol from 0.376 to 77 g L-1 (4 mM – 820 mM) feed solution revealed the presence of
catechol and cis,cis-muconic acid as intermediates. Figure 7.2 illustrates a typical HPLC
chromatogram recorded for samples taken after 7 hours for phenol degradation by C.
tropicalis RETL-Cr1 in medium (pH 6.5) under aerobic condition at 30oC, at an initial
phenol concentration of 3 mM.
2
catechol
cis,cismuconic
acid
Intensity
3
phenol
unidentified
products
01
1
02
Retention time
Figure 7.2 Typical HPLC chromatogram recorded for an aerated suspension: (01 & 02)
unidentified products; (1) cis,cis-muconic acid ; (2) catechol ; and (3) phenol during
phenol degradation in batch culture of C. tropicalis RETL-Cr1 at initial phenol
concentration of 3 mM after 7h incubation.
152
The formation of other products (unidentified) was also observed. These products;
peaks 01 and 02 could be assumed to be other possible intermediates produced in the
ortho-cleavage pathway such as muconalactone, 3-oxioadipic acid (ß-ketoadipate), or
0.8
21
0.7
18
0.6
15
0.5
12
0.4
9
0.3
6
0.2
3
0.1
0
ccMA and Catechol conc.
-1
(mg L )
-1
Phenol conc. (g L ), OD600
succinic acid as suggested by Bugg and Winfield, (1998).
0
0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18
Time (h)
Figure 7.3 Time course of phenol (Ŷ) degradation in batch system (shake-flask) using C.
tropicalis RETL-Cr1 at IPC of 3 mM, pH 6.5, and detection of intermediates; catechol
(Ɣ) and cis,cis-muconic acid (ccMA) (Ÿ). Biomass concentration (Ƒ) of C. tropicalis
RETL-Cr1.
As clearly seen in Figure 7.3, catechol and cis,cis-muconic acid (ccMA) were not
formed simultaneously. Catechol was formed at the earlier stage of the reaction while
cis,cis-muconic acid was formed at the later stage of the biodegradation process. The
maximum concentration of catechol was 20.4 mg L-1 after 7 h of incubation. This
maximum concentration of initial phenol recorded was similar to that reported by Chung
et al., (2003) where the concentration of catechol produced by P. putida CCRC14365
was always less than 21 mg L-1 (0.223 mM) at 30oC and pH 6.8. In contrast, cis,cismuconic acid was only formed at later stage of the incubation period. The onset of the
formation of ccMA was after 6 hours of the degradation process and increased
exponentially as phenol concentration decreased. The catechol concentration finally
153
leveled off after 11 h of incubation period thus appeared to accumulate in the medium.
7.3.2
Phenol metabolic pathway of C. tropicalis RETL-Cr1
Many studies have focused on biodegradation of phenol and phenolic compounds
with respect to degradation pathways (Dagley, 1985; Katayama-Hirayama et al., 1991).
The general principle of pathways for aerobic aromatic catabolism is best described by
Dagley, 1986 and Harayama and Timmis, 1992). This aerobic aromatic catabolic
pathway generally consists of three stages. The three stages are: (1) The conversion of the
growth substrate to catechol, (2) ring-cleavage and (3) metabolism of the ring-cleavage
product to central metabolites by either the ortho or meta-pathways (Figure 7.4).
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Figure 7.4 General principle of aerobic aromatic catabolism in bacteria. (Dagley, 1986;
Harayama & Timmis, 1992) (Adapted from Williams and Sayers, 1994).
154
The first reaction in phenol degradation is catalyzed by phenol hydroxylase (PH)
(EC 1.14.13.7) whereby one oxygen atom of molecular oxygen into the aromatic ring to
form catechol as the central intermediate. This catechol is then converted to cis, cismuconic acid through ortho-cleavage pathway. This pathway is also known as ßketoadipate pathway which is catalayzed by catechol 1,2-dioxygenase (C1,2D) (EC
1.13.11.1) or converted to 2-hydroxymuconic semialdehyde (2-HMSA) through the metapathway which is catalyzed by catechol 2,3-dioxygenase (C2,3D) (EC 1.13.11.2). After
several subsequent reactions, ortho pathway will lead to succinyl-CoA and acetyl-CoA.
On the other hand, meta pathway will lead to pyruvate and acetyldehyde and finally both
pathways will incorporated into the tricarboxylic acid cycle (TCA) or Krebs cycle
(Shingler, 1996). Therefore, from this phenol metabolic pathway, cis,cis-muconic acid
(ccMA) is considered the indicator for ortho-cleavage pathway and 2-hydroxymuconic
semialdehyde (2-HMSA) is the indicator for meta-cleavage.
The HPLC chromatography and enzymatic assays revealed that C. tropicalis
RETL-Cr1 appeared to metabolize phenol via ortho cleavage pathway (Figure 7.5). 2hydroxymuconic semialdehyde (2-HMSA) has been reported to be responsible for colour
change in culture medium (greenish to yellow) during phenol degradation (Li and
Humphrey, 1989; Mörsen and Rehm, 1990). However, in this present study no colour
changes of media was observed, which further suggests that Candida tropicalis RETLCr1 probably did not metabolize phenol through meta-pathway. Hence it could be
postulated that ortho-pathway for phenol degradation taken by C. tropicalis RETL-Cr1
could be similar to one reported for Candida tropicalis by Hashimoto, (1970, 1973),
Neujahr et al., (1974); Middelhoven, (1993), Krug and Straube, (1986) and Bastos et al.,
(2000a).
155
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%
Figure 7.5 Postulated ortho-pathway for degradation of phenol by C. tropicalis RETLCr1. [Metabolic sequences for phenol catabolism: PH = phenol hydroxylase, C1,2D =
catechol 1,2-dioxygenase, ccMALe = cis,cis muconic acid lactonizing enzyme (Gaal and
Neujahr, 1979; Bugg and Winfield, 1998), and Reaction: phenol + O2 + NADPH +H+
NADP+ + H2O + catechol (Mörtberg and Neujahr, 1987), catechol + O2
ccMA + 2H+ (Ngai et al., 1990)], NADP = Nicotinamide adenine dinucleotide
phosphate.
156
7.4
Conclusion
Phenol catabolism was confirmed through the detection of the intermediary
products namely catechol which was formed in the early stage and cis,cis-muconic acid
in the later stage of incubation. HPLC chromatography and enzymatic results showed
that this indigenous phenol-degrading yeast, C. tropicalis RETL-Cr1 seemed to
metabolize phenol via ortho-cleavage pathway.
157
CHAPTER 8
CONCLUSIONS AND FUTURE RESEARCH
8.1
Conclusions
The main objectives of this study have been met. The findings are summarized as
follows:
Of the 35 strains isolated aerobically, 8 were selected to be studied for growth in
RM broth containing phenol. Isolates RAS-Cr1, RETL-Cr1 and RETL-Cr3 were further
screened for tolerance of phenol and survivality test. The most prominent degraders of
phenol RETL-Cr1 was able to grow profusely on RM agar at pH 6.5 at 37oC in the
presence or absence of 3 mM (282 mg L-1) with inadditional carbon source with
percentage survival of 97%. This result is a good indicator that RETL-Cr1 was able to
degrade phenol as a sole carbon and energy source.
Isolate RETL-Cr1 consists of oval-shape, budding cells which were found to be
the best phenol-utilizer with 100% removal efficiency at µ = 0.313 h-1 followed by
RETL-Cr3 (16%) at µ = 0.359 h-1 and RAS-Cr1 (14%) at µ = 0.322 h-1. The most
distinctive features of the isolate RETL-Cr1 is that it has an extremely high tolerance
level towards phenol up to 60mM. Based on a BLASTN search of GenBank, the
complete sequences of ITS1-5.8S rDNA-ITS2 regions and portions of I8S and 28S for
158
the purified DNA products of RETL-Cr1 shared 98% similarity with Candida tropicalis.
Isolate RETL-Cr1 was redesignated Candida tropicalis RETL-Cr1. The strain was
deposited into GenBank database under the accession number AY725426.
The capability of C. tropicalis RETL-Cr1 to degrade phenol at high levels
prompts the next stage of study.
In the second phase of the study, batch shake-flask experiments to investigate the
influence of process variables like pH, temperature, initial phenol concentrations in the
presence or absence of glucose, on phenol biodegradation were carried out and optimum
process conditions has been established. C. tropicalis RETL-Cr1 was found to degrade
phenol completely in media either in the presence and absence of glucose. For substrate
mixture of phenol and glucose, the degradation of the compounds was simultaneous. This
yeast strain was able to degrade phenol at temperatures ranging between 30oC to 37oC
and at wide range of pH values from 5.5 up to 7.0. The optimum physiological conditions
for aerobic degradation of phenol by C. tropicalis RETL-Cr1 was 30oC, pH 6.5 without
glucose.
The fact that this yeast was able to utilize phenol as sole carbon source or in
mixture over a wide range of temperature and pH, is an advantage because these
conditions may resemble the unfavourable conditions encountered in some environments
contaminated by industrial aromatic compounds such as that generated by the oil
refinery. Furthermore, C. tropicalis RETL-Cr1 possess simultaneous substrate utilization
pattern for substrate mixtures of phenol and glucose. Thus, these characteristics made this
indigenous yeast strain suitable microorganism for treating industrial wastes such as
industrial processes effluents especially that which contain phenol.
The catabolism of phenol was confirmed with the detection of intermediary
products, catechol and cis,cis-muconic acid. High catmax, catechol productivity, ccMAmax
and ccMA productivity were achieved at temperatures ranging from 30oC to 37oC and pH
from 5.5 to 7.0 suggesting that these physiological conditions were optimum for phenol
159
hydroxylase (PH) and catechol 1,2-dioxygenase (C1,2D) activities. Temperature of above
37oC may have caused denaturation of C1,2D of C. tropicalis RETL-cr1 and hence
catechol was not converted to ccMA. The catechol may have accumulated in the medium
and exerted inhibition on phenol hydroxylase which is responsible for phenol conversion
to catechol.
In the third phase of the study, a detailed study of phenol degradation in batch
system both in shake-flask and bioreactor were carried out, and phenol degradation
potentials of C. tropicalis RETL-Cr1 was determined and compared. Phenol was depleted
faster in bioreactor than in shake-flask cultures by 1.5-fold owing to the improved growth
conditions due to mixing and aeration. The highest removal efficiency in shake-flask
cultures was 100% with a degradation rate of 0.028 g L-1 h-1 at 3 mM after 14 h
cultivation. Complete degradation of phenol was improved to 39.5 mg L-1 h-1 after 10
hours cultivation with a mean generation time of µ = 0.5381 h-1.
The kinetics of phenol degradation by C. tropicalis RETL-Cr1 in batch cultures
was investigated over a wide range of IPC (3 mM to 10 mM (282-941 mg L-1)). Under
batch condition, the phenol degradation rate and specific growth rates (µ) decreased as
the initial phenol concentrations increased suggesting substrate inhibition after 5 mM
(470 mg L-1) initial phenol concentration. Phenol shows substrate inhibition as reported
elsewhere in other literatures. Respective kinetic parameters were reported as µ in the
range of 0.1557 to 0.4116 h -1 and Yx/s = 24 – 39 g g -1.
Under the conditions tested, the C. tropicalis RETL-Cr1 was capable of achieving
a very high phenol degradation rate of 2.39 g L-1 h-1, which was 85 times improvement of
degradation rate compared to the shake-flask batch system and 61 times fold as compared
to bioreactor batch system. As shown from batch system, high biomass yield on phenol
(Yx/s) was observed and varied from 0.7 to 4.5 g g-1 for fed-batch system. This high
biomass yield obtained could probably be due to high metabolic activity of C. tropicalis
RETL-Cr1 and high carbon/oxygen ratio in phenol.
Phenol biodegradation obtained by fed-batch was characterized by a high
160
degradation efficiency (96-100%), rate (2.3 g L-1 h-1) and substrate yield (Yx/s = 0.714.48 g g -1) but with lower product yields (Ypc/s = 1.6x10-4 – 2.1x10-3 g g -1; Ypc/x =
3.5x10-5-1.4x10-3 g g -1), catechol productivity (1.2x10-5 -5.3x10-5) and ccMA (1.4x10-52.6x10-5 g L-1 h-1). The low product yield was probably due to the utilization of catechol
simultaneously with phenol and thus detected at low concentration. On the other hand,
ccMA product from catechol was observed to be low as it is difficult to utilize catechol
due to its toxicity. These results also indicate further degradation of these metabolites.
The variation in biomass yield could probably be linked to inhibition caused by
accumulation of ccMA in the medium. It has been shown that a drop in pH to 3.9 could
have inhibited the synthesis of the ccMA lactonizing enzyme (muconate cycloisomerase).
Hence, ccMA was not converted to muconolactone thus get accumulated in the medium.
This accumulation of ccMA may affect the primary flows of electrons and energy during
phenol degradation by C. tropicalis RETL-Cr1.
The results achieved certainly proved the ability of C. tropicalis RETL-Cr1 to
degrade phenol at high concentration using fed-batch system. Its biodegradation capacity
can be regarded as extremely significant. Thus, it has significant potential in the
application of phenol degradation in industrial waste streams e.g. in industrial effluent
treatment and decontamination of polluted sites.
The chromatographic and enzymatic results indicated that this indigenous phenoldegrading yeast, C. tropicalis RETL-Cr1 degraded phenol via ortho-cleavage pathway.
The search for pollutant-degrading microorganisms from contaminated sites has
been proven successful. This support the idea that this contaminated environments is a
valid choice for the isolation of microorganisms useful in bioremediation as indicated in
the literature search. Therefore the biodegradation capability of this C. tropicalis RETLCr1 is truely significant. This is the first report in Malaysia of its kind of an indigenous
phenol-degrading yeast.
161
The overall representation of this study is presented in Figure 8.1 of Appendix
B1.
8.2
Future Research
This study has opened up several avenues of research which is as follows:(i)
For further enhancement of phenol biodegradation, to perform fed-batch by
variable feeding or by cell- immobilization of C. tropicalis RETL-Cr1 and its
application of different types of bioreactors.
(ii)
Production, purification and characterization of biosurfactant from C.
tropicalis RETL-Cr1.
(iii)
Development of cell-based biosensor using C. tropicalis RETL-Cr1 that able
to detect phenol.
(iv)
Development of bench-scale biofilter using C. tropicalis RETL-Cr1 for
cleaning of phenol from industrial wastewater.
(v)
Isolation, identification and characterization genes of C. tropicalis RETL-Cr1
responsible for the degradation of phenol.
(vi)
Isolation, identification and characterization of phenol hydroxylase and
catechol 1,2 dioxygenase and muconate cycloisomerase from C. tropicalis
RETL-Cr1 responsible for phenol degradation.
(vii)
Further study on biodegradative capability of C. tropicalis RETL-Cr1 towards
other recalcitrant aromatic hydrocarbons such as arthracene and crysene.
162
(viii) Optimization study on production of cis,cis-muconic acid from phenol or
other sources such toluene, benzene and benzoic acid using C. tropicalis
RETL-Cr1.
(ix)
Bioaugmentation for treatment of petroleum hydrocarbon in sludge farm of an
oil refinery.
163
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APPENDIX A1
0.8
y = 0.0723x
R2 = 0.9784
0.7
0.6
OD600
0.5
0.4
0.3
0.2
0.1
0
0.18
0.32
0.36
0.48
0.68
0.8
1
1.2
1.23
g dw L-1
Figure A1 Plot of OD600 Vs dry weight during batch cultivationCalibration curve for calculation of dry cell weight of C. tropicalis
RETL-Cr1.
215
APPENDIX A2
0.4
R2 = 0.9979
Absorbance at 450nm
0.35
0.3
0.25
0.2
0.15
0.1
0.05
0
0
0.25
0.5
0.75
1
Glucose conc. (mM)
Figure A2 Standard Curve use to calculate glucose using Shimadzu Spectrophotometer
Model based on Sigma® procedure 510 (Sigma® Diagnostics, St Louis, MO).
216
APPENDIX A3
DETERMINATION OF GLUCOSE USING SIGMA® PROCEDURE 510 (SIGMA®
DIAGNOSTICS, ST. LOUIS, MO)
A. Principle
The testing is based on the oxidation of glucose with glucose oxidase. When the
reaction occurs a brown-colored complex is formed. The intensity of the color is
directly proportional to the glucose concentration in the sample.
Principle of the reaction:
Glucose oxidase
Glucose + 2 H2O + O2 ———————————> Gluconic acid + 2 H2O2
Peroxidase
H2O2 + o-Dianisidine ————————————> Oxidized o-Dianisidine
[Colorless]
[Brown]
B. Preparation of Glucose Standards
Concentration
Volume of glucose Volume of distilled water
(mM)
(mL)=(µL)
(mL)
0
0.25
0.50
0.75
1.0
0
0.005 = (5)
0.010 = (10)
0.015 = (15)
0.020 = (20)
20
19.995
19.990
19.985
19.980
Total volume
(mL)
20
20
20
20
20
C. Preparation of Combined Enzyme Color Reagent (CECR)
1.“The ENZYME SOLUTION is prepared by adding the contents of 1 capsule of
PGO Enzymes (glucose oxidase and peroxidase) to 100 mL of distilled water
in an amber bottle. Invert bottle with gentle shaking to dissolve.
2. COLOR REAGENT SOLUTION is prepared by reconstituting one vial of
o-Dianisidine Dihydrochloride with 20 mL of distilled water.
3. COMBINED ENZYME-COLOR REAGENT (CECR) SOLUTION is
prepared by combining 100 mL of Enzyme Solution and 1.6 mL of Color
Reagent Solution. Mix by inverting several times or with mild shaking.
217
APPENDIX A3 - continue
D. Procedures for glucose determination
1. Label each test tube (Eppendorf @ 2 mL) accordingly.
2. Pipette 100 µL (0.1 mL) standard or sample solution to the test tube
3. Add 1.0 mL of the CECR solution to the standard or sample solution
4. Mix thoroughly each of the test tubes and incubate all tubes at 37oC for 30
minutes. Protect from light (dark).
5. Read absorbance for each test tube at 450 nm against the blank within 30
minutes.
6. Save and print results
218
APPENDIX A4
HPLC-ANALYTICAL PARAMETERS FOR DETERMINATION OF PHENOL,
CATECHOL AND CIS,CIS MUCONIC ACID
The HPLC analytical parameters used were:
Detector
: Waters UV detector 2487
Detection
: UV-Absorption Ȝ = 280 nm.
HPLC-pump
: Waters HPLC-pump model 600
Flow rate
: 1.0 mL min-1
Reversed-phased column
: Waters Hypersil C18 5µm (4.6.mm x 250 mm)
Mobile phase (solvent)
: A (1% acetic acid in H2O)
B (1% acetic acid in Acetonitrile)
Composition of mobile phase : A =70%, B= 30%
Temperature
: Room temperature
Elution
: Isocratic
Run time
: 10 minutes
Data analysis
: Millennium 32® Chromatography Manager 3.2.
219
APPENDIX A5
HETEROTROPHIC PLATE COUNT
TEST METHOD APHA 9215
Media used
-
: Ramsay Medium Agar (as described by Ramsay et al., 1983).
Mix thoroughly all components except MgSO4.7H2O and glucose in 1000 mL
distilled water and then autoclaved for 15 minutes at 121oC at 15 pounds per sq.
in (psi) steam pressure. Filter-sterilized MgSO4.7H2O and glucose added after
autoclaving.
Media and glassware for sterilization
-
Media
-
Measuring cylinder and beaker
-
Pipette tips
-
Distilled water
-
Dilution bottles
Procedure
-
Dispense agar onto Petri dishes and incubate at 30oC for 24h.
-
1 mL sample was diluted in 9 mL distilled water
-
Prepare dilution from 100, 101, 102, 103, 104, 105, 106, 107, 108, 109 (1000µL for
each sample of dilution)
-
Take a dilution from 103, 104, 105 and make a duplicate
-
Take 100µL inoculum onto Petri dishes and spread using a bend glass rod
-
Incubate inoculated Petri dishes at 37oC for 1-7 day (after 7 days no checking
necessary.
220
APPENDIX A6
CELLULAR MORPHOLOGY AND BIOCHEMICAL TESTS BASIC
PROCEDURES
1.
Cellular morphology
1.1
Gram staining
A fixed smear was prepared from a 24 hour microbial culture then covered with 1
or 2 drops of crystal violet solution. The stain was allowed to remain on the smear for 1
minute. The stained was poured off and the preparation washed cautiously with distilled
water. The remaining water on the slide was blotted and 1 or 2 drops of Lugol’s solution
were added to the preparation and allowed to stand for 1 minute. The preparation was
then washed with 95% ethyl alcohol until the fluid running off the slide is colourless.
Safranin was then added to the preparation and was left for 30 seconds. The preparation
was washed with water and blot dry. Gram-positive cells appear violet and Gramnegative cells appear red when observed under the light microscope.
1.2
Endospore staining
A slide smear from a 24 hour culture was prepared and fixed thoroughly by
placing the slide above boiling water. The smear was covered with a strip of filter paper.
Malachite green was poured on to the preparation so that the whole slide was covered.
The slide was heated for 5 minutes. The smear was rinsed thoroughly with distilled water.
Safranin was then added to the preparation and was left for 20 seconds. The preparation
was washed with water and blot dry.
221
1.3
Motility
Motility was determined using the culture method. The inoculating needle was
used to transfer the bacteria to the semi-solid media. The semi-solid media was stabbed
using a straight “in and out” movement. The culture was then incubated for 24 hours. If
growth occurs only on the stab, it is negative and if growth occurs away from the stab, it
is positive.
2.
Biochemical Tests
Biochemical characterization of the isolates include the test for fermentation
reaction (lactose) and enzyme activity (catalase, citrate, methyl red, oxidase, urease and
Vogues Proskauer) were carried out as described by MacFaddin, (1980).
2.1
Catalase test
Catalase is found in most aerobic and facultative anaerobic bacteria (Gunsalus
and Stainer, 1961); the main exception is Streptococcus spp. Catalase test was determined
whether the organism has the enzyme catalase which neutralizes hydrogen peroxide
(H2O2) into oxygen and water. A RM plate was inoculated and incubated at 37oC for 24
hours. After incubation, three drops of 10% v/v hydrogen peroxide were added directly to
a colony and if the peroxide effervesces (appearance of gas bubbles) the organism is
catalase positive.
2.2
Oxidase test
Some bacteria contain the enzyme cytochrome oxidase which activates the
222
oxidation of reduced cytochrome by molecular oxygen (Steel, 1961; Wilson and Miles,
1964a) which in turn acted as an electron acceptor in the terminal stage of the electron
transfer system (Mahler and Cordes, 1966; Wilson and Miles, 1964a). A few drops of
1% v/v tetramethyl-ȡ-phenylenediamine-dihydrochloride (TPD) were place onto a piece
of filter paper. The filter paper was smeared with bacterial culture taken in a wire loop. A
purple colour usually developed within 5-10 seconds is recorded as a positive result. A
positive 10-60 seconds is considered a delayed result (Steel, 1961). Steel (1961) stated
that a later development of colour past 60 seconds period denotes a negative oxidase test.
2.3
Methyl Red test
The methyl red test (MR) is based on the used of a pH indicator, methyl red, to
determine the hydrogen ion concentration (pH) present when an organism ferments
glucose (Wilson and Miles, 1964b). An MR broth was inoculated and incubated at 37oC
for 48 hours. After incubation, 5 drops of methyl red reagent was added to the culture
and the tube was shaken gently. A red colour indicated acid formation (positive) and a
yellow colour indicated a negative result.
2.4
Vogues-Proskauer test
The Vogues-Proskauer (VP) test is to determine the ability of some organisms to
produce a neutral end product, acetylmethylcarbinol (acetoin) from glucose fermentation.
The production of of acetoin is one pathway for glucose degradation occurring in bacteria
(Abd-el-Malek and Gibson, 1948). VP broth was inoculated and incubated at 37oC for 48
hours. After incubation, 1 ml of culture was transferred into a sterile tube, and 0.6 ml ĮNapthol and an exact amount of 0.2 ml potassium hydroxide solution was added, shaken
gently. A red colour developing within 15-30 minutes indicated a positive reaction.
223
2.5
Lactose fermentation test
Lactose test is to determine whether a bacteria able to ferment lactose producing
lactic acid, carbon dioxide and water. Tubes containing lactose and an inverted Durham
vial and phenol red indicator was inoculated and incubated at 37oC for 24 hours. Acid
production is indicated if the phenol red turns yellow and gas production is noted by a
bubble in the inverted vial.
2.6
Urease test
Urease test is to determine the ability of an organism to split urea, forming two
molecules of ammonia by the action of the enzyme urease with resulting alkalinity.
Urease is considered constitutive enzyme since it is synthesized by certain bacteria
regardless of the presence and absence of its substrate urea (White and Hill, 1941;
Burrows and Moulder, 1968). A tube of urea medium was inoculated and incubated at
37oC for 24 hours. The appearance of dark pink colour indicated a positive test, and a
negative reaction will be either yellow or orange colour.
2.7
Citrate test
Citrate test is to determine the ability of an organism of utilizing citrate as the sole
carbon source for metabolism with resulting alkalinity. Energy can be supplied to some
bacteria in the absence of fermentation or lactic acid production (Gunsalus and Stainer,
1961) by the use of citrate as the sole source of carbon. A tube of a Koser citrate agar was
inoculated and incubated at 37oC for 24 hours. A colour change in the medium from
green to blue is an indication of a positive result.
224
2.8
Triple sugar iron (TSI)
TSI agar provides information concerning glucose fermentation, utilization of
sugars lactose and sucrose, and the anaerobic respiratory process that uses sulphur as the
final electron acceptor to produce hydrogen sulphide. The medium was prepared as a
slope and a deep butt. The medium was inoculated heavily with a straight wire by stab
inoculation deep down into agar of the butt. When removing the wire, the slant was
streaked. All tubes were incubated for 18 h at 37oC. After incubation, the tubes were
examined for colour changes indicating the reactions which may have occurred and
recorded according to system described by MacFaddin (1980).
2.9
Growth on special media
2.9.1
MacConkey agar (MCA)
This test is used to differentiate microorganisms which are lactose and non-
lactose fermenters. MCA was inoculated and incubated at 37oC for 24 to 48 hours. A
lactose-fermenter produced red colonies, and non-lactose fermenter produced cream
colonies.
2.9.2
Blood agar
This test is used an indicator of various haemolytic properties of some bacteria. It
is one of the techniques to screen biosurfactant-producing bacteria (Passeri et al., 1991).
Blood agar was inoculated and incubated at 37oC for 24 hours. A narrow zone of
discolouration of surrounding colony (alpha (Į) haemolysis ) disclose a partial
heamolyzed red cells, and a relatively wide (2 to 4 mm) clear colourless zones (beta (ȕ)
haemolysis) around the colony. No discolouration or haemolyis of the blood agar
medium is referred as gamma (Ȗ) haemolysis.
CO,WO,ETL,SSF,
SMSF, BTL,STL
Location 2:EMOR
SCREENING AND CHARACTERIZATION
OF
3 SELECTED STRAINS
( RAS-Cr1, RETL- Cr1, RETL-Cr3)
Plating out on RM agar at 37oC
(54 strains obtained)
(RM) at 37oC for 24 hrs
(aerobic and anaerobic)
CULTURE
Samples
MOLECULAR IDENTIFICATION OF MICROBE OF
INTEREST (RETL-Cr1)
DEGRADATION STUDY OF 3 SELECTED STRAINS
(RAS-Cr1, RETL-Cr1, RETL-Cr3)
AB, AS, FTA
Location 1: TPSB
ISOLATION AND SCREENING
Batch
(Bioreactor)
-ve results
Test-tubes
method
Spray
method
Enzymatic assays
To detect 2,3
dioxygenace activity
for meta pathway
RETL-Cr1 isolated from Exxon Mobil Oil Refinery treatment plant
Figure 8.1 Schematic representation of the biodegradation of phenol by C. tropicalis
Kinetics and performance of phenol
degradation by C. tropicalis RETL-Cr1
Orthopathway
ccMA
detected
HPLC Chromatography
to detect cis,cis muconic
as indicator of ortho
POSTULATION OF POSSIBLE PATHWAY
OF DEGRADATION
Fed-batch
(Bioreactor)
Determination and quantification of
Catechol and ccMA
IPC
3mM,5,mM,
7mM,10mM
pH
4.5,5.5,6.5
,7.0, 8.0
+&glucose
Temp ( oC)
30, 37, 40
Optimization
Batch (Shakeflask)
BATCH AND FED-BATCH BIODEGRADATION
BIODEGRADATION OF PHENOL BY CANDIDA TROPICALIS RETL-Cr1 ISOLATED FROM EXXON
MOBIL OIL REFINERY TRATMENT PLANT
APPENDIX B1
225
226
APPENDIX C: Publications
Piakong M.T., Adibah, Y., Madihah, M.S., Noor Aini, A.R., Haryati, J.
Roslindawati, H. and S. Hasila, H. (2002). Isolation and characterization of oildegrading bacteria from oil and oil samples. Proceedings: Towards
Commercialization of Microbiology Research-25th Malaysia Microbiology Society
Symposium,Kota Bahru, Kelantan 8-11 Sept 2002.
Piakong, M. T., Haryati, J., S. Hasila, H., Roslindawati, H., Adibah, Y., Madihah,
M.S. and Noor Aini, A. (2004). Biodegradation of phenol by locally isolated
strains from petrochemical wastewater treatment plants. In: Ujang, Z. and
Henze, M. (Eds.) Water & Environ. Management Series. – Environmental
Biotechnology : Advancement in water and wastewater application in the tropics.
IWA Publishing, U.K. pp. 109-114.
Piakong M.T, Noor Aini A.R, Adibah Y., Madihah M. S, Aishah, H. and Sharifah
Norhafizah S. M.R. (2004). Molecular identification of Candida tropicalis RETLCr1 by PCR amplification of ribosomal DNA. Borneo Sci. 15: 15-22.
Piakong M.T, Noor Aini A. R., Adibah Y., Madihah M.S, Aishah H. and Sharifah
Norhafizah S.M.R. (2004). Phenol biodegradation by a yeast Candida tropicalis
RETL-Cr1. Proceedings: Role of Environmental Science and Technology in
Sustainable Development of Resources. KUSTEM 3rd Annual Seminar on
Sustainable Science and Management. 4-5 May 2004 Kuala Terengganu,
Terengganu.
Fernandez, C.C., Noor Aini A.R., Zaharah, I. and Piakong, M.T. (2005). Development
of enzyme assay and preliminary kinetic studies for the enzyme(s) from Candida
tropicalis RETL-Cr1 involved in phenol degradation. Pak. Biol. Sci. 8( ) CCCC.
Zaharah I, Noor Aini A. R, Adibah Y, Madihah M. S., Fahrul Z. H., Piakong M. T.,
Azaliza S. W. and Salwa M. (2005). Bioremediation of target pollutants from
industrial wastes using locally isolated microbes. Research Focus- Bull. R&D RMC,
Universiti Teknologi Malaysia. 16: XX