Isolation and Identification of Tannase producing bacteria from environmental soil sample (1)

Isolation and Identification of Tannase
producing bacteria from environmental soil
sample
Submitted by
Sunehera Sarwat
Student ID - 1030251047
Fall-2014
DEPARTMENT OF BIOLOGY AND CHEMISTRY
NORTH SOUTH UNIVERSITY
DHAKA BANGLADESH
December, 2014

The Project Entitled
Isolation and Identification of Tannase producing
bacteria from environmental soil sample
Is submitted to the Department of Biology and Chemistry, North
South University for Partial Fulfillment of the requirement for the
Degree of BS in Biochemistry and Biotechnology for the session of
“Fall-2014”
---------------------------------
Sunehera Sarwat
ID: 1030251047
DEPARTMENT OF BIOLOGY & CHEMISTRY
NORTH SOUTH UNIVERSITY
DHAKA BANGLADESH
December, 2014

Approval Certificate
North South University
Dhaka, Bangladesh
This is to certify that we the members of the Thesis Committee have carefully read
and recommended to the Department of Biology and Chemistry, North South
University, Dhaka for the approval of this project entitled:
Isolation and Identification of Tannase producing bacteria from
environmental soil sample
Submitted by Sunehera Sarwat, ID-1030251047 for the partial fulfillment of the
requirements for the Degree Bachelors of Science in Biochemistry and
Biotechnology.
----------------------------------
Obaidur Rahman, PhD
Assistant Professor
Dept. of Biology & Chemistry
North South University, Dhaka
Supervisor (Project)
----------------------------------
Sohidul Islam, PhD
Assistant Professor
Dept.of Biology & Chemistry
Supervisor (Project)
----------------------------------
Abdul Khaleque, PhD
Associate Professor
Dept. of Biology & Chemistry
(Member of the Committee)
----------------------------------
Prof. Dr. G.U. Ahsan
Dean, School of Health and Life
Sciences
(Chair of the Committee)

ii
Acknowledgements
Before I begin I would like to express my gratitude to Almighty Allah, the most
Merciful and Beneficent. Without His blessings, compassion and mercifulness I
would not have been able to come this far.
I would like to express the deepest appreciation to my research supervisor and mentor
Dr. Obaidur Rahman, Assistant Professor, Department of Biology and Chemistry,
North South University for guiding me very meticulously in each and every steps of
the project work. It was a great privilege and pleasure for me to carry on my research
work under his supervision who always took out his time for me. He taught me what
research is truly like, and his enthusiasm for research always encouraged me to pursue
me dream more. His guidance, skilled advice and constructive criticism greatly helped
me to finish my research work and taught me the valuable lesson in patience.
I would also like to express my thanks to my second supervisor, Dr. Sohidul Islam,
Assistant Professor, who believed in me. He always pushed me to my limits which
made me stronger each day. I am glad he was there to teach me the hardest lessons in
my life and help me to come out of my comfort zones and overcome them.
My earnest and deepest respect to Dr. Abdul Khaleque, Chairman and Associate
Professor and Dr. Kazi Nadim Hasan, Associate Professor, for their amiable behavior,
guidance, support and kindness from the very start of my undergraduate life. I truly
consider myself to be lucky for being their student.
I owe my deepest and most genuine appreciation to Dr. Donald James Gomez, former
Dean of School of Health and Life Sciences. He has been like a father figure to me
from the very beginning of my undergraduate life, without whose guidance and
support some of the hardest times of my life would have been unbearable.
I am obliged to thank two of my dearest friends, Nawseen Tarannum and Tasnuva
Ahmed with them I have spent the last four years of my undergraduate life. They were
truly great supports for me in times of my great needs. I would also thank my lab
peers Md. Tanvir Ibn Ashad and Md. Shamsuzzaman Julhas who always helped me
during my research work and made the time spent in the lab much more enjoyable and
fun. I really appreciate all the trouble that he two lab coordinators Mrs. Simi Tasnim

iii
Khan and Mrs. Rahela Zaman had gone through so much because of me. They always
took out their time for my work and guided me.
I would also like to thank my other friends, Tasnim Sultana and Sudeshna Nandi for
always encouraging and supporting me. Also, I thank my dearest and closest friend
Assadul Haque who always believed in me and constantly reminding me that I could
finish what I had started. Special thanks goes to Muntasir Hashim, who has been my
solid support spending countless hours helping me. He contributed greatly in my
writings by giving me his insights and correcting my mistakes. I can never truly
express how lucky I am to have him as a part of my life.
Finally I like to thank my parents and family for their constant inspiration and
blessings. I have come this far because of them and can proudly say that I have
achieved something I can stand tall to. This is for them and will eternally be theirs.

iv
Abstract
Tannase is an inducible enzyme, greatly affected in its production capacity when
considering environmental conditions. Industrial application of tannase in food,
beverage, pharmaceutical and bioremediation, has made it one of the most important
enzymes in research. Previous studies have revealed that tannase distribution in
bacteria is concentrated as genera specific in Gram-positive organisms within the
genera Bacillus, Lactobacillus, Staphylococcus, and few Gram-negative organisms in
the genera Klebsiella, Serratia, Pseudomonas. Present study has revealed the Gram-
positive phylum Firmicutes has the highest number of organisms with tannase
containing enzymes than any other phylum; further more Gram-negative tannase
containing bacteria are concentrated within the phylum Proteobacteria. The study also
identified organisms with tannase in genera Neisseria and Escherichia for the first
time.
The project focuses on the isolation, identification and the distribution of tannase
producing organisms in the bacterial kingdom. These tannase producers were isolated
from the fruit dump waste soil and were characterized through biochemical and
morphological experiments. Eleven new species were identified among which nine of
these belonged under the genus Staphylococcus. The distributions of these tannase
producing organisms in Staphylococcus were predicted by utilizing bioinformatics
methods.
The findings of this project opened up a new perspective to understanding the
evolution of the tannase protein and the distribution of this gene in the wider Gram-
negative and Gram-positive bacterial kingdom.

v
Table of Contents
Chapter :1 INTRODUCTION
1.1 Bacterial Enzymes................................................................................................ 1
1.1.1 Localization of Bacterial Enzymes ................................................................... 2
1.1.2 Types of enzymes used in the industry ............................................................. 4
1.2. Tannase................................................................................................................ 6
1.2.1. Tannase substrate ............................................................................................. 6
1.2.2. Sources of tannase............................................................................................ 8
1.2.3 Mechanism of tannase action............................................................................ 9
1.2.4 Regulation of tannase...................................................................................... 12
1.2.5 Molecular characteristics of tannase ............................................................... 12
1.2.5.1 The alpha/beta (α/β) hydrolase superfamily................................................. 13
1.2.5.2 The catalytic domain of the active site ……………………………….…....14
1.2.6 Industrial application of tannase……………………………………….……..15
1.2.6.1 Preparation of Instantaneous Tea................................................................. 15
1.2.6.2 Beverage clarification .................................................................................. 15
1.2.6.3 Gallic acid production for Pharmaceutical industry…………………….….15
1.2.6.4 Animal Feed Preparation………………………………………….………..16
1.2.6.5 Bioremediation of Tannin-Contaminated Wastewater…………….……….16
1.2.6.6 Other Potential Applications of Tannase………………………….………..16
1.3 Distribution of tannase in nature……………………………………….………18
1.3.1 Distribution of tannase in Gram-positive bacterial kingdom……….………..21
1.3.2 Distribution of tannase in Gram-negative bacterial kingdom………….…….22
1.4 Aims ans Objectives…………………………………………………….……...23
Chapter :2 METHODS AND MATERIALS
2.1 Introduction……………………………………………………………….……24
2.2 Media and stain preparations……………………………………….…………..26
2.2.1 Staining preparation: Gram’s stain……………………………….…………..26
2.2.2 Staining preparation: Endospore stain…………………………….………….27
2.2.3 Nutrient agar……………………………………………………….…………28
2.2.4 M-FC Agar………………………………………………………….………..29

vi
2.2.5 MacConkey agar…………………………………………………….……….30
2.2.6 Mannitol Salt agar…………………………………..………………………..31
2.2.7 Eosin Methylene Blue agar…………………………………………………..32
2.2.8 Starch agar………………………………………………………….………...33
2.2.9 Skim Milk agar…………………………………………………….…………34
2.2.10 Gelatin hydrolysis Medium……………………………………….………...35
2.2.11 Simmons Citrate Agar……………………………………...……….………36
2.2.12 Motility Indole Urease Agar……………………………….……….……….37
2.2.13 Triple Sugar Iron Agar……………………………………………………...38
2.2.14 MRVP medium……………………………………………………….……..40
2.2.15 Tannic acid supplemented media…………………………………………...42
2.3 Sample collection for tannase producing bacteria…………………….………..43
2.3.1 Environmental sample collection…………………………………….………44
2.3.2 Environmental sample serial dilution………………………………….……..45
2.3.3 Spread plate…………………………………………………………….…….45
2.3.4 Growth of colonies on Nutrient Agar (NA)………………………….………45
2.2.5 Growth of colonies on Tannic acid supplemented minimal media (TA).……45
2.3.6 Selection of colonies……………………………………………….………...45
2.4 E.coli isolation and identification……………………………………………...47
2.4.1 Environmental sample collection…………………………………………….48
2.4.2 Sewage waste water sample collection………………………………………48
2.4.3 Spread plate…………………………………………………………………..48
2.4.4 Streak plate and growth of colonies………………………………………….48
2.5 Biochemical test for bacterial identification…………………………………...49
2.5.1 Morphological test: Grams’ staining…………………………………………49
2.5.2. Morphological test: Endospore staining…………………………….…….....49
2.5.3 Catalase test………………………………………….……………………….50
2.5.4 Oxidase test…………………………………………………………………..50
2.5.5 MacConkey Agar Test……………………………………………….……….50
2.5.6 Mannitol Fermentation Test………………………………………………….51
2.5.7 Eosin Methylene Blue Agar Test…………………………………………….51
2.5.8 Starch hydrolysis test………………………………………………………...51
2.5.9 Skim Milk hydrolysis test…………………………………………….……...52
2.5.10 Gelatin hydrolysis test………………………………………………….…..52

vii
2.5.11 Citrate Test (Simmons Citrate Medium)…………………………………....52
2.5.12 Motility Indole Urease test………………………………………………….53
2.5.13 Triple Sugar Iron (TSI) Agar Test………………………………….……….53
2.5.14 Methyl Red Test and Voges-Proskauer Test………………………………..54
2.6 Secondary screening for tannase producing bacteria…………………………..55
2.7 Bioinformatics analysis for tannase enzyme identification…………………….56
2.7.1 Searching for tannase protein sequences in identified bacterial strains……...57
2.7.1.1 Protocol for searching ExPASy database to search for protein sequences...58
2.7.2 Finding homologue of a protein in particular genome…………………….....59
2.7.2.1 Blast search………………………………………………………….……...59
2.7.3 Finding the conserved domain family present in the homologue sequences
found through BLAST…………………………………………….………………..61
2.7.4 Methods for phylogenetic tree construction………………………………….62
2.7.4.1 Collection of 16S rRNA gene sequencing of organism from different
family……………………………………………………………………………….62
Chapter :3 RESULTS
3.1 Control Optimization…………………………………………………………...64
3.1.1 Bacterial strain for optimization……………………………………………...64
3.1.2 Bacterial biochemical identification………………………………………….64
3.1.3 Tannase activity presence in identified control strain………………………..66
3.2 Natural strains isolation for tannase activity…………………………………...67
3.2.1 Natural samples………………………………………………………………67
3.2.2 Soil characterization………………………………………………………….67
3.2.3 Isolation of tannase containing bacterial strain from natural soil samples…...68
3.3 Growth and isolation of pure culture…………………………………………...70
3.3.1 Sub-culture of the tannase positive isolated colonies………………………...70
3.3.2 Selection of isolated colonies………………………………………………...70
3.4 Biochemical tests for the tannase positive isolated colonies…………………...71
3.4.1 GRAMS’s stain test…………………………………………………………..71
3.4.2 Catalase test…………………………………………………………………..72
3.4.3 Oxidase test…………………………………………………………………..72
3.4.4 Growth of selective agar MacConkey agar…………………………………..73
3.4.5 Growth of selective agar Mannitol Salt agar………………………………....74

viii
3.4.6 Growth on Starch hydrolysis agar……………………………………………75
3.4.7 Growth on Skim Milk agar…………………………………………………...76
3.4.7 Citrate test…………………………………………………………………….77
3.4.8 Motility Indole Urease test…………………………………………………...78
3.4.9 Triple Sugar Iron test…………………………………………………………79
3.4.10 Methyl Red and Voges-Proskauer test……………………………………...80
3.4.11 Summary of the Biochemical test results…………………………………...81
3.4.12 Biochemicaltest identification………………………………………………83
3.4.13 Tannase activity in isolated strains………………………………………….84
3.5 Isolated E.coli strain Biochemical test result…………………………………..85
3.5.1 Growth on M-FC agar………………………………………………………..85
3.5.2 Growth on Eosin Methylene Blue agar………………………………………85
3.5.3 Growth on MacConkey agar…………………………………………………86
3.5.4 GRAMS’s stain test………………………………………….……………….86
3.5.5 Catalase test…………………………………………………………………..87
3.5.6 Oxidase test…………………………………………………………………..87
3.5.7 Mannitol fermentation test…………………………………………………...88
3.5.8 Starch hydrolysis test………………………………………………………...88
3.5.9 Gelatin hydrolysis test………………………………………………………..89
3.5.10 Citrate test………………………………………………………………......89
3.5.11 Motility Indole Urease test………………………………………………….90
3.5.12 Triple Sugar Iron test………………………………………………………..91
3.5.13 Methyl Red and Voges-Proskauer test……………………………………...92
3.5.14 Summary of the Biochemical test results for E.coli………………………...93
3.5.16 Tannase activity in isolated E.coli strain……………………………………95
3.6. Bioinformatics analysis for tannase enzyme identification……………………96
3.6.1 Finding the domain of tannase protein……………………………………….96
3.6.2 Selection of reference tannase protein for query……………………………..99
3.6.3 Predicting tannase proteins in sample organisms/species…………………..101
3.6.4 Distribution pattern of the tannase protein within the bacterial kingdom…..103
3.6.4.1 Distribution of tannase protein in Gram-positive bacteria………………..103
3.6.4.1.1 Distribution among the Staphylococcus genera………………………...103
3.6.4.2 Distribution among the wider Gram-Positive bacterial kingdom………...105

ix
3.6.4.3 Distribution among the Gram-Negative bacteria…………………………106
Chapter :4 DISCUSSION
4.1 Optimization..................................................................................................... 108
4.1.1 System optimization...................................................................................... 108
4.1.2 Control optimization ..................................................................................... 109
4.1.3 Primary and secondary screening.................................................................. 110
4.2 Environmentsl samples .................................................................................... 111
4.2.1 Isolated bacterial colonies from the soil and their biochemical test for
identification………………………………………………………………………111
4.2.2 The tannase enzyme………………………………………………………...115
4.2.3 Prediction of presence of tannase in the isolated Staphylococcus strains…..116
4.2.4 Isolated E.coli strain from the sewage and the biochemical test for
identification………………………………………………………………………117
4.3 Tannase distribution pattern in the bacterial kingdom………………………..119
4.3.1 Distribution of tannase in the Staphylococcus genera………………………119
4.3.2 Distribution of tannase in wider Gram-positive bacterial kingdom………...120
4.3.3 Distribution of tannase in wider Gram-negative bacterial kingdom………..121
4.9 Future Study:………………………………………………………………….122
REFERENCES.....................................................................................................123
ANNEXURE.…………………………………………………………………….132

x
List of Figures
Figure Number Figure List Page number
Figure 1.1 Classification of microbial enzymes based on spatial
localization.
2
Figure 1.2 Chemical structures of tannin classification 6
Figure 1.3 The ester and depside bonds present in tannins 9
Figure 1.4 Methyl gallate and epicatechin gallate hydrolyzed to
gallic acid, epicatechin and methanol in (a), digallic
and epigallocatechin gallate hydrolyzed to gallic acids
and epigallocatechin in (b) and rosacyanin hydrolyzed
to degalloylated rosacyanin in (c).
10
Figure 1.5 The reaction mechanism of tannase 11
Figure 1.6 The canonical diagram of alpha/beta hydrolase fold
showing also the catalytic triad
13
Figure 1.7 Gram-positive bacterial kingdom tannase distribution
shown with Red lines.
21
Figure 1.8 Gram-negative bacterial kingdom tannase distribution
shown with Red lines.
22
Figure 2.1 Flowchart summarizing the overall research
methodology.
25
Figure 2.2 Overview of the isolation and identification of
Tannase producing bacteria from soil.
43
Figure 2.3 Overview of isolation and identification of E.coli from
sewage water.
47
Figure 2.4 Bioinformatics analysis to map the conserve domain
with the sequences
56
Figure 2.5 Searching ExPASy database where in right side shows
links to many different resources.
58
Figure 2.6 NCBI- BLAST search home page showing Genome
BLAST. Microbes (Red outlined box) was selected as
is goes directly to microbe genome BLAST.
60
Figure 2.7 Searching homologue of a protein from NCBI microbe
genome protein BLAST, with query sequence in the
search box (Red arrow) and selected organism genome
highlighted in the Red outlined box.
60
Figure 2.8 NCBI conserve domain BLAST 61
Figure 2.9 ExPASy proteomics tool ProDom. 61
Figure 2.10 16s rRNA sequence obtained from LPSN database. 63

xi
Figure 2.11 Finding 16s rRNA sequence from Genbank. 63
Figure 3.1 Growth and zone of hydrolysis of E.coli DH5α
showing positive result (right side of both the plates)
while S. aureus and B.cereus showing negative result
with no growth and zone of hydrolysis (left side of
both the plates) after adding FeCl3 solution on the
media to observe zone of hydrolysis.
66
Figure 3.2 Comparison between the numbers of colonies of the
different serial dilutions of the soil in Tannin
supplemented minimal media (TA) and Nutrient agar
(NA) (control).
69
Figure 3.3 16 colonies that have been sub-cultured from the serial
dilution plates (10-3
and 10-4
) in nutrient agar (NA) to
obtain pure culture Tn stands for sample identifier.
70
Figure 3.4 Gram’s stained cells showing Gram-positive (purple)
cocci in clusters and cocci isolated in 3.4a (i &ii) and
Gram-negative cocci in clusters and cocci isolated in
3.4b (i &ii).
71
Figure 3.5 Formation of bubbles showing the presence of catalase
enzyme.
72
Figure 3.6 T5 and T7 showing oxidase positive with purple blue
colony 3.6(a) (on the left side) and T1, T2, T3, T4, T8,
T9, T10, T11, T13, T14, T15, T16 showing oxidase
negative with colorless colony 3.6(b) (on the right
side).
72
Figure 3.7 Growth on MacConkey agar with color change to
bright pink colonies in both (a) and (b). Tn represents
the respective isolated 14 strains.
73
Figure 3.8 Growth of colonies showing salt-tolerance and media
changing color to bright yellow showing mannitol
fermentation in both (a) and (b). Tn represents the
respective isolated 14 strains.
74
Figure 3.9 No zone of hydrolysis showing no presence of α-
amylase enzyme both in (a) and (b) after the addition
of Gram’s Iodine.
75
Figure 3.10 No zone of hydrolysis showing no production of
ceasinase in both (a) and (b). Tn represents the
76
Figure 3.11 Citrate utilization causing the change in media color
from green to blue (a) and (b). Tn represents the
77

xii
Figure 3.12 Motility Indole Urease test result of all the 14 strains.
Tn represents the respective isolated 14 strains. Some
of the strains such as T1, T4, T5, T8, T9, T14 and T15
may seem as though they are negative for urease test,
but this is merely due to the addition of indole reagent
which caused the light pink color to slowly fade.
78
Figure 3.13 Triple sugar iron test result for 14 strains. Tn
represents the respective isolated 14 strains.
79
Figure 3.14 MR-VP test result for 14 strains. Tn represents the
80
Figure 3.15 Tannase activity showed by the strains isolated in this
study. After growth and incubated secondary screening
is done by adding FeCl3 solution which form
complexes with tannin present in the media, while
showing hydrolysis zone due to presence of tannase.
84
Figure 3.16 Colonies of fecal coliform in M-FC agar with blue
colonies.
85
Figure 3.17 Colonies in EMB agar showing metallic green sheen
with black centers (characteristic of E.coli).
85
Figure 3.18 Colonies in MacConkay agar with pink colonies
showing lactose fermentation.
86
Figure 3.19 Gram’s stained cells showing Gram-negative (pink)
rods.
86
Figure 3.20 Formation of bubbles showing the presence of catalase
enzyme.
87
Figure 3.21 Colony with no color change after the addition of
Kovacs™ Oxidase reagent.
87
Figure 3.22 Colony with no color change as there is no mannitol
fermentation.
88
Figure 3.23 Colony with no zone of hydrolysis as no production of
extracellular enzymes α-amylase. Comparison is made
between the control DH5α strain and the isolated
E.coli strain to observe the zone of hydrolysis.
88
Figure 3.24 Colony with no zone of hydrolysis of the nutrient
gelatin as there is no liquefaction of the media.
89
Figure 3.25 Colony with no change in color showing citrate not
utilized.
89
Figure 3.26 Colony with no change within the media as it is unable
to produce urease enzyme that is capable of
hydrolyzing urea. There is also diffused growth of the
colony within the media as the organism is motile and
90

xiii
thus moves around the media. After the addition of
Kovacs’s™ Indole reagent there is a formation of
cherry red ring on top of the media.
Figure 3.27 Colony capable of fermenting glucose, lactose and
sucrose as the media turns yellow due to production of
acid and the pH lowering. Gas is also produced as
there are cracks within the media and it moves
upwards from the bottom where the gas production is
the most.
91
Figure 3.28 Colony showing MR and VP test results. The methyl
red test is positive as shown in (a) due to the media
remaining red after the addition of methyl red solution.
The Voges-Proskauer test giving negative result as the
broth remains yellow after the addition of α- napthol
followed by KOH shown in (b).
92
Figure 3.29 Growth and zone of hydrolysis of E.coli showing that
it has tannase producing ability.
95
Figure 3.30 Phylogenetic tree of Staphylococcus genera where Red
lines showing the strains identified in this study, Aqua
lines showing organism with predicted tannase protein
and Green lines showing the presence of esterase/
lipase. Navy blue showing the negative control used in
this study with absence of tannase protein.
104
Figure 3.31 Distribution of tannase among the Gram-negative
phyla with the Red lines showing its presence among
the families. And the Magenta line showing the family
with the tannase activity found in this study.
107

xiv
List of Tables
Table Number Table List Page Number
Table: 1.1 Classification of enzymes based on their reaction
types.
4
Table: 1.2 Distribution of tannase producing bacteria 19
Table: 2.1 Ingredients for Gram’s staining. 26
Table: 2.2 Ingredients for Endospore staining. 27
Table: 2.3 Ingredients for Nutrient Agar 28
Table: 2.4 Ingredients for M-FC Agar Base 29
Table: 2.5 Ingredients for MacConkey Agar 30
Table: 2.6 Ingredients for Mannitol Salt Agar 31
Table: 2.7 Ingredients for Eosin Methylene Blue Agar 32
Table: 2.8 Ingredients for Starch Hydrolysis Agar. 33
Table: 2.9 Ingredients for Skim Milk Agar. 34
Table: 2.10 Ingredients for Gelatin hydrolysis Medium. 35
Table: 2.11 Ingredients for Simmons Citrate Agar 36
Table: 2.12 Ingredients for Motility Indole Urease Agar 37
Table: 2.13 Ingredients for Triple Sugar Iron Agar 39
Table: 2.14 Ingredients for MRVP Medium 41
Table: 2.15 Ingredients for Tannic acid supplemented minimal
media.
42
Table: 3.1 Biochemical test results of the control strains. 65
Table: 3.2 Different colonies in different serial dilutions in both
Nutrient agar plate (NA) and Tannin supplemented
minimal media (TA).
68
Table: 3.3 Summary of the biochemical test results of the
selected tannase positive strains.
81
Table: 3.4 Biochemically identified bacterial strains 83

xv
Table Number Table List Page Number
Table: 3.5 Summary of the biochemical test results of the E.coli
strain isolated.
94
Table: 3.6 Bacterial Tannase protein sequences catalytic
domains and their protein family
98
Table: 3.7 Tannase protein sequences of the query. 100
Table: 3.8 Putative sequences with tannase domain activity
identified within the sequences obtained for the
identified bacterial strains after genome BLAST and
ProDom search.
102

xvi
List of Abbreviations
U: urease
SH: Starch hydrolysis agar
GH: Gelatin hydrolysis agar
MR: Methyl red
VP: Voges-Proskauer
TSI: Triple sugar iron
G: Glucose
L: Lactose
S: Sucrose
MIU: Motility indole urease
M: motility
I: indole
MAC: MacConkey agar
MSA: Mannitol salt agar
EMB: eosin methylene blue agar
CT: Catalase test
OT: Oxidase test
CU: citrate utilization
α/β hydrolase: alpha/beta hydrolase

1
1.1 Bacterial Enzymes
Enzymes are biocatalysts that are produced by a living organism which acts to bring
about a specific biochemical reaction (Gurung, et al., 2013). They are employed
largely in different industrial processes for a large number of reactions as they offer
many advantages over chemical catalyst particularly; the microbial enzymes which
have widespread uses in various industries. The global market has an immense need
for industrial enzymes that is estimated to be worth about 3.3 billion dollars in 2010.
This market is expected to reach more than 4 billion dollars by 2015 (Gurung, et al.,
2013 and Jana, et al., 2013). Recently, with the evolution of green biotechnology the
relevance of enzyme for the production of chemicals, fuels, bioremediation, food
industry, secondary metabolites, and other industries has been highly promoted (Jana,
et al., 2013).
The majorities of industrial enzymes used nowadays are of microbial origin as for
one, the microbial enzymes are generally more active and stable than plant and animal
enzymes also exemplifying an alternative source of enzymes because they can be
cultured in large quantities in a short amount of time by fermentation. Thus, with the
development of fermentation processes that targeted particularly for the production of
microbial enzymes by use of specifically selected strains, it is now possible to
produce purified, well-characterized enzymes on a large scale basis. This
development granted the access of microbial enzymes into genuine industrial products
and processes, for example, within the detergent, textile and food industries (Gurung,
et al., 2013).
Second, owing to their biochemical diversity and susceptibility to gene manipulation,
the microbial enzymes are growing more desirable for the industry as well (Alves, et
al., 2014). The recombinant DNA technology allowed the production of enzymes that
were not commercially produced previously by meliorating the production process. In
addition, the growth in biotechnology, such as protein engineering and directed
evolution, encouraged the revolution of the commercialization of industrial important
enzymes. Here, enzymes play the key roles in numerous biotechnology products and
processes that are commonly encountered in the production of food and beverages,
cleaning supplies, clothing, paper products, transportation fuels, pharmaceuticals, and
monitoring devices. This advancement in biotechnology is providing different kinds

2
of enzymes exhibiting new activities, adaptability to new conditions leading to their
increase use in industrial purposes (Gurung, et al., 2013).
Apart from this the evolution of microorganisms coupled with selective pressures in
different habitats has produced their incomparable physiological and biochemical
diversity, in which enzymes play a key role in microbial adaptation and evolution
(Alves, et al., 2014). So, industries are now looking for new microbial strains in order
to produce the highest quality of different type enzymes to fulfill the current enzyme
requirements (Gurung, et al., 2013).
1.1.1 Localization of Bacterial Enzymes
Enzymes are ubiquitous, meaning that they are essential components for animals,
plants and microorganisms, catalyzing and coordinating the complex biochemical
reaction required for cellular metabolism and survival. The localization of these
enzymes is therefore an important factor, not only for the cell’s metabolism but also
for their application in the industry. Bacterial enzymes are mainly localized in two
different regions as shown in the Figure 1.1 below.
Figure 1.1: Classification of microbial enzymes based on spatial localization.
Intracellular or Cytoplasmic: intracellular enzymes are the common metabolic
enzymes that are responsible for catalyzing all those processes needed within the
cells. These enzymes are responsible for the release of energy from nutrients such as
glucose or carbohydrates, lipids, proteins. DNA synthesis and replication are also
Bacterial enzymes
Intracellular enzymes Extracellular enzymes
Ecto-enzyme
Truly Extracellular
enzyme

3
done by these enzymes, thus they ensure mostly the bacterial cell metabolism and
survival.
Extracellular or Secretory: while many enzymes are retained within the cell, others
are released into the surrounding environment. In some cases, microbes produce small
amounts of extracellular enzymes, regardless of substrate availability, as a mechanism
to detect substrate within the environment. But, if the substrate is present, these
constitutive enzymes produce signals that induce additional enzyme synthesis and
secretion. Since the cell cannot transport complex substrate thus, they are first
hydrolyzed outside of the cell into simpler components which are then transported in
(Cunha, et. al., 2010 and Burns et. al., 2010) as the passive transport through bacterial
cell wall and cell membrane is restricted to very small and chemically simple
compounds. For Gram-negative bacteria, extracellular enzymes are contained within
the periplasmic space, associated with the outer cell wall, or released into the outside
(Burns et. al., 2010). Presence of trimeric proteins (porins) are also there as channels
between the outer membrane and the periplasmic space. For Gram-positive bacteria,
the cell wall is not as restrictive, in terms of permeability, as it is in the case of the
outer membrane of gram negative bacteria (Cunha, et. al., 2010). Extracellular
enzymes are further classified based on their physical relation with the cell as
ectoenzymes and truly extracellular enzymes.
1. Ectoenzymes are associated with living cells and include enzymes inserted in
or spanning the plasma membrane with transmembrane domains, associated
with the cell wall. These enzymes, in Gram-negative bacteria, are attached to
the outer membrane surface or retained within the periplasmic space (Cunha,
et. al., 2010).
2. Strict-sense or truly extracellular enzymes occur in free form and catalyze
reactions detached from their producers. Bacterial extracellular enzymes may
be actively secreted by intact viable cells, into the environment by viral lysis
(Cunha, et. al., 2010).
The majority of enzymes used for industrial application purpose are of extracellular or
secretory type. The main reason behind this is due to them being easier to extract thus
cheaper to purify for commercial use. Also large amount of the enzyme can be
obtained. When it comes to intracellular enzymes, there is an extra downstream step

4
for cell lysis and purification, with the extraction of very little amount of the enzyme
when compared to that obtained from the secretory type.
1.1.2 Types of enzymes used in the industry
Enzymes are highly selective and specific catalysts which can significantly accelerate
both the rate and specificity of metabolic reactions (Gurung, et al., 2013). Most
enzymes are much larger than the size of the substrates they act upon, and only a
small portion of the enzyme (around 2 to 4 amino acids) organized in precise three-
dimensional arrangement is directly involved in catalysis. Enzymes are classified
according to their nature of catalyzing a reaction into six different groups shown in
Table 1.1 below.
Table 1.1: Classification of enzymes based on their reaction types. (Gurung, et al,
2013)
Class Description Important subclass
Oxidoreductase
(EC1)
Transfer of reducing equivalent hydrogen
and electron from one redox system to
another.
Dehydrogenase,
oxidase, reductase
Transferase
(EC2)
Transfer of other group from one
molecule to another.
Glycosyl-transferase,
amino-transferase
Hydrolase
(EC3)
Cleaves bond using water molecule. Easterase, peptidase
Lyase
(EC4)
Cleaves bonds by elimination, leaving
double bonds or rings, or conversely
adding groups to double bonds, this
cleavage does not require water like
hydrolase.
C-C lyases, C-N
lyases, C-O lyases
Isomerase
(EC5)
Moves groups within the molecule thus
giving rise to isomers of the molecule.
Epimerase, isomerase
Ligase
(EC 6)
Catalyzes ligation and are energy
dependant therefore always coupled with
hydrolysis of nucleoside triphosphates.
C-C ligases, C-N
ligases, C-O ligases

5
Currently, the dominant types of enzymes in industrial application are classified as
“hydrolase”, which are being used for the degradation of various natural substances.
The pivotal enzyme type here remains Proteases, because of their wide use in the
detergent and dairy industries. These enzymes catalyze the cleavage of peptide bonds
in other proteins, and microorganisms elaborate a large array of intracellular and/or
extracellular proteases (Alves, et al., 2014). Various hydrolases that degrades
carbohydrates, such as amylases and cellulases, are also used in industries such as the
starch, textile, detergent, and baking industries, representing the second largest group
(Gurung, et al., 2013).
Another group of hydrolytic enzymes includes esterases, which catalyze the cleavage
and formation of ester bonds and are known as α/β-hydrolases (Alves, et al., 2014);
tannase belongs to this class of enzymes.

6
1.2 Tannase
Tannase or tannin acly hydrolase (E.C.3.1.1.20), is an inducible microbial enzyme
that catalyzes the hydrolysis of ester and depside bonds between varied substrate like
gallo-tannin, epigallocatechin-3-gallate, gallic acid esters and hydrolysable tannins to
release gallic acid and glucose (Jana, et al., 2013, Belur, et al., 2011 and Banerjee, et
al., 2012).
1.2.1 Tannase substrate
The substrates for tannase are tannins which are naturally occurring plant phenolic
compound distributed in different parts of the vascular plants (Belmares, et al., 2004).
Tannins are the fourth most abundant plant constituents coming after cellulose,
hemicelluloses and lignin and are mostly accumulated in the vulnerable parts of the
plants, where they provide immunity to the plants from microbial attacks (Mohapatra,
et al. 2006). Thus tannins are considered to be anti-microbial by nature, as it is toxic
to animals and inhibit the growth of a number of microorganisms (Pepi, et al., 2009).
However, many microorganisms have developed mechanisms to overcome the effects
of tannins which include tannin modification, degradation, and dissociation of tannin-
substrate complexes, tannin inactivation by high affinity binders, membrane
modification, and metal ion sequestration (Belur, et. al., 2010 and Smith et al., 2005).
Tannins have the ability to precipitate macromolecules such as protein, cellulose,
gelatin and starch by forming non-reversible complexes with them (Belur, et al.,
2011). Currently tannins have been classified in four groups, shown below Figure 1.2.
Figure 1.2: Chemical structures of tannin classification (Aguilar, et al., 2007).

7
1. Hydrolysable tannins or Gallotannins are the simplest form of these
polyphenols characterized by the presence of several molecules of organic
acids, such as gallic, digallic, and chebulic acids, esterified to a molecule of
glucose. Gallotannins can be easily hydrolyzed under mild acid or alkaline
conditions, either in hot water or enzymatically thus tannase generally act
upon gallotannins (Aguilar, et al., 2007 and Ashok, et al., 2012). Tannic acid
is an example of this type on tannin.
2. Ellagitannins, on the other hand have building blocks made of ellagic acid
units linked to glucosides. Molecules with a core of quinic acid instead of
glucose have also been considered as ellagitannins and so are more stable than
gallotannins (Aguilar, et al., 2007 and Ashok, et al., 2012).
3. Complex tannins are generated through reactions between several units of
gallic or ellagic acids with catechins and glucosides. An example of this kind
of tannin is catechin- gallate with hydrolysable and condensed bonds (Aguilar,
et al., 2007).
4. Condensed tannins or proanthocyanidins are oligomeric and polymeric
complex compounds made of flavonoid building blocks (from 2 over 50) that
are not considered to be easily hydrolyzable. Among the major constituents
are catechin derivatives such as cyaniding and delphinidin, which are
responsible for the astringent taste of fruit and wines. Condensed tannins are
not hydrolyzed by “classical tannases”, with initial degradation steps carried
out by mono- or di- oxygenases (Aguilar, et al., 2007 and Ashok, et al., 2012).
All tannins, especially the hydrolysable ones are very soluble in water

8
1.2.2 Sources of tannase
Tannase can be obtained from tannin rich plants and animal tissues, but, for its
industrial purpose, microbial sources are preferred as these enzymes are usually more
stable than their plant or animal counterparts. In addition, the fermentation process
can produce large amounts of enzymes in a constant environment and can be
controlled more easily (Rodrıguez-Duran, et al., 2011). The major sources of tannase
producers are mainly from the two groups: Fungi and bacteria.
1. Fungi: the first tannase protein was extracted and identified from the fungi
strain now known as Aspergillus niger. Apart from this several fungal tannase
have also been indentified from the genus Aspergillus, Penicillium,
Trichoderma, Fusarium, Paecilomyces and Rhizopus (Belur, et al., 2011).
2. Bacteria: over the last 25 years a number of bacterial strains have been
isolated that contained tannase activity. Several species were identified among
which organisms from the genus Bacillus, Lactobacillus, Staphylococcus,
Serratia, Pseudomonas and some of the genera falling in the
Enterobacteriaceae family are predominant (Belur, et al., 2011 and Jana, et al.,
2013).

9
1.2.3 Mechanism of tannase action
Tannase has been reported to have dual activities catalyzing the hydrolysis ofester
bonds (galloyl ester of an alcohol moiety) and depside bonds (galloyl ester of gallic
acid) present in gallotannins, complex tannins, and gallic acid esters but they do not
affect the carbon-carbon bonds, which is why tannase is unable to hydrolyze
condensed tannins (Haslam, et al., 1966, Rodrıguez-Duran, et al., 2011, Belur, et al.,
2011 and Ren, et al., 2013). The Figure 1.3 below illustrates the bonds that are
hydrolyzed by tannase.
Figure 1.3: The ester and depside bonds present in tannins (Haslam, et al., 1966).
Tannase from different sources, have different molecular masses such as in the case of
fungi and yeasts are glycoproteins and often form hetero- or homo-oligomers with
two to eight subunits. Bacterial tannases on the other hand, exist mainly as monomers
(Ren, et al., 2013).
The dual activities of tannase cause this enzyme to have a wider range of substrate
specificity. This specificity depends on the source and the methods utilized for its
production and isolation (Rodrıguez-Duran, et al., 2011 and Ren, et al., 2013). Apart
from it being a bi-functional enzyme, tannase also exists as isoenzymes. Tannase
hydrolyzes other substrates such as methyl gallate, propyl gallate, digallic acid,
epicatechin gallate, and epigallocatechin gallate-releasing gallic acid (Curiel, et al.,
2009). Tannase also acts on ellagitannins such as rosacyanin or phyllanemblinin. In
those cases, tannase selectively hydrolyses the galloylmoieties, yielding gallic acid
and degalloylated ellagitannins (Lu, et al., 2007). The different types of substrates
hydrolyzed by tannase are shown in Figure 1.4 (a), (b) and (c) below (Rodrıguez-
Duran, et al., 2011).

10
Figure 1.4: Methyl gallate and epicatechin gallate hydrolyzed to gallic acid,
epicatechin and methanol in (a), digallic and epigallocatechin gallate hydrolyzed
togallic acids and epigallocatechin in (b) and rosacyanin hydrolyzed to degalloylated
rosacyanin in (c).
Tannase are a family of serine esterases, with a catalytic triad having its serine residue
presentin the conserved pentapeptide motif (-Gly-X-Ser-X-Gly-) which is necessary
for its catalytic activity (Rodrıguez-Duran, et al., 2011 and Ren, et al., 2013). The
enzyme’s mechanism of action was best described by Ren, et al., (2013) for the
Lactobacillus plantarum tannase. After the substrate binds to the enzyme, the
(a)
(b)
(c)

11
hydroxyl group of Ser163 starts a nucleophilic attack on the carbonyl unit of the
galloyl unit. This attack is assisted by His451 that acts as a general base. This causes
the formation of a tetrahedral intermediate, stabilized by hydrogen-bonding
interactions with Gly77 and Gly164 that form the oxyanion hole. His451-H acts as a
general acid, the tetrahedral intermediate then collapses to produce the alcohol
product and the acyl-enzyme intermediate. A water molecule is then activated by
His451 to attack the acyl-enzyme to form the second tetrahedral intermediate, which
then collapses to release gallic acid and regenerate the enzyme. This mechanism is
shown in Figure 1.5.
Figure 1.5: The reaction mechanism of tannase (Ren, et al., 2013).

12
1.2.4 Regulation of tannase
Tannase is an inducible enzyme, and the presence of tannins causes the increase of its
production. Although there is a contradiction regarding this as according to Mondal,
et al., (2000) low levels of glucose, lactose and sucrose were not repressive of the
enzyme production but did repress in higher concentration. Whereas according Sabu
et al., (2006) ant carbon source present in the media other than tannic acid as a carbon
source inhibits the production of tannase.
1.2.5 Molecular characteristics of tannase
The catalytic function of an enzyme is greatly dependant on the molecular structure of
the protein. Although a great variety has been seen in case of the fungal and bacterial
tannase where most of the fungal tannase had their alpha/beta hydrolase domain in
association with feruloyl esterase while most of the bacterial only had the alpha/beta
hydrolase domain. Tannase (E.C. 3.1.1.20) and feruloyl esterase (E.C. 3.1.1.73)
belong to the same protein family (Pfam IPR011118 and Pfam PF07519), but have
different catalytic functions. This enzyme family is expressed by certain bacteria and
fungi, many of which are plant pathogens. The enzymes hydrolyze the ester bonds of
hydrolysable tannins and feruloyl-polysaccharides, releasing the bound
macromolecules that were previously indigestible. At the sequence level though
tannase and feruloyl esterase are currently indistinguishable, causing confusion when
annotating the genes without functional characterization. The majority of the tannase
sequences on record are often given both names but while both have similar function,
they act upon different plant polyphenols (Udatha, et al., 2012).
Tannase, when characterized through protein family search belonged to the serine
esterase family (Ren, et al., 2013 and Jana, et al., 2014). The serine esterase family
has a conserved serine residue within the pentapeptide motif (-Gly-X-Ser-X-Gly-)
which is part of the catalytic triad consisting of serine, aspartic acid and histidine.
When tannase sequences of bacterial strains were compared one common link was
their relationship with the alpha/beta (α/β) hydrolase superfamily which could be
explained as these protein evolved from a “common ancestor” (Nardini, et al., 1999).

13
1.2.5.1 The alpha/beta (α/β) hydrolase superfamily
The α/β hydrolase superfamily of enzymes consists of α/β hydrolase fold within their
catalytic domains. It includes the family of proteases, lipases, esterases,
dehalogenases, peroxidases and epoxide hydrolases, making it one of the most
versatile and widespread protein folds to be discovered. The typical α/β hydrolase fold
has been described to be consisting of a mostly parallel, eight-stranded β sheet
surrounded on both sides by α helices with onlythe second β strand being antiparallel
(Nardini, et al., 1999). Although, differences may be also present in the spatial
position of the α helices connecting the β strands of the central β sheet. In some cases,
one or more of these helices may even be completely absent. Only helix α C appears
to be well conserved; as it has a strategic position in the center of the β sheet and
plays an important role in the correct positioning of the nucleophilic residue within
the active site (Nardini, et al., 1999). The canonical secondary structure diagram of
α/β hydrolase fold is shown in Figure 1.6 below.
Figure 1.6: The canonical diagram of alpha/beta hydrolase fold showing also the
catalytic triad (Nardini, et al., 1999).
According to the crystal structure study of Lactobacillus plantarum tannase by Ren et
al., (2013), only half of the bacterial tannase is composed of the α/β hydrolase fold.
Fungal tannase on the other hand also have a functional domain belonging under the
feruloyl esterase. Thus when comparison was made between the fungal and the
bacterial tannase very little sequence similarity was found but not only that even when
different tannase sequences of several bacterial strains were also compared very little
significant sequence similarity was also found (Ren, et al., 2013 and Jana, et al.,

14
2014). Apart from this the tannase α/β hydrolase fold also consists of a cap domain
involved in substrate binding. Whereas in case of lipase this flap is usually
amphipathic and directly covers the catalytic site, while for the feruloyl esterases,
however, the catalytic site is usually exposed.
1.2.5.2 The catalytic domain of the active site
Tannase protein sequences of different bacterial genus and species also have a varied
different other protein domains present along with the α/β hydrolase domain such as
carboxylesterase/thioesterase, peptidase, proline iminopeptidase like, etc (Jana, et al.,
2014). But regardless of having these domains the catalytic functional domain is still
represented by the α/β hydrolase domain which consists of a catalytic triad build up of
serine, aspartic acid and histidine residues. The nucleophile (serine) is located in a
very sharp turn, called the ‘nucleophile elbow’, where it can easily approach the
substrate, as well as the hydrolyticwater molecule. The nucleophile elbow is identified
by the consensus sequence motif Sm-X-Nu-X-Sm (Sm = small residue, X = any
residue and Nu = nucleophile). The tightness of this strand-turn-helix motif induces
the nucleophilic amino acid residue to adopt an energetically unfavorable main chain
torsion angles and imposes steric restrictions on residues located in its proximity. The
geometry of the nucleophile elbow also contributes to the formation of the oxyanion-
binding site, which is necessary to stabilize the negatively charged transition state that
occurs during hydrolysis process (Nardini, et al., 1999).

15
1.2.6 Industrial application of tannase
1.2.6.1 Preparation of Instantaneous Tea
Tea is the second most highly consumed beverage worldwide after water. During the
production of tea beverages, hot and clear tea infusions tends to form turbid
precipitates after cooling. These precipitates, called tea cream, are formed by a
complex mixture of polyphenols. Tea polyphenols also form hydrogen bonds with
caffeine, which also leads to the cream formation. This cream formation is a quality
problem and may have anti-nutritional effects. Thus when treated with tannase, it can
hydrolyze the ester bonds of catechins to release free gallic acid and water-soluble
compounds with lower molecular weight, reducing turbidity and increasing solubility
of tea beverage in cold water. This treatment of tea beverage leads to a better color
appearance, less cream formation, better taste, mouth feeling, and overall acceptance.
Also tannase treated green tea shows higher antioxidant properties than normal green
tea or black tea (Rodrıguez-Duran, et al., 2011 and Belur, et al., 2011).
1.2.6.2 Beverage clarification
New fruit juices (pomegranate, cranberry, raspberry, etc.) have recently been
acclaimed for their health benefits, in particular because of their antioxidant
properties. However, the presence of high tannin content in those fruits is responsible
for haze and sediment formation, as well as for color, bitterness, and astringency of
the juiceupon storage. When enzymatically treated with tannase this improve the
quality of these juices. Tannase is used as clarifying agent in refreshing drinks with
coffee flavor, and recently, this process for the enhancement of the antioxidant
properties of coffee by using tannase and other enzymes has also been patented
(Belur, et al., 2011).
1.2.6.3 Gallic acid production for Pharmaceutical industry
One of the major applications of this enzyme is in the production of gallic acid. Gallic
acid (3,4,5-trihydroxybenzoic acid) is a phenolic compound and the monomeric unit
of the gallotannins and complex tannins. Gallic acid and its related compounds
possess many potential therapeutic properties including anticancer and antimicrobial
properties. Its major application is in the area of manufacturing the antibacterial agent
trimethoprim. It is also used inleather industry, in manufacturing gallic acid esters,

16
such aspropyl gallate, a potent antioxidant utilized as antioxidantin fats and oils.
Gallic acid is also used in the manufacture of pyrogallol and as a photosensitive resin
in semiconductor production as well as ink and photographic developer (Belur, et al.,
2011, Weetal, et al., 1985 and Raghuwanshi, et al., 2011).
1.2.6.4 Animal Feed Preparation
High levels of dietary tannins have negative effects on animal nutrition; as tannin bind
to macromolecules. Tannins form strong complex with enzymes, minerals, and other
nutrients. They are also responsible of a bitter taste, which considerably reduces the
feed intake. Tannins are ubiquitous in nature and are widely found in feedstuffs,
forages, fodders, and agro-industrial wastes, affecting livestock production. This
antinutritional effect of tannin can be reduced by treating with tannase or tannase
producing microorganism. Enzymatic extract containing tannase when applied to
several flours used as animal feed (barley, bran, maize, oat, rye, soya, and wheat
flour) this released similar amounts of reducing sugars from all flours when compared
with a commercial enzymatic additive used in animal feeding. These observations
concluded that tannase-containing preparation has a high potential as supplements for
animal feeding (Rodrıguez-Duran, et al., 2011).
1.2.6.5 Bioremediation of Tannin-Contaminated Wastewaters
Tannins occur commonly in the effluents derived from several agro-industries one
such includes tanneries. The treatment of this kind of wastewaters is usually difficult
because tannins are highly soluble and inhibit the growth of many microorganisms.
Tannase can be potentially used for the degradation of tannins in these effluents.
Enzymatic treatment of tannery wastewater removed about 42% of the tannin content
and 20% of the color. These findings suggest than tannase or tannase producing
microorganism could be utilized for a pretreatment of tannin-rich wastewaters
(Rodrıguez-Duran, et al., 2011 and Murugan, et al., 2010).
1.2.6.6 Other Potential Applications of Tannase
Ethanol as a fuel production from agro-industrial wastes has gained attention in recent
years. When these feed stocks are pretreated for delignification, simple or oligomeric
phenolics and derivatives are generated from lignin. These compounds can inhibit the
hydrolysis catalyzed by cellulases. Thus, tannase can be utilized for degradation of

17
these oligomeric phenolics and, by doing so, alleviates the inhibition on cellulolysis
(Tejirian, et al., 2011). Tannase gene and tannase activity can be utilized for the
identification of Staphylococcus lugdunensis in humans and as an indicator of colon
cancer (Noguchi, et al., 2007). Tannase has been utilized for the production of
molecules with therapeutic applications, such as some esters derived from prunioside
A with anti-inflammatory activity. Other potential applications of tannase are found in
the manufacture of laundry detergents as an additive, in cosmetology to eliminate the
turbidity of plant extracts, and in the leather industry to homogenize tannin
preparation for high-grade leather tannins (Rodrıguez-Duran, et al., 2011).

18
1.3 Distribution of tannase in nature
The first reported a tannase gene from bacteria Staphylococcus lugdunensis was done
by Noguchi, et al., (2010). They cloned and sequenced a novel gene (tanA) from that,
which encodes a polypeptide of 613 amino acids with tannase activity. The tanA gene
was found to be specific for S. lugdunensis and had no significant similarity with the
genes coding for fungal tannases (Noguchi, et al., 2010). Later, Iwamoto and
coworkers cloned and sequenced the tannase gene from Lactobacillus plantarum
(tanLpl). The tanLpl gene was almost identical to a nucleotide sequence of L.
plantarumWCFS1 encoding a hypothetical protein but with a single base substitution
at four positions and was similar (46.7%) to tanA from S. lugdunensis (Iwamoto, et
al., 2008). More recently, Sharma and John reported the characterization of the
tannase gene from Enterobacter sp. (Sharma, et al., 2011). Multiple sequence
alignment showed that Enterobacter sp. tannase is not very much similar to tannase of
S. lugdunensis or L. plantarum, since only 10% and 13% amino acid residues of
Enterobacter sp. tannase are similar to those of S. lugdunensis and L. plantarum
tannases, respectively. Additionally, bacterial tannase are not closely related to fungal
tannases, either.
Tannase is widely distributed in diverse families of microorganisms and the
predominant members are the family of Bacillus sp. (Mondol, et al., 2001), Klebsiella
sp. (Banerjee, et al., 2012), Pseudomonas sp. (Selwal, et al., 2010), Enterobacter sp.
(Sharma, et al., 2011), Pantoea sp. (Pepi, et al., 2009), Lactobacillus sp. (Osawa, et
al., 2000).
More genera still need to be investigated for their availability of tannase through
phylogenetic tree (Figure 1.7 and Figure 1.8). Tannase distribution can be observed
from genus to genus and species to species to understand the distribution pattern
better for the entire bacterial kingdom, Gram-positive and Gram-negative
phylogenetic tree was constructed with the bacterial families already identified before
labeled with red lines. The distribution of tannase pattern among different species has
been studied for some time now, and has been summarized in Table1.2 and Figure1.7
and Figure1.8.

19
Table 1.2: Distribution of tannase producing bacteria
Organism Tannase predicted/reported Reference
Bacillus
B. cereus reported Mondol, et al., 2001
B. subtilis Sequence submitted/ reported Jana, et al., 2013
B. licheniformis reported Mondol, et al., 2000
B. sphearicus reported Raghuwanshi, et al., 2011
B. polymyxa reported Deschamps, et al., 1983
B. pumilus reported Deschamps, et al., 1983
B. massiliensis reported Belur, et al., 2010
Azotobacters sp. reported Gauri, et al., 2012
Pantoea sp. reported Pepi, et al., 2010
Pseudomonas
P. aeruginosa Selwal, et al., 2010
P. stutzeri Sequence submitted/ reported
P. citronellolis reported Chowdhury, et al., 2004
P. mendocina Sequence submitted/ reported
P. syringae Sequence submitted/ reported
P. savastanoi Sequence submitted/ reported
P. plecoglossicida reported Chowdhury, et al., 2004
Corynebacterium sp. reported Deschamps, et al., 1983
Paenibacillus polymyxa reported Deschamps, et al., 1983
Klebsiella
K. planticola reported Deschamps, et al., 1983
K. pneumoniae Sequence submitted/ reported Deschamps, et al., 1983
Selemonas ruminanticum reported Skene and Brooker, 1995
Citrobacter freundii reported Kumar, et al., 1999
Microbacterium terregens reported Belur, et al., 2010
Serratia
S. ficaria reported Belur, et al., 2010
S. marcescens reported Belur, et al., 2010
Serratia sp. Sequence submitted/ reported Pepi, et al., 2010
Providencia rettgeri reported Belur, et al., 2010
Lactobacillus
L. apodemi reported Osawa, et al., 2006

20
Organism Tannase predicted/reported Reference
L. plantarum Sequence submitted/ reported Ren, et al., 2013
L. paraplantarum Sequence submitted/ reported Iwamoto, et al.,2008
L. pentosus Sequence submitted/ reported Nishitani, et al., 2004
L. animalis reported Sasaki, et al., 2005
L. murinus reported Sasaki, et al., 2005
L. brevis reported Mathews, et al., 2006
L. buchneri reported Mathews, et al., 2006
L. casei reported Mathews, et al., 2006
L. helveticus reported Mathews, et al., 2006
L. hilgardii reported Mathews, et al., 2006
Entrobacter
E. asburiae reported Mandal and Ghosh, 2013
E. cloacae Sequence submitted/ reported Beniwal et al., 2010
E. ludwigii reported Singh, et al., 2012
Leuconostac
L. fallax reported
L. mesenteroides reported
Pediococcus
P. acidilactici reported
P. pentosaceus reported
Rhodococcus sp. reported Nadaf and Ghosh, 2011
Streptococcus gallolyticus Sequence submitted/ reported Iwamoto, et al.,2008
Staphylococcus lugdunensis Sequence submitted/ reported Noguchi, et al.,2007
Gluconacetobacter sp. Sequence submitted/ reported
Oenococcus oeni Sequence submitted/ reported

21
1.3.1 Distribution of Tannase in Gram-positive bacterial kingdom
The Gram-positive bacteria are divided into two phyla, one is phylum Firmicutes
consisting of low G+C bacteria, and second phylum Actinobacteria consisting of high
G+C bacteria. So far several Gram-positive bacteria have been discovered with
tannase activity and reported in several journals (Table 1.2). Among the Firmicutes,
organisms from the genus Bacillus, Lactobacillus, Streptococcus, Staphylococcus,
Leuconostoc and Paenibacillus have been reported to have tannase activity. While
from the Actinobacteria, organisms from only two genera Microbacterium and
Corynebacterium have been reported to have tannase activity. The tannase producing
bacterial distribution pattern among different species of the Gram-positive bacterial
kingdom has been summarized in Figure1.7.
Figure 1.7: Gram-positive bacterial kingdom tannase distribution shown with Red
lines.

22
1.3.2 Distribution of Tannase in Gram-negative bacterial kingdom
The Gram-negative bacteria consists of the phyla Proteobacteria, Aquificae,
Chlamydiae, Bacteroidetes, Chlorobi, Cyanobacteria, Fibrobacteres,
Verrucomicrobia, Spirochetes, Planctomycetes, Acidobacteria, Thermotogae and
Chloroflexi. So far several Gram-negative bacteria have been discovered with tannase
activity and reportedin several journals (Table 1.2). However, most of the organisms
discovered were from phyla Proteobacteria, consisting species from the genera
Enterobacter, Klebsiella, Serratia, Citrobacter, Pantoea, Providencia, Lonepinella,
Pseudomonas, Azotobacter, Rhodococcus, and Gluconacetobacter. The tannase
producing bacterial distribution pattern among different species of the Gram-negative
bacterial kingdom has been summarized in Figure1.8.
Figure 1.8: Gram-negative bacterial kingdom tannase distribution shown with Red
lines.

23
1.4 Aims and Objective
Tannase, is relatively a new enzyme when it comes to bacteria and thus not much
research has been done on it so far, apart from a few notables such as Deschamps
(Deschamps, et al., 1983), Kumar (Kumar, et al., 1999), Osawa (Osawa, et al., 2000),
Mondol (Mondol, et al., 2000), Nishitani (Nishitani, et al., 2005) and Belur (Belur, et
al., 2010); who themselves isolated and identified the strains. Tannase has several
industrial applications thus one of the aims of this project was to isolate and identify
bacterial strains that are capable to producing tannase. Among these strains, the
highest producing tannase strain will be used further for enzyme extraction, assay and
purification.
The second aim of this project was to develop a bioinformatics process that can help
identify the following:
a. Tannase containing enzyme in bacteria. The main process here would be to
identify the active conserved domain necessary for the enzyme’s catalytic
activity. As so far various different types of bacteria and fungi have been
identified with tannase activity, thus the interest here is to see the similarity
among their catalytic domain and what differences if there, that can affect the
enzyme’s activity.
b. Distribution of tannase enzyme in bacterial kingdom. Here, the objective is to
see how bacterial evolution has affected the distribution of the gene necessary
for tannase activity. As tannase is an enzyme that is not constitutively express
neither the enzyme’s structure or protein sequence has been found to be
conserved among several genera, thus a phylogenic study is in fact necessary
to further understand the evolution and distribution of this protein among the
bacterial kingdom.
c. Filling the gaps in phylogenetic tree of the distribution pattern of the enzyme
among the already identified genera to see if any other species are capable of
producing tannase as well.

24
2.1 Introduction
This chapter discusses the methodology that has been used in this research. The first
part (2.2), describes the different types of stains and media used in this research, their
composition and preparation methods. The second part (2.3), describes about sample
collection and processing such as measurement of different soil parameters, serial
dilution and spread plate. The third part (2.4), discusses about the E.coli isolation
procedure from sewage water while the fourth part describes (2.5), biochemical test
procedures used in order to identify the natural bacterial strains that were isolated.
The fifth part (2.6), describes the process of secondary screening of tannase producing
bacteria and the final part (2.7) describes about the bioinformatics tool used in order
to predict the presence of the protein in the strains isolated.
This research study was conducted based on the methodology which has been first
summarized in a flowchart.The details of the methodology are explained elaborately
through out the chapter.

25
Figure 2.1: Flowchart summarizing the overall research methodology.
Sample processing and serial dilution
Spread plate of the dilutions and bacterial growth
Screening of the microorganism on their
capability to produce the enzyme
Take the selected strains for biochemical test
identification
Take the identified bacterial strains and look for
sequence homology through genome BLAST
with known/reported tannase protein sequence
Predict whether the catalytic protein domain is
present for the bacteria to have tannase
producing ability
Predict the tannase distribution pattern in the
bacterial kingdom
Collection of environmental samples

26
2.2 Media and stain preparations
2.2.1 Staining preparation: Gram’s stain
The Gram’s stain is used to differentiate between Gram-positive and Gram-negative
bacteria. The main principle of Gram’s staining was followed as per GRAM’s original
protocol (Gram, 1884) and with modified adjustments in (Cappuccino & Sherman,
2005). The main reagents required for the Gram’s staining are given in the Table 2.1.
Table 2.1 Reagents for Gram’s staining.
Reagents gm/100ml
Primary Stain: Crystal Violet Staining Reagent
Crystal violet (certified 90% dye content) 2.0
Ethanol, 95% (vol/vol) 20.0
Ammonium oxalate 0.8
Mordant: Gram's Iodine
Iodine 1.0
Potassium iodide 2.0
Decolorizing Agent:
Ethanol, 95% (vol/vol)
Counterstain: Safranin
Stock solution:
Safranin O 2.5
95% Ethanol 100 ml
Working Solution:
Stock Solution 10 ml
Distilled water 90 ml

27
2.2.2 Staining preparation: Endospore stain
The main purpose of the endospore staining is to observe endospore forming bacteria
such as Bacillus and Clostridium which is an intracellular spore formed for reaching a
high degree of resistance to deleterious agents (Schaeffer & Fulton, 1933 and
Mormak & Casida, 1985). The modified endospore staining protocol was followed of
(Cappuccino & Sherman, 2005). The main reagents required for the Endospore
staining are given in the Table 2.2.
Table 2.2 Reagents for Endospore staining.
Reagents gm/100ml
Primary Stain: Malachite green staining Reagent
Malachite green 0.5
Distilled water 100
Decolorizing Agent:
Tap water
Counterstain: Safranin
Stock solution:
Safranin O 2.5
95% Ethanol 100 ml
Working Solution:
Stock Solution 10 ml
Distilled water 90 ml

28
2.2.3 Nutrient agar
The nutrient agar that was used is Nutrient Agar bought from Oxoid™. The agar was
prepared by suspending 75 g in 1 liter of distilled waterthen boiling to dissolve the
contents completely as mentioned by the Oxoid™ manual. Then, the media is
sterilized by autoclaving at 121°C for 15 minutes. If required the pH may be adjusted
to 6.5 by the addition of 1% sodium bicarbonate solution. The formula of the Nutrient
agar media is given in Table 2.2.
Table 2.3: Ingredients for Nutrient Agar*.
Ingredients gm/liter
Yeast extract 4.0
Tryptone 5.0
Glucose 50.0
Potassium dihydrogen phosphate 0.55
Potassium chloride 0.425
Calcium chloride 0.125
Magnesium sulphate 0.125
Ferric chloride 0.0025
Manganese sulphate 0.0025
Bromocresol green 0.022
Agar 15.0
*(http://www.oxoid.com/UK/blue/prod_detail/prod_detail.asp?pr=CM0309&org=107
&c=UK&lang=EN)

29
2.2.4 M-FC Agar
M-FC Agar Base is used for the detection and the enumeration of faecal coliforms.
This medium is based on the property of faecal coliforms to grow at 44-45°C and the
ability to ferment lactose (Grabow, et al., 1981). Proteose peptone, tryptose and yeast
extract provide the necessary nutrients for the growth of faecal coliforms. Bile salts
inhibit the growth of contaminating gram-positive microorganisms. Aniline blue,
suppresses the growth of many Gram-positive microorganisms and along with rosolic
acid forms the indicator system of the medium. Although, the average countson M-FC
agar without rosolic acid are higherthan on standard M-FC agar, thus here the rosalic
acid was not used (Grabow, et al., 1981). After incubation at 44-45°C coliforms will
form blue colonies whereas non-coliforms will form gray colored colonies on M-FC
Agar Base.
The M-FC Agar Base that was used was bought from HIMEDIA™. The media is
prepared by suspending 52.1 grams in 1000 ml distilled water after which heating to
boiling with gentle swirling to dissolve completely as according to the HIMEDIA™
manual. This is then cooled to 45°C and pour into sterile Petri plates. The formula of
the M-FC Agar Base media is given in Table 2.4.
Table 2.4: Ingredients for M-FC Agar Base*.
Tryptose 10.0
Proteose peptone 5.0
Yeast extract 2.0
Lactose 12.5
Bile salts mixture 1.5
Sodium chloride 5.0
Aniline blue 0.1
Agar 15.0
*(http: //himedialabs.com/TD/M1122.pdf.)

30
2.2.5 MacConkey agar
MacConkey agar is used for the isolation of Gram-negative enteric bacteria and the
differentiation of lactose fermenting from lactose non-fermenting Gram-negative
bacteria. The selective action of this medium is attributed to crystal violet and bile
salts, which are inhibitory to most species of Gram-positive bacteria (Zimbro, et al.,
2009). Gram-negative bacteria usually grow well on the medium and are
differentiated by their ability to ferment lactose. Lactose fermenting strains grow as
red or pink colonies and may be surrounded by a zone of acid precipitated bile. The
red color is due to the production of acid from lactose, absorption of neutral red and a
subsequent color change of the dye when the pH of medium falls below 6.8 (Zimbro,
et al., 2009).
The MacConkey Agar that was used was bought from HIMEDIA™. The agar is
boiling with gentle swirling to dissolve the agar completely as mentioned in the
HIMEDIA™ manual. Then, the media was sterilized by autoclaving at 121°C for 15
minutes. The formula of the MacConkey agar media is given in Table 2.5.
Table 2.5: Ingredients for MacConkey Agar*.
Pancreatic digest of gelatin 17.0
Casein enzymic hydrolysate 1.50
Pancreatic digest of gelatin 17.0
Peptic digest of animal tissue 1.50
Lactose 10.0
Bile salts 1.50
Sodium chloride 5.0
Neutral red 0.03
Crystal violet 0.001
Agar 15.0
*(http://himedialabs.com/TD/M081.pdf)

31
2.2.6 Mannitol Salt agar
Mannitol Salt agar is a selective medium prepared for the isolation of presumptive
pathogenic staphylococci. Most other bacteria are inhibited by the high salt
concentration with the exception of some halophilic marine organisms. Presumptive
coagulase-positive staphylococci produce colonies surrounded by bright yellow zones
whilst non- pathogenic staphylococci produce colonies with reddish purple zones
(Zimbro, et al., 2009). The Mannitol Salt agar that was used was bought from
Oxoid™. The agar was prepared by suspending 111g in 1 liter of distilled water then
boiling to dissolve completely as mentioned by the Oxoid™ manual. Afterwards
sterilization by autoclaving is done at 121°C for 15 minutes. The formula of the
Mannitol Salt agar media is given in Table 2.6.
Table 2.6: Ingredients for Mannitol Salt Agar*.
`Lab-Lemco’ powder 1.0
Peptone 10.0
Mannitol 10.0
Sodium chloride 75.0
Phenol red 0.025
Agar 15.0
*(http://www.oxoid.com/uk/blue/prod_detail/prod_detail.asp?pr=CM0085&org=153
&c=uk&lang=en)

32
2.2.7 Eosin Methylene Blue agar
Eosin Methylene Blue Agar is used for the isolation and differentiation of Gram-
negative enteric bacteria.
In this media methylene blue and eosin-Y inhibit Gram-positive bacteria to a limited
degree. These dyes serve as differential indicators in response to the fermentation of
carbohydrates. The ratio of eosin and methylene blue is approximately to 6:1. Lactose
and sucrose are the sources of energy by being fermentable carbohydrates. The
coliforms that produce purplish black colonies due to taking up of methylene blue-
eosin dye complex, when the pH drops which is absorbed into the colony. Non-
fermenters raise the pH of surrounding medium by oxidative deamination of protein,
which solubilize the methylene blue-eosin complex resulting in colorless colonies
(Zimbro, et al., 2009).
The Eosin Methylene Blue Agar that was used was bought from HIMEDIA™.The
agar is prepared by suspending 35.96 grams in 1000 ml distilled water after which
heating to boiling with gentle swirling to dissolve the agar completely as mentioned
by the HIMEDIA™ manual. Then, the media is sterilized by autoclaving at 121°C for
15 minutes.The formula of the Eosin Methylene Blue Agar media is given in Table
2.7.
Table 2.7: Ingredients for Eosin Methylene Blue Agar*.
Dipotassium phosphate 2.0
Lactose 5.0
Sucrose 5.0
Eosin - Y 0.40
Methylene blue 0.065
Agar 12.50
* (http://www.himedialabs.com/TD/M317.pdf)

33
2.2.8 Starch agar
Starch agar is a differential medium that tests the ability of an organism to produce
the extracellular enzymes (exoenzymes) α-amylase and oligo-1, 6-glucosidase that are
secreted out of the bacteria and diffuse into the starch agar. These enzymes hydrolyze
starch by breaking the glycosidic linkages between glucose subunits and allow the
products of starch hydrolysis to enter the cell (Hemraj, et al., 2013).
When bacteria capable of producing α-amylase and oligo-1, 6-glucosidase are grown
on starch agar, they secrete these enzymes into the surrounding areas and hydrolyze
the starch (Hemraj, et al., 2013).To detect the hydrolysis of starch, Gram’s iodine is
used. Gram’s iodine reacts with starch to form a dark blue, purple, or black complex
depending upon the concentration of iodine.
The agar is prepared by suspending the required amount of the ingredients in 1 L of
distilled water and mixed thoroughly. Heat is applied to dissolve with frequent
agitating the contents and then the media is autoclaved at 121°C for 15 minutes. The
melted medium is then poured into petri plates and the agar is allowed solidify
(Zimbro, et al., 2009).The required amount of the ingredients is given in Table 2.8.
Table 2.8: Ingredients for Starch Hydrolysis Agar.
Beef extract 2.0
Soluble Starch (Merck) 10.0
Agar (Bacto) 12.0

34
2.2.9 Skim Milk agar
The enzyme caseinase is secreted out of the cells (an exoenzyme) into the surrounding
media, catalyzing the breakdown of milk protein, called casein, into small peptides
and individual amino acids which are then taken up by the organism for energy use or
as building material.(Hemraj, et al., 2013)
1 g of agar is suspended in 50 ml distilled water with 5 g skim milk powder
suspended in 50 ml distilled water to make 100 ml skim milk agar. Both of the two
medium are autoclaved at 121°C for 15 minutes, mixed and then poured into plates
(Zimbro, et al., 2009). The required amount of the ingredients to make skim milk agar
are given in Table 2.9.
Table 2.9: Ingredients for Skim Milk Agar.
Skim milk powder (Titon biotech) 50.0
Agar (Bacto) 10.0

35
2.2.10 Gelatin hydrolysis Medium
Gelatin is a protein derived from the connective tissues of vertebrates, that is, collagen
which is produced when collagen is boiled in water. Gelatin hydrolysis detects the
presence of gelatinase (Hemraj, et al., 2013). Gelatinase are proteases secreted
extracellularly by some bacteria which hydrolyze or digest gelatinwhich is detected
using a nutrient gelatin medium. This medium contains peptic digest of animal tissue
(peptone), beef extract, and gelatin. Gelatin serves as both solidifying agent and
substrate for gelatinase activity. When nutrient gelatin tubes are stab-inoculated with a
gelatinase-positive bacterium, the secreted gelatinases will hydrolyze the gelatin
resulting in the liquefaction of the medium (Clarke, et al., 1952).
Since gelatin is digested and is no longer able to gel, the medium will remain liquid
when placed inside a refrigerator or in an ice bath. A nutrient gelatin medium
inoculated with a gelatinase-negative bacterium will remain solid after the cold
treatment (Hemraj, et al., 2013).
The agar is prepared by suspending the required amount of the ingredients in 1 L of
distilled water and mixed thoroughly. Heat is applied to dissolve with frequent
agitating the contents and then the media is autoclaved at 121°C for 15 minutes. The
melted medium is then poured into petri plates and the agar is allowed solidify
(Zimbro, et al., 2009). The required amount of the ingredients to make gelatin
hydrolysis agar are given in Table 2.10.
Table 2.10: Ingredients for Gelatin hydrolysis Medium.
Peptone (Oxoid) 5.0
Beef extract (Oxoid) 2.0
Gelatin (power food grade) 120.0

36
2.2.11 Simmons Citrate Agar
The citrate test is commonly a part of a group of tests, the IMViC tests that screens
bacterial isolates for the ability to utilize citrate as its carbon and energy source.
Citrate is the sole source of carbon in the Simmons citrate medium while inorganic
ammonium salt (NH4H2PO4) is the sole fixed nitrogen source (Zimbro, et al., 2009).
Upon uptake by the cell, citrate is cleaved by citrate lyase to give oxaloacetate and
acetate. The oxaloacetate is further metabolized to pyruvate and CO2. The carbon
dioxide that is released will subsequently react with water and the sodium ion in the
medium to produce sodium carbonate, an alkaline compound that will raise the pH.
In addition, ammonium hydroxide is produced when the ammonium salts in the
medium are used as the sole nitrogen source. The bromothymol blue pH indicator is a
deep forest green at neutral pH. With an increase in medium pH to above 7.6,
bromothymol blue changes to blue (Zimbro, et al., 2009).
The Simmons Citrate Agar that was used was bought from BBL™. The agar is
boiling with gentle swirling to dissolve the agar completely as according to the
BBL™ manual.Then, the media is sterilized by autoclaving at 121°C for 15
minutes.The formula of the Simmons Citrate Agar mediais given in Table 2.11.
Table 2.11: Ingredients for Simmons Citrate Agar*.
Ammonium dihydrogen phosphate 1.0
Dipotassium phosphate 1.0
Sodium chloride 5.0
Sodium citrate 2.0
Magnesium sulphate 0.2
Bromothymol blue 0.08
Agar 15.0
*(http://www.bd.com/ds/technicalCenter/inserts/L007504(07)(201101)pdf)

37
2.2.12 Motility Indole Urease Agar
MIU medium base is used for detection of motility, urease and indole production. In
the media casein enzymic hydrolysate provide amino acids and other nitrogenous
substances. Dextrose is the fermentable carbohydrate while phenol red is the pH
indicator which turns pink- red in alkaline conditions. Motility and urease reactions
are read before testing for Indole production. Motile organisms show either diffused
growth or turbidity extending away from stab inoculation line while non-motile
organisms grow along the stabline. Organisms that utilize urea produce ammonia
which makes the medium alkaline, showing pink-red color by change in the phenol
red indicator. Indole is produced from tryptophan present in casein enzymic
hydrolysate. The indole produced combines with the aldehyde present in the Kovac's
reagent to form a red complex (Hemraj, et al., 2013).
The Motility Indole Urease Agar mediathat was used bought from HIMEDIA™. The
agar is prepared by suspending 18 grams in 950 ml distilled water after which heated
to boiling with gentle swirling to dissolve the agar completely.Then 95 ml amounts
are dispensed into flasks and sterilize by autoclaving at 121°C for 15 minutes. The
media is then cooled to about 50-55°C and aseptically5 ml sterile 40% Urea (Merck)
solution is added per 95 ml basal medium which are then dispensed into sterile test
tubes as according to the HIMEDIA™ manual.The formula of the Motility Indole
Urease Agar mediais given in Table 2.12.
Table 2.12: Ingredients for Motility Indole Urease Agar*.
Dextrose 1.0
Sodium chloride 5.0
Phenol red 0.010
Agar 2.0
* (http://himedialabs.com/TD/M1076.pdf)

38
2.2.13 Triple Sugar Iron Agar
Triple Sugar Iron Agar is used for the identification of Gram-negative enteric bacilli
on the basis of dextrose, lactose and sucrose fermentation and hydrogen sulphide
production (Zimbro, et al., 2009).
In the media lactose, sucrose and dextrose are the fermentable carbohydrates. Sodium
thiosulphate and ferrous ions make H2S indicator system. Phenol red is the pH
indicator. Organisms that ferment glucose produce a variety of acids, turning the color
of the medium from red to yellow. More amounts of acids are liberated in butt
(fermentation) than in the slant (respiration) (Zimbro, et al., 2009 and Hemraj, et al.,
2013).
Growing bacteria also form alkaline products from the oxidative decarboxylation of
peptone and these alkaline products neutralize the large amounts of acid present in the
butt. Thus the appearance of an alkaline (red) slant and an acid (yellow) butt after
incubation indicates that the organism is a glucose fermenter but is unable to ferment
lactose and/or sucrose. Bacteria that ferment lactose or sucrose (or both), in addition
to glucose, produce large amounts of acid enables no reversion of pH in that region
and thus bacteria exhibit an acid slant and acid butt (Zimbro, et al., 2009 and Hemraj,
et al., 2013).
Gas production CO2is detected by the presence of cracks or bubbles in the medium,
when the accumulated gas escapes. Thiosulphate is reduced to hydrogen sulphide by
several species of bacteria and H2S combines with ferric ions offerric salts to produce
the insoluble black precipitate of ferrous sulphide (Zimbro, et al., 2009 and Hemraj, et
al., 2013).
The Triple Sugar Iron Agar that was used was bought from HIMEDIA™. The agar is
boiling with gentle swirling to dissolve the agar completely as according to the
HIMEDIA™ manual. This is then distributed into test tubes which are sterilized by
autoclaving at 121°C for 15 minutes.The formula of the Triple Sugar Iron Agar media
is given in Table 2.12.

39
Table 2.13: Ingredients for Triple Sugar Iron Agar*.
Yeast extract 2.0
Beef extract 2.0
Lactose 10.0
Sucrose 10.0
Dextrose 1.0
Sodium chloride 5.0
Ferrous sulphate 0.20
Sodium thiosulphate 0.30
Phenol red 0.024
Agar 12.0
* (http://himedialabs.com/TD/M021.pdf)

40
2.2.14 MRVP medium
This test is used to detect the ability of an organism to produce and maintain stable
acid (Methyl red) and acetoin (Voges-Proskauer) in a bacterial broth culture as an end
product from glucose fermentation. This test is also a part of a group of tests, the
IMViC tests (Zimbro, et al., 2009).
For Methyl Red test: organisms ferment sugars present in the broth by the mixed acid
pathway that gives 4 mol of acidic products (mainly lactic and acetic acid), 1 mol of
neutral fermentation product (ethanol), 1 mol of CO2, and 1 mol of H2 per mol of
glucose fermented. The large quantity of acids produced causes a significant decrease
in the pH of the culture medium. When the culture medium turns red after addition of
methyl red, because of a pH at or below 4.4 from the fermentation of glucose, the
culture has a positive result for the Methyl Red test. A negative Methyl Red test is
indicated by a yellow color in the culture medium, which occurs when less acid is
produced (pH is higher) from the fermentation of glucose (Zimbro, et al., 2009).
For Voges-Proskauer test: Bacteria that ferments sugars via the butanediol pathway
produce acetoin (i.e., acetyl methyl carbinol or 3-hydroxybutanone) as an
intermediate which can be further reduced to 2,3-butanediol. In the presence of KOH
the intermediate acetoin is oxidized to diacetyl, a reaction which is catalyzed by α-
naphthol. Diacetyl reacts with the guanidine group associated with molecules
contributed by peptone in the medium, to form a pinkish-red-colored product. The α-
naphthol in the Barritt’s modification of the Voges-Proskauer test serves as a color
intensifier (Zimbro, et al., 2009).
The MRVP Medium that was used was bought from Oxoid™.The broth is prepared
by suspending 17 grams in 1000 ml distilled water after which heating to boiling with
gentle swirling to dissolve completely as according to the Oxoid™ manual. This is
then distributed into test tubes which are sterilized by autoclaving at 121°C for 15
minutes. The formula of the Methyl Red and Voges-Proskauer media is given in
Table 2.14.

41
Table 2.14: Ingredients for MRVP Medium*.
Peptone 7.0
Glucose 5.0
Phosphate buffer 5.0
Methyl red test
Methyl red 0.1
Ethanol 0.30
Deionized water 0.20
Voges-Proskauer test
Barritt’s reagent A
α-naphthol 0.05
Ethanol 0.1
Barritt’s reagent B
KOH 0.4
Deionized water 0.1
*(http://www.oxoid.com/uk/blue/prod_detail/prod_detail.asp?pr=CM0043&org=71&
c=uk&lang=en)

42
2.2.15 Tannic acid supplemented media
Tannic acid containing media is made to isolate and screen bacteria that are capable of
degrading tannic acid as a sole carbon source by producing the enzyme tannase to
yield glucose and gallic acids.
For the preparation of the tannic acid supplemented media first the mineral media is
made following the composition of Mondol et al. (2001). The mineral media was
autoclaved at 121˚C for 20 minutesafter which it was allowed to slightly cool and
Cycloheximide of 0.01 g was added to it using a sterile autoclaved pipette (Walsh, et
al., 2013). The media is mixed thoroughly while avoiding the formation of bubbles.
Cycloheximide inhibits the fungal growth that might be present in the soil sample.
For Tannic acid supplementation the protocol followed by Pepi et al. (2010) was used
where Tannic acid powder (LOBA chemie) was made into solution of 2% (2 g in 100
ml) by dissolving in autoclaved distilled water. pH is then adjusted to 5.5 by adding
10% NaOH drop-wise which results in the change of the color of the solution from
light straw yellow to darker straw yellow.When the media has been set they are
flooded with freshly prepared 2% tannic acid solution for 1 minute according to
modified Pepi et al. (2010). Following this the tannic acid solution is discarded and
the media plates are allowed to dry. The composition of the Tannic acid supplemented
media is given in Table 2.15.
Table 2.15: Ingredients for Tannic acid supplemented minimal media.
Dipotassium phosphate(Merck) 0.5
Monopotassium phosphate(Merck) 0.5
Magnesium sulphate(Merck) 0.5
Ammonium nitrate(Merck) 2.0
Bacto agar (BBL) 25
Tannic acid (LOBA chemie) 20
pH 5.5± 0.2 at 25°C

43
2.3 Sample collection for tannase producing bacteria
The procedure for collection of fruit dump sample to isolate tannase producing
bacteria and the downstream process are summarized in the Flowchart 2.2.
Figure 2.2: Overview of the isolation and identification of Tannase producing
bacteria from soil.
Collect fruit dump waste soil
Measure soil pH Measure soil moisture
content
Sample processing and serial
dilution from 10-1
to 10-5
Spread plate of the dilutions in
Nutrient agar (NA) plate
Spread plate of the dilutions in
Tannic acid supplemented
minimal media (TA) plate
Take well isolated colonies and plate in
Nutrient agar (NA) to obtain pure colonies
Take the selected strains for
morphologicaland biochemical test
identification
Identify the strains based on biochemical
test
Secondary screening of the micro-organism
on their capability to produce tannase by
adding FeCl3 solution onto the agar
Primary screening of tannase producing
bacteria by observing growth in TA plate

44
2.3.1 Environmental sample collection
Soil samples, deep brown or black in color, were collected in sterile zip lock bags
using a sterile spatula from a fruit waste dump located at Baridhara Bashundhara,
Dhaka Bangladesh. The fruit dump is a good source for tannin-rich plant deposit
(Chowdhury, et al., 2004). Mostly the soils with it are rich in tannase containing
microorganisms required to bio-degrade the plant tannin (Chowdhury, et al., 2004).
The soil sample was subjected to characterization by following parameters.
i) pH: The pH of the soil was determined by dissolving 1 g of the sample in 10 ml of
sterile distilled water (Chowdhury, et al,. 2004) and then measuring the solution with
a pH meter.
ii) Moisture: The moisture content was determined by first weighing the clean dry
beaker, after which a representative quantity of soil is placed in it. Aluminium foil
was placed over the top of the beaker containing the soil sample, which was then
placed inside an oven (O’Kelly, et al, 2014). After drying overnight the moisture
content was measured with the following formula:
MC % = W2 - W3
W3- W1
Where:
W1 = Weight of container (g)
W2 = Weight of moist soil + container (g)
W3 = Weight of dried soil + container (g)
X100

45
2.3.2 Environmental sample serial dilution
The soil sample was taken and aseptically 2 g of the soil was dissolved in 10 ml of
autoclaved 0.85% phosphate buffered saline (PBS) solution (Agustini, et al., 2012).
This solution or stock solution was first vortexed and then let to sit so that the solid
contents of the soil would sediment while the liquid supernatant can be used. 1 ml of
sample stock was taken and added to 9 ml of autoclaved 0.85% PBS. This was then;
vortexed at the maximum speed (Agustini, et al., 2012).This tube was labeled as 10-1
.
From this test tube again 1 ml was taken to another 9 ml of 0.85% PBS and label this
to 10-2
. This process was repeated a few more times to get 10-3
, 10-4
and 10-5
dilution
(Agustini, et al., 2012).
2.3.3 Spread plate
Inoculums from each dilution (100 µl) from 10-1
to 10-5
were taken and spread onto
Nutrient agar plate NA (control) and 2% Tannic acid supplemented agar plate TA
following the general spread plate technique (Cappuccino, & Sherman, 2005).
2.3.4 Growth of colonies on Nutrient Agar (NA)
The Nutrient Agar plateswere incubated at 37˚C for 1 day while checking the growth
of the colonies and then counting the number of colonies per plate dilutions using a
colony counter.
2.3.5 Growth of colonies on Tannic acid supplemented minimal
media (TA)
The Tannic acid supplemented agar plates were incubated at 37˚C for 2 days while
checking the growth of the colonies each day and then counting the number of
colonies per plate dilutions using a colony counter.
2.3.6 Selection of colonies
Appropriate and well isolated colonies were picked from the tannic acid
supplemented minimal media spread plates of 10-3
and 10-4
dilutions. Sixteen different
isolates were selected and were then sub-cultured several times in Nutrient Agar to
obtain pure colonies (Cappuccino, & Sherman, 2005). These sixteen colonies were
numbered as follows: T1, T2, T3, T4, T5, T6, T7, T8, T9, T10, T11, T12, T13, T14,
T15 and T16. Among these sixteen isolates, based on their colony morphology,

46
similar looking colonies were omitted, and fourteen different and dissimilar colonies
were chosen for the further biochemical test. These colonies number were: T1, T2,
T3, T4, T5, T7, T8, T9, T10, T11, T13, T14, T15 and T16.

47
2.4 E.coli isolation and identification
The procedure for collection of sewage waste water sample for isolation of E.coli that
produces tannase and the downstream process are summarized in the Flowchart 2.3
below.
Figure 2.3: Overview of isolation and identification of E.coli from sewage water.
Collection of sewage water
Sample processing or serial dilution
Spread plate the dilutions up to 106
on
m-FC agar; incubate at 44.5°C for 24
hours
Take the well isolated blue colonies
from the M-FC agar; streak them again
onto M-FC agar; incubate at 44.5°C for
24 hours
Take the well isolated blue colonies
from the M-FC agar; streak them onto
MacConkey agar then followed by EMB
agar; incubate at 37°C for 24 hours
Select greenish colonies and perform
morphological & biochemical test to
identify bacteria
Take the identified E.coli strain and find
whether it produces tannase

48
2.4.1 Environmental sample collection
Sewage water sample murky and dark in color, were collected in sterile plastic
bottlesusing a sterile spatula from a sewage waste site located at Baridhara,
Bashundhara, Dhaka Bangladesh. The sewage water is a good source for E.coli as it is
a part of the faecal coliform (Farasat, et al., 2012).
2.4.2 Sewage waste water sample collection
The sewage waste water sample was taken and aseptically 2 g of the water was
dissolved in 10 ml of autoclaved 0.85% phosphate buffered saline (PBS) solution
(Agustini, et al., 2012). This solution or stock solution was first vortexed and then let
to sit so that any solid contents present would sediment while the liquid supernatant
can be used. 1 ml of sample stock was taken and added to 9 ml of autoclaved 0.85%
PBS. This was then, vortexed at the maximum speed (Agustini, et al., 2012 and
Farasat, et al., 2012). This tube was labeled as 10-1
. From this test tube again 1 ml was
taken to another 9 ml of 0.85% PBS and label this to 10-2
. This process is repeated a
few more times to get 10-3
, 10-4
and 10-5
dilution (Agustini, et al., 2012 and Farasat, et
al., 2012).
2.4.3 Spread plate
From each of these test tubes of different dilution 1 ml of the sample was taken and
spread plated followed by streak plating them on M-FC agar. The plates were
incubated at 44.5˚C for 24 hours (Grabow, et al., 1981).
2.4.4 Streak plate and growth of colonies
Appropriate and well isolated blue colonies were picked from the 10-3
and 10-4
dilutions spread plates and were sub-cultured several times on M-FC agar followed by
Eosin Methylene Blue Agar and MacConkey Agar to obtain pure colonies until E.coli
characteristic colonies were observed whichwerethen used for the biochemical test
identification (Cappuccino, & Sherman, 2005 and Zinnah, et al., 2007).

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