Characterization of ureolytic bacteria isolated from … culture technique was used in this study to...
Transcript of Characterization of ureolytic bacteria isolated from … culture technique was used in this study to...
CHARACTERIZATION OF UREOLYTIC
BACTERIA ISOLATED FROM LIMESTONE
CAVES OF SARAWAK AND EVALUATION
OF THEIR EFFICIENCY IN
BIOCEMENTATION
By
ARMSTRONG IGHODALO OMOREGIE
A thesis presented in fulfilment of the requirements for the
degree of Master of Science (Research)
Faculty of Engineering, Computing and Science
SWINBURNE UNIVERSITY OF TECHNOLOGY
2016
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ABSTRACT
The aim of this study was to isolate, identify and characterise bacteria that are capable
of producing urease enzyme, from limestone cave samples of Sarawak. Little is known
about the diversity of bacteria inhabiting Sarawak’s limestone caves with the ability of
hydrolyzing urea substrate through urease for microbially induced calcite precipitation
(MICP) applications. Several studies have reported that the majority of ureolytic
bacterial species involved in calcite precipitation are pathogenic. However, only a few
non-pathogenic urease-producing bacteria have high urease activities, essential in MICP
treatment for improvement of soil’s shear strength and stiffness.
Enrichment culture technique was used in this study to target highly active urease-
producing bacteria from limestone cave samples of Sarawak collected from Fairy and
Wind Caves Nature Reserves. These isolates were subsequently subjected to an
increased urea concentration for survival ability in conditions containing high urea
substrates. Urea agar base media was used to screen for positive urease producers
among the bacterial isolates. All the ureolytic bacteria were identified with the use of
phenotypic and molecular characterizations. For determination of their respective urease
activities, conductivity method was used and the highly active ureolytic bacteria
isolated comparable with control strain used in this study were selected and used for the
next subsequent experiments in this study. Effects of cultural conditions on urease
activity and evaluation of biocementation potential of these locally selected ureolytic
isolates were also performed.
Out of the ninety bacteria subcultured from enriched cultures containing the cave
samples, thirty-one bacterial isolates were selected based on their respective abilities of
producing urease enzyme by completely turning the colour of urea agar base medium
from yellow to pink in comparison to other isolated urease producing bacteria and the
control strain (Sporosarcina pasteurii, DSM33) used in this study. The microscopic
analysis using Gram staining technique showed that majority of the bacterial isolates
were Gram-positive bacteria while only three of the isolates were Gram-negative
bacteria. In addition, majority of the bacterial cells were rod-shaped except for one
bacterial isolate which was a coccus. Endospore staining test results indicate also
indicated that all except one isolate were spore forming bacteria.
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The BLAST results from molecular characterization of the ureolytic isolates suggested
that they were closely related to bacteria from the Sporosarcina pasteurii group,
Pseudogracilibacillus auburnensis group, Staphylococcus aureus group, Bacillus lentus
group, Sporosarcina luteola group and Bacillus fortis group when compared to the 16S
rRNA sequencing data in NCBI nucleotide BLAST database.
Specific urease activity determination from the calculation of conductivity and urease
activity showed that out of all the bacterial cultures, bacterial isolates designated as
NB33, LPB21, NB28, NB30 and the control strain had 19.975, 23.968, 19.275, 20.091
and 17.751 mM urea hydrolysed.min-1.OD-1 respectively, suggesting they had the
highest specific urease activities when compared to the rest isolates. The effect of
cultural conditions on urease activities involving the aforementioned local isolates and
control strain showed that incubated these conditions: at 25 to 30oC; pH 6.5 to 8.0;
incubation period at 24 hr; and urea concentration of 6 to 8%, maximum specific urease
activities for the selected ureolytic bacteria isolates and control strain were obtained.
The biocement treatment test using isolates NB33, LPB21, NB28, NB30 and the control
strain on poorly graded soil clearly showed that MICP is microbially induced and not
chemically induced. The results presented in this study showed that out of all the sand
columns treated, all except the columns containing negative control (only cementation
solution) had calcium carbonate precipitation shown on the top surfaces of their
respective columns. Each column treated with microbial cultures and cementation
solution (containing 1 M or urea and CaCl2) were able to bind the sand particles
together. However, it was observed that there was higher cementation level at positions
close to the injection points which resulting in more calcite contents to be obtained at
this layers of the biocemented sands. Based on the surface strength using penetrometer
test and compressive strength using UCS test, samples treated with isolates LPB21 and
NB28 showed significant strengths when compared to other isolates, consortia, and the
control strain. However, the rest isolates showed similar performance with the control
strain. The application of these newly isolates highly active ureolytic bacteria can be
used to for other MICP treatments in civil and geotechnical industries. The findings in
this study suggest that the isolated ureolytic bacteria (NB28, LPB21, NB33, and NB30)
have the potential to be used as alternative microbial MICP agents for biocement
applications.
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ACKNOWLEDGEMENT
Foremost, I would like to express my deepest gratitude to my principal coordinating
supervisor: Assoc. Prof Dr Peter Morin Nissom (Associate Dean, Science) for all the
valuable discussion, brainstorm, helpful advice, critics, challenges and encouragements
throughout this research study. His overwhelming supervision made me develop new
insights and ideas during this research. His quest for “high-quality work”, made me stay
active, focused and enthusiastic. He also provided critical reviews of my experiments
and writing, prompting me to improve problem solving and writing skills. I would also
like to thank my associate supervisor: Dr Irine Runnie Ginjom for her insightful
discussion and comments on my experimental progress. Her invaluable advice, co-
supervision, and encouragement throughout this study helped made this thesis a
success.
I would like to gratefully acknowledge Assoc. Prof Dr Dominic Ek Leong Ong
(Director, Swinburne Sarawak Research Centre for Sustainable Technologies) and Dr
Ngu Lock Hei (Course coordinator, Chemical Engineering Department) for their
financial support (SSRG) used to partially fund my research project. I am thankful for
the continuous moral support and helpful discussion from Assoc. Prof Dr Dominic Ek
Leong, especially with the idea of going to the caves to screen for calcite-precipitating
microorganisms.
I extend my appreciation to Sarawak Biodiversity Centre (SBC) and Sarawak Forestry
Department (SFD) for issuing the permits (SBC-RA-0102-DO and NCCD.907.4.4
[JLD.11]-37) which enabled me to collect samples from Fairy Cave (N 01°22’53.39” E
110°07’02.70”) and Wind Cave (N 01°24’54.20” E 110°08’06.94”) Nature Reserves,
located in Bau, Kuching Division, Sarawak, Malaysia. The collection of the samples
from these extreme environments to conduct biological research stipulated the
potentials of screening, identifying and characterising highly active isolated ureolytic
bacteria. I am thankful to Dr Paul Mathew Neilsen, Associate director of graduate
studies and research education. His thoughtful guidance and warm encouragement,
especially during my confirmation of candidature helped make me achieve my research
goals. I am sincerely grateful for his continual willingness of finding time out of his
busy schedule to meet me and discuss on how I could tackle research challenges and
improve my research study.
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I would also like to acknowledge Assist. Prof Salwa Al-Thawadi, Dr Ralf Cord-
Ruwisch, PD Dr David Schleheck and Assist. Prof Leon van Paassen for providing
indispensable guidance on how to measure urease activity, the appropriate way of
determining specific urease activity and selective investigation of cultural conditions on
urease activities. I am very thankful for taking your time to reply my inquiries via
emails and researchgate.net.
I am thankful to the science laboratory officers and technicians: Chua JiaNi, Nurul
Arina Salleh, Cinderella Sio and Marclana Jane Richard, for providing me with
experimental materials and allowing me to make use of some apparatus during the
course of my research study. Without their enormous assistance, my research would not
have been completed on time. An exceptional gratitude goes to Hasina Mohammed
Mkwata for being a helpful research lab mate and an amazing girlfriend. Her assistance
while I carried out my experiment, specifically during the measurement of conductivity,
biomass concentration and effect of cultural conditions on urease activity made my
experiments very convenient. I also extend my appreciation to Ghazaleh
Khoshdelnezamiha for playing a significant role during the in vitro biocement test. Her
efforts and a keen interest in my research made my experiment successful. An extensive
appreciation goes to Dr Noreha Mahidi and Holed Juboi for their vehement assistance
during molecular characterization of the isolated ureolytic bacteria. It was a pleasure
working with her. Big thanks also go to my fellow lab colleagues: Nurnajwani Senian
and Ye Li Phua, for providing assistance during sample collection and when I
conducted my experiments in the laboratory.
I would like to thank my amazing parents: Mr Cletus and Mrs Margaret Omoregie, for
their amazing love, care, patience and their financial supports used to partly fund my
research. Their sacrifices in sponsoring my postgraduate study are forever appreciated. I
also warmly appreciate my siblings: Jennifer, Sharon, and Thelma, for their tender
affection and supports during the years I conducted my experiments and wrote on my
thesis. I am obsequiously grateful to God Almighty for all the blessings and abundances
bestowed on me and for making my MSc research a success.
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DECLARATION
I hereby declare that this research entitled “Characterization of ureolytic bacteria
isolated from limestone caves of Sarawak and evaluation of their efficiency in
biocementation” is original and contains no material which has been accepted for the
award to the candidate of any other degree or diploma, except where due reference is
made in the text of the examinable outcome; to the best of my knowledge contains no
material previously published or written by another person except where due reference
is made in the text of the examinable outcome; and where work is based on joint
research or publications, discloses the relative contributions of the respective workers or
authors.
(ARMSTRONG IGHODALO OMOREGIE)
DATE: 06 June 2016
In my capacity as the Principal Coordinating Supervisor of the candidate’s thesis,
I hereby certify that the above statements are true to the best of my knowledge.
(ASSOCIATE PROFESSOR DR. PETER MORIN NISSOM)
DATE: 06 June 2016
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SCIENTIFIC OUTPUT
PUBLICATIONS
Omoregie, AI, Senian, N, Ye Li, P, Hei, NL, Leong, DOE, Ginjom, IRH & Nissom, PM, 2016, 'Screening for Urease-Producing Bacteria from Limestone Caves of Sarawak', Borneo Journal of Resource Science and Technology, 6 (1): 37-45. Omoregie, AI, Senian, N, Ye Li, P, Hei, NL, Leong, DOE, Ginjom, IRH & Nissom, PM, 2016, ‘Ureolytic Bacteria isolated from Sarawak Limestone Caves show High Urease Enzyme Activity comparable to that of Sporosarcina pasteurii (DSM 33)’, Malaysian Journal of Microbiology. (in press). CONFERENCE PAPERS AND PROCEEDINGS
Omoregie, AI, Senian, N, Li, PY, Hei, NL, Leong, DOE, Ginjom, IRH & Nissom, PM, 2015, 'Isolation and Characterization of Urease Producing Bacteria from Sarawak Caves and Their Role in Calcite Precipitation,' International Congress of the Malaysian Society for Microbiology (ICMSM2015), Malaysian Society for Microbiology, pp. 16-21. Senian, N, Omoregie, AI, Peter Morin Nissom, Ngu, L-H & Ong, DEL, 2014, 'Identification of locally found bacteria for potential use in ground improvement works by microbially induced calcite precipitation (MICP) technique,' The 19th International Conference on Transformative Science and Engineering, Business and Social Innovation, Society for Design and Process Science, pp. 261-266. Omoregie, AI, & Nissom, PM, 2016, ‘Cross disciplinary research: developing biocement applications using local bacteria’, The fourth Borneo Research Education Conference, Universiti Teknologi Mara Sarawak, pp. 1-8. Senian, N, Khoshdelnezamiha, G, Omoregie, AI, Ong, DEL, Ngu, LH, Nissom, PM & Henry-Ginjom, IR, 2016, ‘Development of Bio-Pavers with Microbial Induced Calcite Precipitation Technique Using Sporosarcina Pasteurii,’ 19th Southeast Asian Geotechnical Conference & 2nd Association of Geotechnical Societies in SouthEast Asia Conference, Malaysian Geotechnical Society, pp. 327-331. Phua, YL, Omoregie, AI, Ong, DEL, Ngu, LH, Nissom, PM & Ginjom, IR, 2016, ‘Ground improvement via Microbial-Induced Calcite Precipitation using Push-Pull Injection System’, 19th Southeast Asian Geotechnical Conference & 2nd Association of Geotechnical Societies in SouthEast Asia Conference, Malaysian Geotechnical Society, pp. 495-498.
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PRESENTATIONS
Oral presenter, Cross disciplinary research: developing biocement applications using local bacteria, The fourth Borneo Research Education Conference (BREC), 17-18 August 2016, Kota Samarahan, Sarawak, Malaysia Poster presenter, Isolation and Characterization of Urease Producing Bacteria from Sarawak Caves and Their Role in Calcite Precipitation, International Congress of the Malaysian Society for Microbiology, 7-10 December 2015, Batu Ferringhi, Penang, Malaysia. Oral presenter, Isolation of Highly Active Urease Producing Bacteria from Sarawak Limestone Caves, The Regional Taxonomy and Ecology Conference, 1-2 December 2015, Kuching, Sarawak, Malaysia. Poster presenter, Isolation and Characterisation of Urease Producing Bacteria from Sarawak Caves and their Role in Calcite Precipitation, Asian Congress on Biotechnology, 15-19 November 2015, Kuala Lumpur, Selangor, Malaysia. AWARDS
BEST PAPER Awarded for the best paper written at the 4th Borneo Research Education Conference (BREC 2016), organised by Universiti Teknologi Mara Sarawak and Swinburne University of Technology, Sarawak campus. 17-18 August 2016, Kota Samarahan, Sarawak, Malaysia. http://www.sarawak.uitm.edu.my/brec2016 PEOPLE’S CHOICE AWARD Awarded for being one of the best oral presenters at the Three Minute Thesis (3MT) Competition organised by Swinburne University of Technology, Sarawak campus. 17 June 2015, Kuching, Sarawak, Malaysia. http://www.swinburne.edu.my/events/3MT-competition BEST POSTER PRESENTER Awarded best poster presenter for the technical session of environmental biotechnology at the Asian congress on biotechnology organised by Asian federation of biotechnology Malaysia Chapter and Universiti Putra Malaysia. 15-19 December, Kuala Lumpur, Selangor, Malaysia. http://www.acb2015.my/web/list-of-acb2015-winners
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TABLE OF CONTENTS Content Page
ABSTRACT i
ACKNOWLEDGEMENT iii
DECLARATION v
SCIENTIFIC OUTPUT vi
TABLE OF CONTENTS viii
LIST OF TABLES xi
LIST OF FIGURES xii
LIST OF ABBREVIATIONS xiv
CHAPTER 1: INTRODUCTION AND LITERATURE REVIEW
1.1 Introduction 1
1.2 Biomineralisation 3
1.2.1 Biologically induced biomineralisation 4
1.2.2 Biologically controlled biomineralisation 5
1.3 Microbially Induced Calcite Precipitation (MICP) 6
1.3.1. MICP via urea hydrolysis 10
1.3.2. Urease enzyme 12
1.3.3. Mechanism of CaCO3 precipitation 15
1.3.4. Urease Source 17
1.4 Factors Affecting the Efficiency of MICP 18
1.4.1. Concentration of reactants 18
1.4.2. pH 19
1.4.3. Temperature 20
1.4.4. Dissolved inorganic carbon 21
1.4.5. Bacteria size 21
1.4.6. Nutrients 22
1.4.7. Availability of nucleation site 22
1.5 Current Biotechnological Application of MICP 23
1.5.1. Biocementation 24
1.5.2. Creation of biological mortars 24
1.5.3. Bioremediation of cracks in concrete 25
1.5.4. Biodeposition on cementitious materials 27
1.5.5. Biogrout 28
1.5.6. Other essential applications of MICP 30
ix
1.6 Diversities of Microbial Communities in Caves 32
1.7 Screening Sarawak’s Limestone Caves for Ureolytic Bacteria 36
1.8 Aim and Objectives of the Study 39
1.9 Significance of the Study 39
1.10 Thesis Outline 39
CHAPTER 2: ISOLATION, IDENTIFICATION AND CHARACTERISATION OF
UREASE-PRODUCING BACTERIA FROM LIMESTONE CAVES OF SARAWAK
2.1 Introduction 41
2.2 Methods and materials 43
2.2.1. Sampling location and collection 43
2.2.2. Biological material 43
2.2.3. Growth medium and sterilisation 43
2.2.4. Enrichment cultures 44
2.2.5. Isolation of urea degrading bacteria 44
2.2.6. Screening for urease-producing bacteria 45
2.2.7. Preliminary identification 45
2.2.8. Molecular identification 46
2.2.9. Measurement of enzyme activity 48
2.2.10. Evaluation of microbial calcite precipitation 49
2.2.11. Bacterial growth profile and pH profile 50
2.2.12. Statistical analysis 51
2.3 Results 52
2.3.1. Sampling location and sample collection 52
2.3.2. Enrichment culturing and bacterial isolation 54
2.3.3. Selection of urease producing bacteria 55
2.3.4. Phenotypic characterisation 58
2.3.5. Molecular characterization 62
2.3.6. Measurement of conductivity 69
2.3.7. Urease Activity Assay 69
2.3.8. Determination of specific enzyme activity 73
2.3.9. Microbial calcite precipitates 77
2.3.10. Calcite estimation 78
2.3.11. Bacterial growth and pH profiles 80
2.4 Discussion 85
2.5 Conclusion 92
x
CHAPTER 3: EFFECTS OF CULTURAL CONDITIONS ON UREASE ACTIVITY
AND EVALUATION OF BIOCEMENTATION POTENTIALS IN SMALL SCALE
TEST
3.1 Introduction 93
3.2 Methods and Materials 94
3.2.1. The Effect of Cultural Conditions On Urease Activity 94
3.2.2. Small Scale Biocementation Test 95
3.3 Results 100
3.3.1. Temperature (oC) 100
3.3.2. Initial medium pH 102
3.3.3. Incubation period (hr) 104
3.3.4. Effect of urea concentration (%) 106
3.3.5. Biocementation treatment test 108
3.3.6. Soil surface strength 115
3.3.7. Compressive strength 117
3.3.8. Calcite confirmation 119
3.3.9. Calcite content Determination 120
3.4 Discussion 123
3.5 Conclusion 131
CHAPTER 4: GENERAL CONCLUSIONS AND RECOMMENDATIONS
4.1 General Conclusion 132
4.1.1. Aim of the thesis 132
4.1.2. Limestone area as source of ureolytic bacteria 133
4.1.3. Enrichment culture and isolation 134
4.1.4. Screening and identification 134
4.1.5. Measurement of urease activity 135
4.1.6. Biocementation competency of local isolates 136
4.2 Future Directions and Recommendations 136
REFERENCES 138
xi
LIST OF TABLES
Table Page
2.1 Description of samples collected from FCNR and WCNR 52
2.2 Hydrolysis of urea by isolates UAB medium 57
2.3 Morphological characteristics of isolated bacterial colonies 59
2.4 Microscopic characteristics of bacterial isolates 60
2.5 Biochemical characteristics of bacterial isolates 61
2.6 Molecular identification based on 16S rRNA sequencing data using NCBI
nucleotide BLAST database
64
2.7 The nomenclatural taxonomy obtained using Ribosomal Database Project-
II database
66
2.8 Measurement of conductivity variation rate and SEM 71
2.9 Conversion of changes in conductivity to urease activity 72
2.10 t-test results comparing the specific urease activity differences
between individual isolated urease-producing bacteria and control strain
76
2.11 t-test results comparing the calcite precipitate differences between
individual isolated urease-producing bacteria and control strain
79
2.12 Kinetics growth of ureolytic bacteria in batch cultures 81
3.1 Selected ureolytic bacteria for biocement test 95
3.2 Biocement treatment components 96
3.3 Sand characteristics 97
3.4 Sand grain size characteristics 109
3.5 Bacteria concentration and urease activity prior to biocement test 110
3.6 t-test results comparing the strength (psi) differences between the
biocemented sands
116
3.7 Unconfined compressive strength (UCS) of the treated sands 117
3.8 t-test results comparing the unconfined compressive strength (UCS)
differences between the biocemented sands
118
3.9 Summary of calcite content and compressive strength of selected
isolates and consortia
121
xii
LIST OF FIGURES Table Page
1.1 Pathway of biominerals secretion and precipitation in a bacterial cell 11
1.2 Genetic organisation of urease operon in Helicobacter pylori and
Sporosarcina pasteurii
13
1.3 Regulation levels for enzyme activity by microorganisms 14
1.4 A simplified representation of Ureolysis-driven CaCO3 precipitation 16
1.5 An in situ application of bacteria based liquid 25
1.6 Self-healing crack from the addition of bacterial metabolism via urea
hydrolysis
26
1.7 1 mm thick calcite crust formed on the surface of the soil 27
1.8 Set-up for large scale (100m3) soil treatment 29
1.9 Calcified structures of biogenic origin discovered in cave regions 34
1.10 Speleothems samples collected from El Toro and El Zancudo limestone
mines located in Cordillera Central, northeast
of Colombia
35
1.11 Map of Borneo Island showing the geographical divisions and
topographical features of Brunei Darussalam, Indonesia (Kalimantan) and
East Malaysia (Sarawak and Sabah)
38
2.1 Sampling collection site situated in FCNR, Bau, Sarawak 53
2.2 Sampling collection site situated in WCNR, Bau, Sarawak 53
2.3 Microorganisms grown on nutrient agar plates supplemented with 2% urea 54
2.4 Pure colonies of urea degrading bacteria after enrichment culture 55
2.5 Urease production test using UAB medium 56
2.6 Phylogenetic tree based on the bacterial 16S rRNA gene sequence data
sequence from different isolates of the current study along with sequences
available in the GenBank database
68
2.7 Relative conductivity of isolate LPB21 measured for a duration of 5 min 70
2.8 Specific urease activity (mM urea hydrolysed.min-1.OD-1) of urease-
producing bacteria and the control strain
75
2.9 Calcite precipitation media 77
2.10 Comparison of calcite precipitated by selected UPB isolates and the control
strain
78
xiii
2.11 Growth profile of selected ureolytic bacterial isolates and control strain
grown in nutrient broth containing 6% urea for 12 hr
82
2.12 pH profile of selected ureolytic bacterial isolates and control strain grown
in nutrient broth containing 6% urea for 12 hr
83
3.1 The effect of different temperature on urease activity 101
3.2 The effect of different pH on urease activity 103
3.3 The effect of different incubation period on urease activity 105
3.4 The effect of different urea concentration on urease activity 107
3.5 Treatment of sand column using locally isolated bacteria, consortia,
positive and negative controls
111
3.6 Sand columns at the end of treatment using ureolytic bacteria and
cementation solution
112
3.7 Treated sand removed from their respective columns 113
3.8 Treated sand sample held after a curing period and columns were
successfully removed
114
3.9 Surface strength of the biocemented sand samples 115
3.10 Confirming calcite precipitates 119
3.11 Comparison of the relative quantity of calcites in the biocemented sands 120
xiv
LIST OF ABBREVIATIONS MICP Microbially Induced Calcite Precipitation
BIM Biologically Induced Mineralisation
BCM Biologically Controlled Mineralisation
DIC Dissolved Inorganic
IAP Ion Activity Product
UDB Urea Degrading Bacteria
UPB Urease Producing Bacteria
UAB Urea Agar Base
FCNR Fairy Cave Nature Reserve
WCNR Wind Cave Nature Reserve
PCR Polymerase Chain Reaction
TE Trix EDTA
NCBI National Centre for Biotechnology Information
DNA Deoxyribonucleic Acid
BLAST Basic Local Alignment Search Tool
RDP Ribosomal Database Project
MEGA Molecular Evolutionary Genetic Analysis
CPM Calcite Precipitating Media
df Dilution Factor
ATP Adenosine Triphosphate
SUA Specific Urease Activity
UA Urease Activity
HCL Hydrochloric Acid
NaOH Sodium Hydroxide
UCS Unconfirmed Compression Strength
ATSM American Society for Testing and Materials
xv
RH Relative Humidity
SEM Standard Error of Mean
SE Standard Deviation ANOVA
Analysis of Variance
Chapter
1 INTRODUCTION AND LITERATURE REVIEW
1
1.1 Introduction Enzyme technology is a well-established branch of biotechnology undergoing a
development phase (Binod et al., 2013), and their functional significance suggests many
novel application especially for environmentally-friendly industrial purposes (Binod et
al., 2013). Enzymes from microorganisms are an essential source of numerous
industrially relevant enzymes (Ibrahim, 2008). Microbial enzymes are relatively more
stable and properties more diverse than other enzymes derived from plants and animals
(Alves et al., 2014). Enzymes produced from microorganisms can be easily controlled
physiologically, physio-chemically, have quantitative production and mostly extracted
with low production cost extracellularly using downstream processes (Ibrahim, 2008,
Pandey et al., 2010). The industrial usage of the microbial enzymatic process are
classified as (i) Enzymes as final products; (ii) Enzymes as processing aids; (iii)
enzymes in food and beverage production; (iv) Enzymes in genetic engineering and (v)
Enzymes as an industrial biocatalyst (Binod et al., 2013).
Microbially induced calcite precipitation (MICP) is a comparatively innovative soil
improvement technique which requires the production of urease enzyme from bacteria
for soil treatment (Soon, 2013). Modern ground improvement techniques have become
increasingly complex due to sustainability consideration and the expedition of reducing
environmental pollution (Kavazanjian and Hamdan, 2015). Established materials and
methods often require replacement or supplemented by innovative materials which are
environmentally friendly (Kavazanjian and Hamdan, 2015). Existing ground
improvement techniques such a chemical grouting has been proven to have an effective
performance in the increment of soil’s shear strength and stiffness, however,
environmental and human health concerns over their applications have deemed them as
unsustainable materials (DeJong et al., 2010). Portland cement is a major construction
material of choice in building, structure and ground improvement applications in order
to meet the increasing demand of rapid industrialisation and urbanisation (Siddique et
al., 2016). However, the use of Portland cement is associated with certain challenges
such as energy , resource conservation, the cost of production and greenhouse gas
emission (Kavazanjian and Hamdan, 2015). It is estimated that production of Portland
cement clinker solely contributes about 7% global CO2 emission, this makes this
construction material an unsustainable construction material (Jonkers et al., 2010).
2
MICP has been exploited in recent decades as an alternative building material to
Portland cement through either direct substitution or complementary usage
(Kavazanjian and Hamdan, 2015, DeJong et al., 2013). MICP applications require lesser
energy for production, low production cost and no contribution to the greenhouse gas
emission, making it an environmentally friendly construction material (Achal, 2015).
Existing research studies suggests that biocementation technology can be used to
address important geotechnical problems in granular soils which include slope stability,
erosion, stiffness and stress-permeability, tunnelling and liquefaction (van Paassen et
al., 2010, DeJong et al., 2010, DeJong et al., 2011).
Bacteria acts as primary agents of geochemical changes due to their high surface area to
volume ratio, their widespread abundant distribution, evolutionary adaptiveness, diverse
enzymatic and nutritional possibilities (Warren and Haack, 2001). Numerous microbial
species from extremely diverse environments have been linked to the process of
microbial precipitation of calcium carbonate (Hammes, 2003). Calcium carbonate is the
most reactive mineral on earth, composing 4% of the earth’s weight (Whiffin, 2004), it
is constantly involved in processes of dissolution and precipitation (Hammes et al.,
2003b, Hammes and Verstraete, 2002). Carbonaceous minerals are frequently found in
oceans, soils, and geological formations, representing an important segment of the
global carbon pool (Hammes, 2003). The primary role of bacteria in calcium carbonate
precipitation has been subsequently ascribed to their capability to create an alkaline
environment through numerous biological and chemical activities (Fujita et al., 2000,
Castanier et al., 2000, Castanier et al., 1999). Characterisation of microorganisms by
genera and species which were previously unachievable through biochemical methods
alone are now being executed with the use of sequence-classifier algorithms (Ercole et
al., 2007). The ease in microbial identification using traditional and molecular
methodology can aid in understanding and identify wider ranges of the microorganism
of a given community (Rajendhran and Gunasekaran, 2011), with the capability of
producing urease enzyme, and induce microbial calcite sufficient for MICP
applications.
3
1.2 Biomineralisation
Biomineralisation is the reformation of chemicals (Anbu et al., 2016) in a
microenvironment caused by the activity of microorganisms which result in the
precipitation of minerals (Phillips et al., 2013, Barkay and Schaefer, 2001, Stocks-
Fischer et al., 1999). In nature, biomineralisation results in the formation of sixty (or
more) various biological minerals, which exists as extracellular or intracellular
inorganic crystals, although some precipitation of inorganic minerals contains trace
elements of organic compounds (Dhami et al., 2013b, Yoshida et al., 2010, Konishi et
al., 2006). It is anticipated that the number of biominerals formed will continue to
increase (Defarge et al., 2009).
Biominerals are distinguished based on their properties such as size, shape, crystalline
nature and elemental composition (isotopes and trace) (Sarayu et al., 2014). Minerals
which are formed through biologically induced mineralisation, through passive surface-
mediation includes iron (Fe), manganese (Mn), carbonates, phosphonates and silicates.
Calcium carbonate (CaCO3) is a biomineral widely secreted by most microorganisms
(Sarayu et al., 2014, Barabesi et al., 2007). Calcium carbonate mineralisation can be
found in natural formations such as corals, ant hills or caves (Dhami et al., 2013d). Out
of the eight polymorphs of calcium carbonate, seven are crystalline and one is
amorphous (Weiner and Dove, 2003). Calcite, aragonite, and vaterite are pure calcium
carbonate, while two-monohydrocalcite and the stable form of amorphous calcium
carbonate contain one water molecule per calcium carbonate (Weiner and Dove, 2003),
however, the temporary forms of amorphous calcium carbonate do not contain water
(Addadi et al., 2003).
Carbonate minerals precipitated by microorganisms contributes about 50% of the total
biominerals formed, while phosphate minerals contribute 25% of the precipitated
minerals by microbial species (Sarayu et al., 2014). These minerals are usually formed
in high quantities and widespread in nature (Ramesh Kumar and Iyer, 2011, Weiss et
al., 2002). Biominerals have unusual morphologies as they are often defined by the
complexity and variety of secreting microorganisms (Bazylinski and Frankel, 2003).
4
Biomineralisation process is divided into two different fundamental groups which are
based on the degree of their biological control (Sarayu et al., 2014). These groups are
known as biologically induced and biologically controlled mineralisation (Weiner and
Dove, 2003). Lowenstam (1981) introduced these two groups as “biological induced”
and “organic matrix-mediated” mineralisation, however, the latter was renamed by
Mann (1983) to “biologically controlled mineralisation”, recognising that the process of
biomineralisation within these conversions varies with different microorganisms.
1.2.1 Biologically induced biomineralisation
Biologically induced mineralisation (BIM) involves the interaction of the environment
and biological activities resulting in mineral precipitation (Sarayu et al., 2014). In this
type of situation, microbial cell surfaces often act as a causative agent for nucleation
and subsequent growth of the minerals (Weiner and Dove, 2003). These type of
biominerals are often secreted to the metabolism of the microorganisms, and the
systems have little or no control over the minerals which are being deposited (Sarayu et
al., 2014). The precipitation of extracellular by-product of the microbial metabolism can
lead to random crystallisation and non-specific crystal morphologies (Provencio and
Polyak, 2001).
The organelles of these microbes take part in the process of BIM, the cell wall acts as
nucleation sites (Sarayu et al., 2014). Once these biominerals are synthesized, the pH,
CO2, and composition of the microenvironments of the microorganisms are often altered
and any changes in the microorganisms will adversely have an effect on the secreted
biominerals because the whole process of BIM depends primarily on the circumstances
prevailing in the microorganism (Frankel and Bazylinski, 2003, Tebo et al., 1997,
Fortin et al., 1997). BIM process results in engulfment of the whole cell of the
microorganisms by biominerals secretions, which causes an encrustation (Sarayu et al.,
2014). The distinctive feature of BIM is that biominerals, when deposited are usually
formed along the surfaces of the microbial cells where they remain firmly attached to
the cell wall and organic components of the cell wall (lipids, proteins, and
polysaccharide) can influence the process in BIM (Mann, 2001).
5
1.2.2 Biologically controlled biomineralisation
Biologically controlled mineralisation (BCM) due to cellular activities of
microorganism are classified into extracellular, intercellular and intracellular
participations of the microbes (Sarayu et al., 2014). In extracellular participation,
macromolecular matrix (made up of proteins, polysaccharides, and glycoproteins)
situated outside the cell acts as the site of mineralisation, which is related to BIM
(Sarayu et al., 2014). The genes which are responsible play effective roles in
determining the structures and compositions which are integrated with the regulation
and organisation of the composite formation (Weiss et al., 2002). The matrix
composition is unique and contains a high proportion of acidic amino acids (Swift and
Wheeler, 1992).
The structures and compositions are genetically programmed to execute vital regulating
roles which result in composite biominerals formation (Weiner and Dove, 2003). The
intercellular participation is seen in a microorganism that lives as communities (Sarayu
et al., 2014). The minerals which are secreted by these microbes nucleates in the
epithelial cells and fill the intercellular space in a particular orientation which resembles
an exoskeleton (Young and Henriksen, 2003). The intracellular involvement is an
extremely controlled mechanism which precipitates minerals that direct the nucleation
of the biominerals inside the cells, these compositions are then governed by the
environments insides the vesicles or vacuoles usually determined by the specificity of
the species (Rodriguez-Navarro et al., 2012). Some of the species-specific
crystallochemical properties include uniform particle sizes, high level of spatial
organisation, complex morphologies, and well-defined structure and composition
(Mann, 2001).
6
1.3 Microbially Induced Calcite Precipitation (MICP)
Natural lithification of sediment occurs due to physical, chemical and biological
processes (Gadd, 2010) which result in deposition of minerals in the sediments, these
minerals compact the sediments together, reducing pore space together, eliminating
water permeability and causing cementation to occur (Paassen, 2009). However,
production of these minerals which results in a compartment of sediments undergoes a
very slow process (Paassen, 2009). On the other hand, mineralization using biological
process can accelerate cementation, the microorganisms (when supplied with suitable
substrates) are able to catalyse chemical reactions leading to a dissolution or
precipitation of inorganic minerals which aids in changing the properties of soil
(Paassen et al., 2009, Paassen, 2009).
Microbially induced calcite precipitation (MICP) is a process that refers to calcite
precipitation from a supersaturated solution in a microenvironment that occurs due to
the occurrence of microbial and biochemical activities (Hamilton, 2003, Bosak, 2011,
Anbu et al., 2016). MICP utilises the biologically induced pathway of biomineralisation
(Whiffin et al., 2007, Whiffin, 2004). During MICP process, microorganisms are able to
produce metabolic products (CO32-) that react with ions (Ca2+) in the microenvironment
which results in consequent minerals precipitated (Anbu et al., 2016). The ability of
microorganisms to induce biomineralisations, both in natural and laboratory conditions
are influenced by the type of microbes involved (Dhami et al., 2012a), salinity and
compositions of nutrients available in the microenvironments (Rivadeneyra et al., 2004,
Knorre and Krumbein, 2000).
CaCO3 is one of the utmost prevalent minerals on earth, mostly found in rocks, fresh or
marine water and soils (Castanier et al., 1999, Ehrlich, 1998). CaCO3 precipitation
occurs usually when the amount of calcium and carbonate ions in the solution exceeds
the product solubility (Cheng, 2012). Comparing contributions of abiotic change such
as a change in temperature, pressure or evaporation and biotic action which involves
microbial activity, it is suggested that biotic actions have a greater level of contribution
in inducing CaCO3 precipitates in most environments on earth (Castanier et al., 2000).
7
CaCO3 precipitation is a rather straightforward chemical process often governed by four
main key factors (Dhami et al., 2013b): (1) the calcium concentration, (2) the
concentration of dissolved inorganic carbon (DIC), (3) the pH and (4) the availability of
nucleation sites (Hammes and Verstraete, 2002). CaCO3 precipitation requires
sufficient calcium and carbonate ions so that the ion activity product (IAP) exceeds the
solubility constant (Kso) as shown in Equations (1.1) to (1.3) (Dhami et al., 2014, Dhami
et al., 2013b). From the comparison of the IAP with the Kso , the saturation state (Ω) of
the system can be defined; if Ω > 1 (Dhami et al., 2014), then an oversaturation and
precipitation will occur in the system as mentioned below by Morse (1983):
(1.1) Ca2+ + CO3
2- ↔ CaCO3
(1.2) Ω = a (Ca2+) a (CO32-) / Kso
(1.3) with Kso calcite, 25oC = 4.8 x 10-9
As previously mentioned, the concentration of DIC and the pH of the microenvironment
influences the concentration of carbonate ions (Dhami et al., 2014, Dhami et al., 2013b).
However, DIC concentration relies on environmental parameters such as temperature
and partial pressure of carbon dioxide for the systems which are exposed to the
atmosphere (Cheng, 2012, Dhami et al., 2013b). The equilibrium reactions and constant
which governs the DIC concentration in aqueous media (25oC and 1 atm) are given in
Equations (1.4) to (1.8) as suggested by Stumm and Morgan (1981):
(1.4) CO2 (g) ↔ CO2 (aqueous) (pKH = 1.468)
(1.5) CO2 (aqueous) + H2O ↔ H2CO3 (pK= 2.84)
(1.6) H2CO3 ↔ H+ + HCO3- (pK1 = 6.352)
(1.7) HCO3− ↔ CO32− + H+ (pK2 = 10.329)
(1.8) With H2CO3 = CO2(aqueous) + H2CO3
8
CaCO3 precipitation is very slow under normal conditions which require a long
geological time, however, MICP can produce a large amount of carbonate in shorter
duration (Dhami et al., 2013b). Exploratory research involving MICP has gained an
increased interest in the last 20 years, with the primary focus of research in
biotechnology, applied microbiology, geotechnical and civil engineering, due to the
numerous applications of MICP (Dhami et al., 2014). Various bacterial species are
capable of inducing calcite precipitates in alkaline environments rich in Ca2+ ions
(Dhami et al., 2013b) and other mechanisms in natural habitats (Rivadeneyra et al.,
2004, Ehrlich, 1996).
There are mainly four groups of microorganisms which are involved in the MICP
process (Dhami et al., 2013b), namely: (i) photosynthetic microorganisms such as
cyanobacteria and algae, (ii) sulphate reducing bacteria responsible for dissimilatory
reduction of sulphates, (iii) microorganism utilizing organic acids, and (iv)
microorganisms involved in nitrogen cycle either by ammonification of amino
acids/nitrate reduction or hydrolysis of urea (Jargeat et al., 2003, Hammes and
Verstraete, 2002, Stocks-Fischer et al., 1999).
In the aquatic environment, MICP is primarily caused by photosynthetic
microorganisms (McConnaughey and Whelan, 1997). Algae and cyanobacterial
metabolic processes utilize dissolved CO2 (Dhami et al., 2013b) and calcium ions to
induce CaCO3 precipitations as shown in Equation (1.9) to (1.12) (Hammes and
Verstraete, 2002). CaCO3 precipitation (dolomites and aragonite) via this route often
happens in the seawater, geological formations, landfill leachates and during biological
treatment of acid mine drainage (Machel, 2001, Warthmann et al., 2000, Wright, 1999).
(1.9) CO2 + H2O −→ (CH2O) + O2
(1.10) 2HCO3- ↔ CO2 + CO3
2− + H2O
(1.11) CO3 2− + H2O ↔ HCO3
- +OH−
(1.12) Ca2+ + HCO3- + OH− → CaCO3 + 2H2O
9
Heterotrophic microorganisms are also capable of inducing CaCO3 precipitation by the
production of carbonate or bicarbonate and modification of the microenvironment
which favours the precipitations (Castanier et al., 1999). The abiotic dissolution of
gypsum provides an environment that is rich in sulphate and calcium ions, the presence
of organic matter and absence of oxygens allows sulphate reducing bacteria to reduce
sulphate to hydrogen sulphite (Whiffin, 2004) as shown in Equation (1.13) and (1.14)
(Wright, 1999, Castanier et al., 1999, Ehrlich, 1998).
(1.13) CaSO4·2H2O → Ca2+ + SO4 2− + 2H2O
(1.14) 2(CH2O) + SO4 2− → HS−+HCO3- +CO2+H2O
The third pathway involved in CaCO3 precipitation involves bacteria which use organic
acids as their only carbon and energy sources wherein some common soil bacteria
species are included (Dhami et al., 2014). The consumption of these acids results in pH
increase which leads to CaCO3 precipitation in the presence of calcium ions as shown
in Equation (1.15) to (1.17) (Braissant et al., 2002, Knorre and Krumbein, 2000).
(1.15) CH3COO− + 2O2 → CO2 + H2O +OH−
(1.16) 2CO2 + OH− → CO2+ HCO3-
(1.17) 2HCO3-+ Ca2+ → CaCO3 + CO2 + H2O
Various heterogeneous bacterial groups are linked to this pathway for MICP process
(Dhami et al., 2014). Braissant et al. (2002) suggested that this pathway might be
extremely common in natural environment due to the abundance of low molecular
weight acids in soils, especially by fungi and plants. The fourth pathway of MICP
process involves microorganisms in nitrogen cycle via hydrolysis of urea. This pathway
is the easiest and most used method of MICP involving several applications (Dhami et
al., 2013b).This is attributed to the ability of the urea hydrolysis pathway to induce a
high amount of CaCO3 precipitates (Sarayu et al., 2014, Qabany et al., 2012, Siddique
and Chahal, 2011).
10
1.3.1. MICP via urea hydrolysis
CaCO3 precipitation by bacteria through urea hydrolysis is the most straightforward and
easily controlled mechanism of MICP with the ability to induce high amount of CaCO3
in a short duration of time (Dhami et al., 2014).
(1.18) CO(NH2)2 + H2O NH2COOH + NH3
(1.19) NH2COOH + H2O → NH3 + H2CO3
(1.20) H2CO3 → 2H++2CO32-
(1.21) NH3 + H2O → NH4++ OH−
(1.22) Ca2+ + 2CO32- →CaCO3 (KSP = 3.8 × 10−9)
KSP is the solubility product shown in Equation (21).
Stocks-Fischer et al. (1999) suggested that during microbial urease activity, 1 mol of
urea is hydrolyzed intracellularly to 1 mol of carbonate, which spontaneously
hydrolyzes to form an additional 1 mol of ammonia and carbonic ions. The ammonia
and carbonic ions equilibrate in water to form bicarbonates, 1 mol of ammonium and
hydroxide ions which allows an increases the pH of the environment as shown in
Equation (1.18) to (1.22) (Stocks-Fischer et al., 1999). Urease enzyme is responsible for
catalysing the hydrolysis of urea to produce ammonia and carbonate ions (Mobley and
Hausinger, 1989).
microbial urease
11
Figure 1.1: Pathway of biominerals secretion and precipitation in the cell of a bacteria. The bacteria serve a nucleation site for CaCO3 precipitation in the microenvironment (Sarayu et al., 2014). An ATP-generating system coupled with urea hydrolysis process in Sporosarcina pasteurii was suggested by Jahns (1996) and Whiffin (2004). The chemical transport processes which are related to microbial urea hydrolysis was (Mobley and Hausinger, 1989). The leading function of bacteria has been linked to their capability to generate an
alkaline microenvironment (Kumari, 2015) through various biological and chemical
activities as shown in Figure 1.1 (Dhami et al., 2014, Dhami et al., 2013b). The
bacteria’s surface plays an essential role in CaCO3 precipitates (Fortin et al., 1997). Due
to the presence of various negatively charged groups, at a neutral pH, positively charged
metal ions are able to bind to bacteria’s surfaces, favouring heterogeneous nucleation
(Douglas and Beveridge, 1998, Bäuerlein, 2003). The precipitation of CaCO3 on the
external surface of the bacterial cells often occurs by successive stratification, which
makes the cells become embedded in growing CaCO3 crystals (Castanier et al., 1999,
Rivadeneyra et al., 1998).
12
1.3.2. Urease enzyme
Urease and its substrate urea represent an important milestone in the early scientific
investigation (Mora and Arioli, 2014). Urease is produced by many diverse bacterial
species which includes normal flora and non-pathogens (Mobley, 2001). The scientific
interest in microbial urease was previously related to the relevance of this enzymatic
activity in infection (Mora and Arioli, 2014). This interest was strongly stimulated since
the discovery of the association of Helicobacter pylori with gastritis and stomach cancer
(Mobley et al., 1995). Urease has also been demonstrated as a potent virulence factor
for some bacterial species which include Proteus mirabilis, Staphylococcus
saprophyticus and Helicobacter pylori (Eaton et al., 1991, Jones et al., 1990, Gatermann
and Marre, 1989).
1.3.2 (a): Molecular characterisation of urease genes
Microbial ureases are multi-subunit metalloenzymes that hydrolyse urea substrates to
form carbonic acid and two molecules of ammonia (Mobley et al., 1995). The
degradation of urea provides ammonium for integration into intracellular metabolites
and enables the survival of the microorganism in acidic environments (Collins and
D'Orazio, 1993, Mobley et al., 1995). The structure of urease was first explained by
Jabri et al. (1995), showing that ureases may be composed of up to three distinctive
types of subunits, indicating that all the proteins are closely related. The structural genes
that encode both the urease subunits, ureA, ureB, and ureC, and the accessory proteins
required for assembly of the urease nickel metallocenter are typically clustered at a
single locus (Mobley et al., 1995). Different patterns of urease expression have been
observed in various bacteria (Wray et al., 1997).
There are eight genes which are necessary for the synthesis of urease enzyme,
designated as ureA; -B; -I; -E; -F; -G; -H and -I (Hu and Mobley, 1993, Hu et al., 1992,
Cussac et al., 1992, Ernst et al., 2007). Urease genes are evolutionarily related to each
other, sharing a common an ancestor (Ernst et al., 2007). Urease of Helicobacter pylori
is composed of two subunits, UreA (27 kDa) and UreB (62 kDa) and the subunits form
a multimeric enzyme complex with spherical assembly (Labigne et al., 1991, Clayton et
al., 1990, Ernst et al., 2007).
13
Figure 1.2: Genetic organisation of urease operon in Helicobacter pylori and Sporosarcina pasteurii. The ureAB genes of the ancestral urease operon are fused and labelled ureA, the ancestral ureC is labelled ureB in Helicobacter pylori (Ernst et al., 2007). In Helicobacter pylori, ureA and ureB are fused together to create ureA gene, while
ureC gene is labelled as ureB as shown in Figure 1.2. on the other hand, in
Sporosarcina pasteurii, the ancestral genes ureA and ureB are not joined together
(Figure 1.2).The ureEFGH genes codes for urease accessory proteins, which aid in
mediating proper formation of the complex quaternary structure and also transport
nickel ions into the urease enzyme active centre (Ernst et al., 2007). The ureI gene
codes for pH which regulates the urea channel situated in the cytoplasmic membrane
(Akada et al., 2000). ureI and ureA also interact during urea hydrolysis at the cell wall
of bacteria, allowing fast diffusion of ammonia and CO2 to occur (Voland et al., 2003).
1.3.2 (b): Activity of urease enzyme
Urease activity (UA) is the urea hydrolysis activity produced by the enzyme urease per
minute (Alhour, 2013). The process of urease production is illustrated in Figure 1.3
(Whiffin, 2004). Enzyme activity regulation is vital for energy efficiency in cell
function, however not all enzymes are mandatory all the time and their synthesis can
either be turned “off” (repressed) or “on” (induced ) depending the presence or absence
of metabolites (Whiffin, 2004). This type of genetic control is often regulated by the
cell at the transcriptional level where messenger RNA is produced from the DNA
template (Ratledge, 2001, Lewin, 1994). Enzymes such as urease can be controlled at
the transcription (inducible/repressible) level are usually repressed under normal
conditions, which helps to converse energy from unnecessary protein synthesis
(Whiffin, 2004). The presence of an inducer, normally its substrate, can strongly induce
an energy up to 1000-fold its level under non-induced conditions (Lowe, 2001).
14
Figure 1.3: Regulation levels for enzyme activity by microorganisms. The enzyme can be regulated at the transcriptional level or modification level (Whiffin, 2004). The genetic control is regulated by the microorganism’s cell where the messenger RNA (mRNA) codes for the enzyme which is produced from the DNA template (Ratledge, 2001, Lewin, 1994).
Whiffin (2004) determined microbial urease activity by measuring the relative change
in conductivity (mS.cm-1) when exposed to urea under standard conditions of 1.11 M
urea at 25oC. A standard curve was generated by determining the conductivity change
resulting from complete hydrolysis of several concentrations (50mM-250mM) of urea
by purified urease (Sigma Cat. No. U-7127) (Whiffin, 2004). From the standard curve
of changes in conductivity (mS.cm-1.min-1), Whiffin (2004) determined the equations
required to calculate the urease activity (mM urea hydrolysed.min-1) and the specific
urease activity (mM urea hydrolysed.min-1.OD-1) of ureolytic bacteria as shown in
Equation (1.23) and (1.24):
(1.23) Urea hydrolysed (mM) = Conductivity variation rate x (df) x (11.11)
(1.24) Specific urease activity = urease activity /Biomass
15
From Equation (1.23), urease activity (mM urea hydrolysed.min-1) was calculated by
multiplying the conductivity variation rate (mS.cm-1.min-1) by dilution factor (df) and
11.11 (correlation rate). According to Whiffin et al. (2007) 1 mS.cm-1.min-1 corresponds
to a hydrolysis activity of 11 mM urea.min-1 in the measured range of activities
considering the dilution of the culture during the activity measurement by a factor of 10
(Cheng and Cord-Ruwisch, 2013). From Equation (1.24), specific urease activity (mM
urea hydrolysed.min-1.OD-1) was calculated by dividing urease activity (mM urea
hydrolysed.min-1) by biomass (OD600). According to Whiffin (2004), the biomass
concentration was measured at the end of incubation period (overnight cultivation).
1.3.3. Mechanism of CaCO3 precipitation
CaCO3 Precipitation involves: (i) The development of supersaturation solution, (ii)
Nucleation (the formation of new crystals) begins at the point of critical saturation
and (iii) Spontaneous crystal growth on the stable nuclei (Alhour, 2013). CaCO3
precipitation occurs at the bacterial cell surface if there are sufficient concentration of
Ca2+ and CO32− in solution (Figure 1.4) (Anbu et al., 2016). The biochemical reaction
that takes places in the urea-CaCl2 medium leads to precipitation of CaCO3 as shown in
Equation (1.25) to (1.27), act as binders in between the substrate particles was
suggested by Stocks-Fischer et al. (1999).
(1.25) Ca2+ + Cell → Cell − Ca2+ (1.26) Cl − + HCO3− + NH3 → NH4Cl + CO3
2− (1.27) Cell − Ca2++ CO3
2− → Cell − CaCO3
16
Figure 1.4: A simplified representation of Ureolysis-driven CaCO3 precipitation. (A) Bacteria uptake urea and release ammonium (AMM) and dissolved inorganic carbon (DIC), bacterial cells attract calcium ions. (B) A local super-saturation occurs in the presence of calcium ions, resulting in CaCO3 precipitation on the bacterial cell wall. (C)The whole cell is encapsulated (De Muynck et al., 2010b). There are different phases of the CaCO3 precipitated by the bacteria which are: the three
anhydrous polymorphs (calcite, vaterite, and aragonite); two hydrated crystalline phases
(monohydrocalcite and ikaite); and various amorphous phases with different hydration
ranges (Rieger et al., 2007, Gower, 2008, Gebauer et al., 2010). Monohydrocalcite and
aragonite have been reported to be secreted by the bacteria (Gebauer et al., 2010,
Sanchez-Navas et al., 2009), It is also suggested that the proteins of Bacillus firmus and
Bacillus sphaericus are present in the extracellular polymeric substances which controls
the aragonite or calcite polymorph selection and calcium carbonate precipitation
(Kawaguchi and Decho, 2002). Lian et al. (2006) have also suggested that the cells and
the extracellular polymeric substances of Bacillus megaterium have controlled the
precipitation of calcite and vaterite. Similarly, Myxococcus sp. was also been reported to
have precipitated vaterite and calcite with varying morphologies along with other
minerals such as phosphate and sulphate, however depending on the medium that was
being used for culturing (Sarayu et al., 2014)
17
1.3.4. Urease Source
In a review by Sarayu et al. (2014), a list of bacteria that have been reported to induce
CaCO3 precipitates was tabularized. Some of these bacteria listed as Pseudomonas
putida, Arthrobacter sp., Desulfovibrio desulfuricans, Phormidium crobyanum and
Homoeothrix crustaceans (Sarayu et al., 2014). Out of the forty-one bacteria, only a few
are known to produce urease enzyme. Most urease producing bacteria which have been
reported to induce CaCO3 precipitates and have been used for MICP applications are of
Bacillus genus. Ureolytic bacteria which have been reported in literature for MICP
applications are as Bacillus sphaericus and Sporosarcina pasteurii used for to heal
concrete cracks(De-Belie and De-Muynck, 2008, Ramachandran et al., 2001, De-
Muynck et al., 2008); Bacillus pseudifirmus and Bacillus cohnii used to treat surfaces of
concrete (Jonkers and Schlangen, 2007, Jonkers, 2007); and Bacillus cereus and
Shewanella as cement mortar (Achal et al., 2011, Achal and Pan, 2011, Ramachandran
et al., 2001).
The majority of urease producing bacteria which have been reported were mostly from
soils and sludge samples. Alhour (2013) reported to have isolated thirty-two ureolytic
bacteria (closely related to Bacillus licheniformis, Bacillus lentus, Bacillus cereus,
Psuedomonas antarcticus, Psuedomonas apiaries, Bacillus carboniphilus, Bacillus
subtilis, Psuedomonas borealis, Bacillus sporothermodrans, Bacillus lequilensis,
Psuedomonas cellulositropicus, Bacillus mycoides, Lysinbacillus sphaericus,
Panibacillus barcinonesis, Bacillus isabeliae and Bacillus fordii)from soil, sludge and
freshly cut concrete surface samples collected at three locations in Gaza Strip. Al-
Thawadi and Cord-Ruwisch (2012) reported they isolated three ureolytic bacteria
(closely related to Bacillus aqaarimus and Sporosarcina pasteurii) from activated
sludge samples from a wastewater treatment plant collected at different locations in
Woodman Point, Perth, Western Australia. Dhami et al. (2013d) reported they isolated
five ureolytic bacteria (closely related to Bacillus megaterium, Bacillus cereus, Bacillus
thuringiensis, Bacillus subtilis and Lysinibacillus fusiformis) from calcareous soil
samples collected at Anantapur District, Andhra Pradesh, India. Hammes et al. (2003b)
reported they isolated twelve ureolytic bacteria (closesly related to Sporosarcina
pasteurii, Bacillus psychrophilus, Planococcus okeanokoites, Bacillus globisporus and
Filibacter limicola from garden soil, landfill soils, freshly cut concrete surface and a
calcareous sludge from a biocatalytic calcification reactor collected at Ghent, Belgium.
18
Ghashghaei and Emtiazi (2013) reported they isolated twelve ureolytic bacteria (closely
related to Enterobacter ludwigii) from soil, freshwater, chalk, cement and activated
sludge samples. Achal et al. (2010b) reported they isolated two ureolytic bacteria
(closely related to Bacillus cereus and Bacillus fusiformis) from cement samples
collected from commercial bags. Achal and Pan (2011) reported they isolated three
ureolytic bacteria (closely related to Sporosarcina pasteurii, Bacillus megaterium, and
Bacillus simplex) from alkaline soil samples collected at Bhagalpur, India. Stabnikov et
al. (2013) reported they isolated three ureolytic bacteria (closely related to Sporosarcina
pasteurii and Staphylococcus succinus) from tropical beach sand (Singapore), garden
sand soil (Kiev, Ukraine) and water samples (The Dead Sea in Jordan resort, resort).
1.4 Factors Affecting the Efficiency of MICP
Urease activity and the amount of calcite precipitated during MICP process are based on
various environmental factors, including pH, temperature, bacterial size and cell
concentration (Anbu et al., 2016, Qabany et al., 2012, Soon et al., 2012).
1.4.1. Concentration of reactants
Calcium ions in bacteria's environment play a major role in inducing calcite
precipitation (Sarayu et al., 2014). Microbial cell surfaces are negatively charged which
acts as scavengers for cations such as Ca2+ and bind to the cell surfaces in aquatic
environments (Ramachandran et al., 2001, Stocks-Fischer et al., 1999). Bicarbonate
which is produced by bacterial cell gets released when it combines with the calcium
ions available in the environment to precipitate CaCO3 (Sarayu et al., 2014). Hence,
calcium ions involved in this mechanism is supplied either by the medium or may result
from the support material to which the bacterium is attached to (Rodriguez-Navarro et
al., 2012). It safeguards the fixation of the surplus toxic calcium in the environment,
which enables the bacteria to survive in unfavourable conditions (Rodriguez-Navarro et
al., 2012). A reaction between urea and calcium ions results in calcite formation.
However, a solution containing equimolar of 1 mole of calcium chloride and 1 mole of
urea provides better conversion to calcite (Nemati et al., 2005).
19
A lower concentration of cementation reagents adds to a satisfactory level of
ammonium decomposition which might enhance microbial activity (Soon et al., 2012).
Higher concentration of cementation reagents (urea and calcium ions) extends the
precipitation of calcite induced during MICP process (Nemati et al., 2005, Okwadha
and Li, 2010). It was also confirmed in a study conducted by De Muynck et al. (2010b),
whereby the weight of soil samples increased when a higher concentration of
cementation reagents was added compared to the addition of lower concentration.
However, a considerable amount of salinity has an inhibitory effect on microbial
activity, urease production, and calcite precipitation which is mainly contributed by
calcium salts (Soon et al., 2012, Rivadeneyra et al., 1998). In some cases, urease
production is still readily available for MICP process at high salinity. However, the
ratio of actual calcite precipitated and abstract calcite composition decreases when there
is an increase in reactant concentrations (Nemati and Voordouw, 2003, De Muynck et
al., 2010b). Salinity has less inhibitory effects on moderately halophilic bacteria
compare to those non-halophilic bacteria (Soon et al., 2012). Several moderate
halophilic bacteria were studied for calcite precipitation in salinity environment
(Rivadeneyra et al., 2000, Stocks-Fischer et al., 1999, Rivadeneyra et al., 1998).
Moderate halophilic bacteria are capable of growing at a wide range of salinity. Hence,
they should be used for soil treatment during biocementation application if the soil
environment contains high salinity (Rivadeneyra et al., 2004).
1.4.2. pH
The pH environmental of urease-producing bacteria is one of the important aspects of
MICP process. The chemical compositions of the in vivo fluids and adjacent to the sites
of the minerals formation is directly influential to the understanding of
biomineralisation processes (Soon, 2013). The pH of the environment controls the
survival and the metabolic activity of the microorganisms that indirectly monitors the
secretion of the products (Soon et al., 2012). High pH conditions favour the formation
of CO32– from HCO3– which leads to calcification of the generated bicarbonate (Knoll,
2003). Stocks-Fischer et al. (1999) stated that the optimum pH for urease ranges
between 7.0 to 8.0, which was further supported by the research findings of Evans et al.
(1991) and Arunachalam et al. (2010).
20
Stocks-Fischer et al. (1999) also reported that urease activity rapidly increased from pH
6.0 to 8.0, until it reached its peak (pH 8.0) and gradually decreased when at higher pH.
However, Soon et al. (2012) stated that urease activity is still viable at pH 9.0. A recent
study by Gat et al. (2014) showed that urea hydrolysis leads to an increase in the pH of
growth medium due to the production of ammonium as was indeed found in treatment
using Sporosarcina pasteurii. On the other hand, co-culture which included Bacillus
subtilis showed a decrease which correlated in time with the exponential growth phase
of Bacillus subtilis. They suggested that and may, therefore, be attributed to increased
respiration, leading to enrichment in CO2, thus acidifying the medium. A study by Sidik
et al. (2015), which focused on the process of bacterial calcium carbonate precipitation
in organic soil showed that when soils samples were treated with the bacterial solution,
the pH values fluctuated between 9 to 9.4 during the period the sand samples were
being treatment. It indicated that this range, that the treatment medium used was
appropriate for MICP process as suggested by DeJong et al. (2010).
1.4.3. Temperature
Enzymatic reactions such as urea hydrolysis by urease are dependent on temperature
(Anbu et al., 2016). The optimum temperature which favours urease hydrolysis ranges
between 20 to 37oC (Okwadha and Li, 2010, Mitchell and Santamarina, 2005),
however, enzymatic reactions for optimum production is influenced by environmental
conditions and the concentration of reactants in the system (Anbu et al., 2016). A study
performed by Mitchell and Ferris (2005) reported that urease activity increased between
5 to 10 times when temperature increased between 10 to 20oC. Ferris et al. (2003) and
Dhami et al. (2014) investigated the kinetic rate of urease and temperature on
Sporosarcina pasteurii. Their findings showed that urease was very stable at 35oC, but
the enzymatic activity decreased by 47% when the temperature increased to 55oC.
However, other studies reported by Chen et al. (1996) and Liang et al. (2005) on
temperature effects on urease activity showed that optimum 60oC was the optimum
temperature for the production of urease. This temperature for urease activity is
impractical on site for soil treatment using MICP (Soon et al., 2012).
21
1.4.4. Dissolved inorganic carbon
Inorganic carbon present in the environment plays a major role in MICP process (Soon,
2013). Dissolved inorganic carbon (H2CO3 + HCO3−+ CO3
2−), is a major product of
microbial respiration which affects microbial activities and its alkalinity (Sauvage et al.,
2014, D'Hondt et al., 2002). The DIC released from the extracellular polysaccharide of
the microorganisms complexes the calcium ions, thus reducing calcium carbonate
saturation enhancing the calcite precipitation (Tourney and Ngwenya, 2009). A study by
Gat et al. (2011), on stimulation of ureolytic MICP in natural soils, reported that
interaction between ureolytic and non-ureolytic bacteria was affected during ureolysis.
Their finding showed an increase in DIC concentration when ureolytic and non-
ureolytic bacteria co-cultured. This result was supported by a recent study by Gat et al.
(2014) on calcite precipitates using co-culture of ureolytic and non-ureolytic bacteria,
namely, Sporosarcina pasteurii, DSMZ33 and Bacillus subtilis, DSMZ 6397. Their
experiment showed that DIC concentrations were affected by three processes: (1)
hydrolysis of urea to produce bicarbonate, (2) bacterial respiration and mineralization of
the NB by ureolytic and non-ureolytic bacteria to produce dissolved CO2, and (3)
precipitation of CaCO3, which led to a reduction in DIC concentration (Engel et al.,
2004). The decrease in dissolved calcium concentration observed in this experiment
may be attributed to the precipitation of CaCO3. A study by Tobler et al. (2011)
reported a similar phenomenon for the induction of urea hydrolysis in a mixed culture
of indigenous soil bacteria.
1.4.5. Bacteria size
The type of bacteria appropriate for MICP application should be able to catalyst the
urea hydrolysis and they are usually urease positive bacteria (Soon et al., 2012). The
typical urease positive bacteria used for MICP are aerobic bacteria, are often selected
for MICP process because of their ability to release CO2 which is essential for the rise
in pH due to the production of ammonium when urea is being broken down (Soon,
2013). Bacterial sizes found in soil ranges from 0.5 to 3.0 μm microbes can move along
soil particles either through self-propelled manner or via passive diffusion (Mitchell and
Santamarina, 2005, Soon et al., 2012).
22
The geometric compatibility of urease producing bacteria is critical whenever the
transportation of bacteria within the soil is required for soil treatment, and mall pore
throat size would limit their free passage, depending on the size of microbes and soil
composition (Sarayu et al., 2014). A significant amount of silt and clay in the ground
would have an inhibitory effect on bacteria’s movement (Soon et al., 2014). It is
imperative to select appropriate soil and bacteria for MICP treatment (Soon, 2013).
1.4.6. Nutrients
Nutrients are the energy sources for bacteria, providing sufficient nutrient the ureolytic
bacteria is critical for precipitation of calcite (Soon et al., 2012). Nutrients are often
supplied to the bacteria during culture and soil treatment stages (Soon, 2013). The most
common nutrients usually provided to bacterial include Potassium, Sodium, Nitrogen,
Calcium, Iron and Magnesium (Mitchell and Santamarina, 2005). The unavailability of
organic constituents in soil limits bacterial growth, hence the supply of sufficient
nutrient to soil containing ureolytic bacteria can promote bacterial growth which can
enhance calcite precipitation required in achieving the desired level of ground
improvement (Soon et al., 2012).
1.4.7. Availability of nucleation site
A nucleation site is isolated from the environment by a restricting geometry limiting the
diffusion in and out of the system, which enable the modification of the activity of at
least a cation, proton, and other possible ions and ensure electro-neutrality (Sarayu et
al., 2014). The ion movement is enabled by active pumping with organelles or passive
diffusion to enable the microorganisms to use a great variety of anatomical
arrangements (Perry, 2003). The biofilm and the extracellular polysaccharide which is
formed by the microorganisms are effective in binding ions from the environment and
act as a heterogeneous nucleation site for the mineral deposition (Sarayu et al., 2014).
The creation of a strong electrostatic affinity to attract cations and enables the
accumulation of calcium ions on the surface of the cell wall which allows sufficient
supersaturation state of calcium ions to be achieved. Thus binding it to the carbonate
ions and results in the formation of calcium carbonate on the cell wall (Obst et al., 2009,
Tourney and Ngwenya, 2009). This mechanism favours the bacterial growth by
reducing the toxic calcium in the environment (Sarayu et al., 2014).
23
Higher bacterial cell concentration (106 to 108) supplied to soil samples would certainly
increase the amount of calcite precipitated from MICP process (Okwadha and Li, 2010).
Urea hydrolysis rate is directly proportional to a concentration of bacteria cell, provided
there will be enough reagent available for the biocement treatment of sand (Soon et al.,
2012). High concentration of bacteria produces more urease per unit volume to
commence the urea hydrolysis (Soon, 2013). Li et al. (2011b) and Stocks-Fischer et al.
(1999) suggested that the cells of the bacteria served as a nucleation site for MICP
occurrence.
The availability of nucleation sites serves as one of the key factors for microbial calcite
precipitation (Knorre and Krumbein, 2000). Lian et al. (2006) studied the crystallization
by Bacillus megaterium. They showed using scanning electron microscopic images that
nucleation of calcite takes place at bacteria cell walls. Stocks-Fischer et al. (1999) also
demonstrated that calcite precipitation relates with the bacteria concentration used.
Stocks-Fischer et al. (1999) were able to relate calcite induced via MICP efficiency with
chemically induced calcite at pH 9.0. Their findings concluded that about 98% of the
initial concentrations of Ca2+ were precipitated via MICP. On the other hand, only 35 to
54% of chemically induced calcite was observed. It was then suggested that the
bacterial cells provided a nucleation site for calcite to be induced which increased the
environment for further calcite to be induced, was responsible for the differences in
calcite precipitated via MICP and chemical processes.
1.5 Current Biotechnological Application of MICP MICP is highly desirable because of its natural availability and lower production of
pollutants (Al-Thawadi, 2008). MICP process is an effective and environmentally
friendly technology which can be applied to solve various environmental problems such
as soil instability and concrete crack (Anbu et al., 2016). Some of the biological
applications of MICP have been discussed by Whiffin (2004), Al-Thawadi (2008) and
in review articles by Phillips et al. (2013), Sarayu et al. (2014) and Anbu et al. (2016).
24
1.5.1. Biocementation
Biocement or biosandstone was proposed as a novel method for cementing loose sands
to produce structural materials, consisting of Alkaliphilic urease producing bacteria, a
substrate solution (urea), a calcium source and sand (Achal, 2015). However, a typical
set-up for sand consolidation experiment to develop biocementation was simplified by
Reddy et al. (2012), where sand is either mixed with bacterial culture or later injected
directly into the sand columns. The sand was plugged through a plastic column, and the
cementation fluid which consisted of nutrient media, urea, and calcium ions were then
injected at a specific rate in the column using gravimetric free flow direction. Another
study on calcite deposition in sand columns using Sporosarcina pasteurii by Achal et al.
(2009b) found that 40% of calcite deposited in the sandstone resulted and led to a
reduction of porosity and permeability in the sandstone. A study by Qian et al. (2010)
on a sand column of a size of 32.10 and 18.40 mm showed the right amount of
compressive strength, measured up to 2 MPa when CaCl2 was used as a calcium source
for biosandstone. The MICP substance in the biosandstone was confirmed using X-ray
diffraction (XRD) and energy dispersion spectroscopy (EDS), and calcite, which was
precipitated in the sandstone as the main microbial induced substance in the
biosandstone. The results of MICP process on biosandstone lead researchers to carry out
investigation beyond this building material (Achal, 2015).
1.5.2. Creation of biological mortars The knowledge obtained with MICP treatments resulted in the development of
biological mortar for remediation of small cavities on limestone surfaces (De Muynck et
al., 2010a). The purpose of using initiating biological mortars was to avoid some of the
problems related to chemical and physical incompatibilities of commonly used mortars
with the underlying materials, specifically in the case of brittle materials (Castanier et
al., 1999). The resistance of mortar specimens and surface deposition to degradation
process can be improved via microbial calcite precipitation (Siddique and Chahal, 2011,
Al-Thawadi, 2011, Chunxiang et al., 2009).
25
Figure 1.5: An in situ application of bacteria based liquid. Ureolytic bacterial culture was used to repair a system on cracked parking decks (Jonkers et al., 2016). A study by Le Metayer-Levrel et al. (1999) showed that they successfully studied
bacterial cementation which aimed at the creation of biological mortars and patinas on
limestones. Their method solely depended on spraying the entire surface of limestone
with bacteria followed by nutritional medium containing urea and calcium. Rodriguez-
Navarro et al. (2003) reported a relatively low penetration depth of 500 μm by
immersing the limestone sample in cementation media. They reported the use of
Myxocccus xanthus (a slow growing bacterium) resulted in CaCO3 precipitation at the
wall of the porous materials without plugging them. A recent in situ application on
cracked was carried out by Jonkers et al. (2016) as shown in Figure 1.5. Their finding
showed that concrete repair using MICP is inexpensive, improved the durability of the
material and also lowered the environmental impact of civil engineering activities.
1.5.3. Bioremediation of cracks in concrete In concrete, cracking is common due to relatively low tensile strength (De-Belie and
De-Muynck, 2008). Several mechanisms such as shrinkage, freeze-thaw reactions,
mechanical compressive and tensile forces lead to the formation of cracks (Alhour,
2013).
26
Cracking on concrete surfaces also results in enhanced deterioration of embedded steel
through easy ingress of moisture and ions that react with reinforcements in concrete and
expansive stressed which leadings to spalling (Gavimath et al., 2012, Achal et al.,
2013). Thus, it is practical to use adhesive for sealing of concrete cracks so that the
strength and durability of the concrete will be improved (Wong 2015). A conventional
approach used in repairing cracks involves injecting epoxy resin or cement grout into
the concrete. However, they result in various thermal expansion, environmental and
health hazards (De-Belie and De-Muynck, 2008).
Figure 1.6: Self-healing crack from the addition of bacterial metabolism via urea hydrolysis. The ureolytic bacterial culture was able to produce minerals which helped to repair and cover the cracks (Sierra-Beltran et al., 2014). Several research groups have investigated the possibility of using MICP as an
alternative effective repair method for cracks in concrete via bioremediation (Alhour,
2013). Investigation on the potential of using bacteria to act as self-healing agent in
concrete to fix a crack. Specifically, with the use of alkali-resistant spore-forming
bacteria, Bacillus pseudofirmus (type strain DSM 8715) and Bacillus cohnii (type strain
DSM 6307) (Jonkers, 2007, Jonkers and Schlangen, 2007, Jonkers et al., 2010).
27
Their findings showed that bacterial cement stone specimens appeared to produce a
solid result of crack-plugging. Other studies by Abo-El-Enein et al. (2013), Bang et al.
(2010), and Siddique and Chahal (2011) have shown that the cracks in concrete filled
with a mixture of Sporosarcina pasteurii and sand showed a significant increase in
compressive strength and stiffness when compared to cracks without cells. In Figure
1.6, Sierra-Beltran et al. (2014) reported self-healed cracks using MICP.
1.5.4. Biodeposition on cementitious materials
The emergence of microbial involvement in carbonate precipitation has led to the
exploration of this process in a variety of fields, including environmental, civil and
geotechnical engineering (De Muynck et al., 2010a). Among these applications, MICP
has been used for biogenic-carbonate-based surface treatments, a process known as
biodeposition (Figure 1.7) (Le Metayer-Levrel et al., 1999, Rodriguez-Navarro et al.,
2003, Dick et al., 2006).
Figure 1.7: 1 mm thick calcite crust formed on the surface of the soil. A successful percolation treatment with ureolytic bacterial culture, a high concentration of urea and calcium solution resulted in a nearly impermeable crust on the surface of the sample (Achal et al., 2010c).
28
Biodeposition of bacterial calcite is a viable method of surface treatment for cement-
based materials that can be explored in a sustainable approach (Wong, 2015).
Considering the size of bacterial cells are around 1 µm, both the cells and their media
containing the reactants (urea and calcium ions) can permeate deep into the pores and
interface between aggregates or paste of the concrete structure (Ramachandran et al.,
2001). Hence, this enables microbial cementation to take place within and on the
surface of such materials which then provides reinforcement and protection (Wong,
2015).
A study by De-Muynck et al. (2011) using ureolytic biodeposition treatment was
applied to five types of limestones so as to investigate the effect of pore structure on the
protective performance of bigenis carbonate surface treatment. Their findings showed
that in macroporous stone, biogenic carbonate formation occurred to a larger extent and
at greater depths than in microporous stone. Hence, exhibiting a greater protective
performance on macroporous stone compared to microporous stones. Precipitation on
microporous stones was limited to the outer surface of a microporous rock. From this
study, it was clear that biodeposition was very effective and more feasible for
macroporous stones than for microporous stones (De-Muynck et al., 2011). Another
study by Li and Jin (2012) on remediation technique of cracked concrete by bacterially
mediated carbonate deposition showed that bio-deposition was able to make
improvement in concrete compressive strength and flexural load using Sporosarcina
pasteurii. Their findings concluded that this can be used to enhance the strength and
flexural load of a faulty concrete specimen.
1.5.5. Biogrout
Nemati and Voordouw (2003) described the use of urease to cement porous medium.
Their study showed that reducing the permeability of porous medium by enzymatic
CaCO3 precipitation using Canvalia ensiformis was successful. Nemati and Voordouw
(2003) used between 0.1 and 1.0 M (>33 g.L-1) calcite together with high urease activity
for a successful plugging of the sand core. Unfortunately, the strength build-up was not
monitored. Stocks-Fischer et al. (1999) reported that injection of bacteria and reagents
together at low flow rates can result in full clogging of the system near the injection
point. An investigation on Biogrout ground improvement using MICP was also
performed by Paassen (2009).
29
This study was successful in developing an unprecedented 100 m3 field scale
experiment (Figure 1.8), and 40 m3 of the sand were treated using MICP process within
a duration of 12 days Although in both scale up experiments significant increase of the
average strength was obtained, different variable mechanical properties were observed
in the sand. It could be affected by induced flow field, bacteria distribution, the supply
of reagents and crystallization process (Paassen, 2009). Another study by Suer et al.
(2009) investigated the potential of using biogrouting as an alternative approach to jet
grouting to seal the contact between sheet pilling and bedrock. Their finding showed
that biogrouting process was cheaper than jet grouting and had much lower
environmental impact. Biogrouting also consumed less water and produced less
landfilled waste.
Figure 1.8: Set-up for large scale (100 m3) soil treatment. The sand was injected 10 times for 12 days with Sporosarcina pasteurii cell and cementation solution (Paassen, 2009). The scale-up demonstration of MICP in 100 m3 of sand to determine the ground improvement abilities and extent of precipitation (Phillips et al., 2013).
30
1.5.6. Other essential applications of MICP 1.5.6 (a): Removal of calcium ions (Ca2+)
Calcium-rich wastewater is a problem some industries face due to calcification during
downstream processing (Hammes et al., 2003c). High concentration of calcium ions
ranging from 500-1500 mg.L-1 in the wastewater can cause substantial scaling in
pipelines and reactors as a result of calcium formation as carbonate, phosphate, and
gypsum (Al-Thawadi, 2008, Dhami et al., 2013e). A novel application for the process of
MICP as an alternative mechanism for the potential removal of Ca2+ from industrial
wastewater instead of chemical precipitation approach has been developed (Hammes et
al., 2001). MICP process facilitated the removal of soluble calcium from calcium-rich
industrial wastewater via urea hydrolysis pathway, mediated by autochthonous bacteria.
Calcium removal more than 90% was achieved throughout the experimental period
while the effluent pH remained at a reasonable level (Hammes et al., 2001, Hammes et
al., 2003c). A recent study by Isik et al. (2010) showed that a significant parameter,
hydraulic retention time, required an optimum condition of 5-6 hr to hydrolyse calcium
successfully from industrial water using MICP in a biocatalytic calcification reactor.
1.5.6 (b): Removal of polychlorinated biphenyls (PBs)
Polychlorinated biphenyls (PCBs) is a recalcitrant contaminant which surfaces on
concrete when PCBs containing oils leaks from the equipment. (Phillips et al., 2013).
The last two decades have seen an increase in the use of bioremediation for the removal
of contaminants, which includes PCBs (Dhami et al., 2013e). The conventional method
previously used to remove PCBs such as solvent washing, hydro-blasting and epoxy
coating have not been very effective due to resurfacing of the oil over a period of time.
Microbial process using MICP process has been initiated as an alternative measure to
remove PCBs (Dhami et al., 2013e). Okwadha and Li (2010) reported the potential use
of Sporosarcina pasteurii for the treatment of PCB-coated cement cylinders leading to
surficial encapsulation of PCB-containing oils. A study by Okwadha and Li (2011)
stated that when Sporosarcina pasteurii containing urea and calcium ions were applied
on the surficial PCB-containing oil, there was no observation of leaching and there was
a reduction of permeability by 1-5 orders of magnitude.
31
1.5.6 (c): Industrial by-products
Construction materials such as concrete, brick and pavement blocks are all produced
from natural existing resources. Their production has affected our environment due to
continuous exploration limitation of natural resources. It has led researchers to explore
other means of building materials which are environmentally friendly, affordable and
sustainable (Aubert et al., 2006). There are different types of waste such as slag, fly ash,
wheat straw, saw milk waste, cotton stalk, mining waste tailing and waste gypsum
which are currently being recycled for potential utilisation (Pappu et al., 2007). The
production of fly ashes during combustion of coal for energy is one of the industrial by-
product recognised as an environmental pollutant (Dhami et al., 2013e). Rice husk ash
obtained from burning of rice husk is another major agricultural by-product (Dhami et
al., 2013e). Both these materials can be used as construction materials (bricks and
blocks) without any degradation in the quality of products (Nasly and Yassin, 2009).
Despite the previous report of the problems associated with ash bricks such as low
strength, high water adsorption and low resistance to abrasion. Dhami et al. (2012b)
studied the application of bacterial calcite on fly ash and rice rush ash bricks and
reported they were very efficient in reducing permeability and decreasing water
absorption which lead to enhanced durability of ash bricks.
1.5.6 (d): Low energy building materials
The construction sector is responsible for primary input of energy resulting in the
release of CO2 emissions into the atmosphere (Reddy and Jagadish, 2003). Hence, it is
essential to reduce the emission of these gases released into the air (Dhami et al.,
2013e). Energy requirements for production and processing of different building
materials and various implications on the environment have been previously studies
(Oka et al., 1993, Debnath et al., 1995, Suzuki et al., 1995). Reddy and Jagadish (2003)
reported soil blocks with 6–8% cement content uses the moving energy efficient
building material. These materials have low production cost, are easily recyclable and
environmentally friendly as the soils are mixed with additives such as lime (Dhami et
al., 2013e).
32
These building materials do not make use of burning during its production, and these
stable mud blocks were able to converse much energy (Dhami et al., 2013e). Building
materials using low energy by application of ureolytic Bacillus sp. have successfully
been performed by Dhami et al. (2013c) which shows the potential of using MICP
technology to produce sustainable, cheap and durable buildings.
1.6 Diversities of Microbial Communities in Caves Caves are natural geological formations considered as extreme environments,
unfavourable for the development of life due to the severe abiotic conditions present
(Tomczyk-Żak and Zielenkiewicz, 2015). However, cave environments constitute
ecological niches for highly specialised microorganisms (Schabereiter-Gurtner et al.,
2004). The most common types of caves known are karst caves, formed from limestone
rocks and cave created as a result of lava cavities (Tomczyk-Żak and Zielenkiewicz,
2015). Caves constitute oligotrophic ecosystems, which are less than 2 mg of the total
organic carbon per litre. These environments have a low level of light, low, stable
temperature and high humidity (Tomczyk-Żak and Zielenkiewicz, 2015). Despite these
oligotrophic conditions, the average number of microorganisms dwelling in these
ecosystems are 106 cells/g of rock (Barton and Jurado, 2007).
The majority of biological communities are dependent on energy and carbon fixation of
photosynthesis. However, the inhibition of sunlight prevents colonisation of
phototrophs in cave environments (Wu et al., 2015). Only limited energy and nutrients
can enter these caves through sinkholes, underground hydrology and drip water (Barton
et al., 2007). These environments only allow for the survival and functioning of species
adapted to oligotrophic conditions (Wu et al., 2015). The limited access of
photosynthetic activities in caves inhibits the production of primary organic matter
essential for the survival of photosynthetic microorganisms. Hence, these cave
microorganisms make use of alternative methods by synthesising their organic
molecules through carbon dioxide fixation to produce their source of food or energy
(Tomczyk-Żak and Zielenkiewicz, 2015).
33
This condition allows these cave microorganisms to derive their main source of energy
from not only hydrogen, nitrogen or volatile compounds, but they also derive their
energy from the oxidation of inorganic molecules such as iron, sulphur or magnesium
present in caves (Gadd, 2010, Northup and Lavoie, 2001). Other sources of organic
compounds from which these microorganisms derive their energy comes from plant
roots or remains of human or animal activities, these organic matter allows the
developments of a heterotrophic microorganism (Tomczyk-Żak and Zielenkiewicz,
2015).
Studies performed on calcified structures (Figure 1.9) are of biogenic origins, their
study showed that microorganisms interacted with minerals, hence playing an important
role in the formation of these calcified structures (Melim et al., 2009). These
interactions help in shaping cave structures such as stalactites, stalagmites, as well
formations of bristles in surfaces of cave rocks (Tomczyk-Żak and Zielenkiewicz,
2015). Some of these cave microorganism precipitates CaCO3 on the surfaces of their
cells, which contributes to formations of limestones in the caves (Sanchez-Moral et al.,
2003). The occurrence and structure of microbial communities in limestone caves are
influenced by factors such as pH, availability of nutrients, sunlight, oxygen, metal
compounds, humidity and susceptibility of the substrate to colonisation (Tomczyk-Żak
and Zielenkiewicz, 2015). Bacteria and archaea constitute a majority of the biodiversity
in caves, found in numerous cave habitats such as sediments, stream waters and rock
surfaces (Barton and Jurado, 2007, Engel et al., 2004).
Chemoautotrophic microbes are mostly responsible for CO2 fixation and potentially
participate in inorganic nitrogen (Tetu et al., 2013, Diaz-Herraiz et al., 2013). Moreover,
the interactions between microorganisms and limestone caves may contribute to
speleogenesis, for example, in sulfidic caves, microorganisms can oxidise of hydrogen
sulphide to produce sulfuric acid, which then reacts with carbonate and causes rock
dissolution (Macalady et al., 2007, Engel et al., 2004). Bacteria can alter the surfaces of
rocks through oxidation of some metal elements such as iron (Fe2+) and manganese
(Mn2+) which result in the formation of deposits on cave walls (Carmichael et al., 2013).
34
Figure 1.9: Calcified structures of biogenic origin discovered in cave regions. (A) pool fingers formation and (B) U-loops formation (Garcia et al., 2016). Cave microbial communities are often extremely variable depending on the
microhabitats (Wu et al., 2015). A study by Barton et al. (2007) showed there were
significant differences between the diversity of bacteria and its composition observed on
rock walls within one single cave, suggesting this was possibly related to the host rock
geochemistry. An alteration of physiochemical conditions can influence a change of in
the composition of microbial species. For example, in the water mats of streams in the
Kane Cave, which is rich in sulfur compounds, the water flowing directly from the
spring into the cave, contains a high concentration of sulfur and low amounts of oxygen,
dominated by Epsilonproteobacteria. On the other hand, the water flowing out of the
cave to external environments contains large quantities of oxygen and low
concentrations of sulfur, dominated by Gammaproteobacteria (Tomczyk-Żak and
Zielenkiewicz, 2015, Engel, 2010, Jones and Bennett, 2014).
Studies Rusznyak et al. (2012) on the effect of the microbial population in Herrenberg
Cave in Germany, a typical karst cave, showed that the occasional or limited human
presence in cave environments does not necessarily affect the compositions of microbial
diversity of a population. However, a study by Adetutu et al. (2012) indicated that the
presence of human activity in regions of Naracoorte Caves in Australia had a consistent
influence in bacterial diversity which was attributed to the presence of exogenous
organic matter of human origin. Various studies have demonstrated that bacteria from
35
cave environments are capable of inducing calcite precipitates in vitro (Garcia et al.,
2016). Different species and genera of bacteria have been isolated from speleothems
samples in caves which include Sporosarcina pasteurii, Bacillus subtilis, Myxococcus
xanthus, Bacillus amyloliquefaciens, Bacillus cereus, Pseudomonas flurescens,
Micrococcus sp., Rhodocucus sp. and Arthrobacter sp. (Rusznyak et al., 2012, Achal et
al., 2010b, Rivadeneyra et al., 2006).
Figure 1.10: Speleothems samples collected from El Toro and El Zancudo limestone mines located in Cordillera Central, northeast of Colombia. The diversity of bacteria from speleothems samples in Colombia and their ability to precipitate carbonates were studied using conventional microbiological methods and molecular tools, such as temporal temperature gradient electrophoresis (Garcia et al., 2016). In addition, Rusznyak et al. (2012) and Cacchio et al. (2004) have suggested that the
microorganism mentioned above have a direct relationship with calcite depositions and
speleothems developments in limestone caves. Speleothem carbonates formation were
normally considered as inorganic precipitates, but recent studies have demonstrated
biological influence in their formations (Baskar et al., 2007). These discoveries can
advance our understanding of the diversity of bacteria in cave environments (Roesch et
al., 2007).
36
Most researchers regarding the profiles of microbial communities in speleothem
samples (Figure 1.10) make use of culture-dependent study based on partial analysis of
the 16S rRNA gene using clone library methods and genetic fingerprinting techniques
such as denaturing gradient gel electrophoresis (DGGE). Their studies suggested that
the most dominant phyla in cave environments are Firmicutes, Proteobacteria,
Actinobacteria and Acidobacteria (Ortiz et al., 2014, Dhami et al., 2014, Ortiz et al.,
2013). A metagenomics approach on the study of microorganisms in karstic cave Ortiz
et al. (2014) suggested that functional bacterial genes were associated with low nutrient,
high calcium adaptations, and nitrogen-based metabolism.
1.7 Screening Sarawak’s Limestone Caves for Ureolytic Bacteria Sarawak is one of the two Eastern Malaysian states situated on the island of Borneo,
known as the world's third largest island and one of the twelve mega-biodiversity
regions (Lateef et al., 2014, Tan et al., 2009). Borneo has a landmass of nearly 740,000
square kilometres, located in the equatorial region of the Pacific Ocean (Rautner et al.,
2005). The Island consist of the independent Sultanate of Brunei Darussalam, the
Indonesian territory of Kalimantan, and the Malaysian states of Sarawak and Sabah
(Rautner et al., 2005, Sulaiman and Mayden, 2012) as shown in Figure 1.10.
Borneo is widely known for its rich floral and faunal diversity. However, many areas of
the island require further exploration (Clements et al., 2010, Garbutt and Prudente,
2007, Mohd et al., 2003, Koh et al., 2010, Karim et al., 2004). Diverse habitats such as
mangrove swamps, peat swamps, an estimated 15, 000 plant species (5, 000 trees, 17,
000 orchid species and over 50 carnivorous pitcher plants) host a great diversity of
endophytic microorganisms in Borneo (State Planning Unit, 2013). In 2007, the
countries situated in Borneo Island made a declaration to protect 220,000 square
kilometres of pristine rainforest habitats which are now known as the “Heart of
Borneo,” to prevent disturbances such as deforestation and plantation development from
affecting the Island’s biodiversity (Sulaiman and Mayden, 2012).
37
Sarawak is the largest state in Malaysia, containing 37.5% of the country’s total land
(Mahidi, 2015). Also, Sarawak has 512, 387.47 hectares of protected areas comprising
of 18 National Parks, four wildlife sanctuaries, five nature reserves and the largest
peatland area in Malaysia (Van der Meer et al., 2013, Forest Department Sarawak,
2013). The rich mega-biodiversity in Sarawak has attracted the attention of researchers
within and outside of Malaysia. The existing scientific studies have focused on peat
soils, plants, corals, microbes in aquatic and forest environments (Sa'don et al., 2015,
Kuek et al., 2015, Cole et al., 2015). Sarawak’s limestone forest is one of the nine main
types of forest documented in Sarawak, covering about 520 m2 or 0.4% of the total area
(Julaihi, 2004, Banda et al., 2004).
The limestone forest is situated with vast numbers of limestone caves. The caves or
limestone areas in Sarawak have become the main focus for researchers to investigate
the diversity of bats indigenous to Wind and Niah caves (Mohd et al., 2011, Rahman et
al., 2010b, Rahman et al., 2010a). Studies on evolution of limestone formation,
biological influence on formation of stalagmite, investigation of trace metal ratios and
carbon isotopic composition on limestone caves have been carried out in Niah and Mulu
caves, which are also situated in Sarawak (Moseley et al., 2013, Dodge-Wan and Mi,
2013, Cucchi et al., 2009). Despite Malaysia’s abundance of limestone regions situated
in places such as Langkawi Island, Kedah-Perlis, Kinta Valley, Perak, Selangor, Gua
Musang, and Kelantan as reported by Bakhshipouri et al. (2009).
There are limited reported studies on the exploitation of microbial diversity from these
regions. Moreover, there have been recent studies on isolation of calcite forming
bacteria from limestone cave samples of Perak and research on soil improvement using
Bacillus megaterium, ATCC 14581 type strain (Soon et al., 2014, Soon et al., 2013,
Komala and Khun, 2013). To date, there have been no recorded studies in Sarawak on
the isolation of urease producing bacteria from limestone caves samples of Sarawak.
This research gap and the possibility of certain microbes able to induce calcite
precipitates from limestone cave environments initiated the relevance of screening for
urease producing bacteria from two cave regions in Sarawak.
38
Figure 1.11: Map of Borneo Island. showing the geographical divisions and topographical features of Brunei Darussalam, Indonesia (Kalimantan) and East Malaysia (Sarawak and Sabah). The island of Borneo, known as the world's third largest island and one of the twelve mega-biodiversity regions (Lateef et al., 2014, Tan et al., 2009, Tan, 2006).
39
1.8 Aim and Objectives of the Study The aim of this research was to screen and characterise urease-producing bacteria that
are capable of inducing calcite precipitation. The objectives set out to achieve the
research aim are:
i. To isolate urease producing bacteria from limestone cave samples of Sarawak
using enrichment culture technique.
ii. To identify urease-producing bacteria.
iii. To characterise urease activity of bacterial isolates.
iv. To determine the effects of cultural conditions on urease activity.
v. To study biocementation ability of selected bacteria in vitro.
1.9 Significance of the Study This study explores the prospect of using urease-producing bacteria which were isolated
from domestic location rich in microbial diversity for possible biocementation
applications. The advantage of using local isolates is because they are well adapted to
native environments, and they are also less likely to become pathogenic when they are
under stressed conditions. Additionally, studies on the isolation of non-pathogenic
highly active urease-producing bacteria species are very limited. This formed the
necessary initiation of this study which could pave the way for a new frontier in the use
of non-pathogenic bacterial species isolated from Sarawak, Malaysia.
1.10 Thesis Outline This thesis presented is divided into four chapters: Introduction and Literature Review
(Chapter 1). Isolation, Identification, and Characterisation of Urease-Producing Bacteria
from Limestone Caves of Sarawak (Chapter 2). Effects of Cultural Conditions On
Urease Activity, and Evaluation of Biocementation Potentials in Small Scale Test
(Chapter 3). General Conclusion and Recommendations (Chapter 4). Concluding
remarks are shown at the end of Chapter 2 and 3 to summarise the contents of theses
chapters.
40
Chapter 1 provides a brief introductory background of the study and a broad review of
the essential literature regarding MICP, which has been reported by other researchers.
The aim, scope, and significance of the research which was to be performed, were also
conferred in this chapter. Chapter 2, presents a detailed study on the isolation, screening
and identification of the ureolytic bacteria which were obtained using enrichment
culture technique. In this chapter, specific focus was given to the quantitative
measurement of specific urease activity by the local isolates. The enzyme activity of
these isolates was compared with that of the representative strain used in this study. The
isolates capable of producing comparable urease activities with that of the
representative strain were selected and used for subsequent experiments. Chapter 3,
presents the results on the effects of cultural conditions on the urease activity. A
laboratory-scale study concerning the application of ureolytic bacteria for MICP process
to treat poorly graded soil. The sole purpose of this chapter was to access whether
sufficient potential exists to warrant the possible usage of the locally isolated ureolytic
bacteria, serving as alternative MICP agents. This knowledge can lead to further
investigation along this line of research such as large-scale microbial production in a
reactor and field applications using MICP agents. In Chapter 4, a succinct overview of
the most significant findings of the experimental studies is presented and are shown
within the context of one another. Perspectives on future research possibilities within
this field are conferred in this chapter as future directions to be considered.
Chapter
2 ISOLATION, IDENTIFICATION AND
CHARACTERISATION OF UREASE-PRODUCING
BACTERIA FROM LIMESTONE CAVES OF SARAWAK
41
2.1 Introduction Limestone caves, known as natural geological formations are considered as extreme
environments which form an ecological niche for the survival of various
microorganisms (Schabereiter-Gurtner et al., 2004). These environments often excluded
from the outside world with limited in nutrient, may contain novel, diverse microbial
populations (Sugita et al., 2005). Hence, it is imperative to perform pioneering
investigations ventured at exploring and isolating microbial species that indigenous to
cave regions. It’s been known and reported that formation of stalagmite and stalactites
are often as a result of microbial and mineral interactions (Tomczyk-Żak and
Zielenkiewicz, 2015). Some microorganisms are able to induce calcites on the surface
of their respective cells, which promotes limestone formation (Schabereiter-Gurtner et
al., 2004). This chapter reports the investigation of bacterial microorganisms isolated
from limestone caves of Sarawak with potential industrial relevance. These
microorganisms, ureolytic bacteria, prefer to live in alkaline environments, produce an
enzyme which primarily allows calcite precipitation to occur (Achal and Pan, 2011).
This process, microbial-induced calcite precipitation (MICP) is usually directed by
urease enzyme (urea amidohydrolase; EC 3.5.1.5) which is produced by some
microorganisms that relies on urea as their primary source of nitrogen (Zhang et al.,
2015, Achal, 2015).
Urease enzyme was previously studied from clinical evaluation on patients infected
with pathogenic microorganisms (Cheng and Cord-Ruwisch, 2013, Lee and Calhoun,
1997, Mobley et al., 1995). However, the usage of urease on biocementation application
for improvement of soil strengthening has been the subject of various research from the
Microbial biotechnology, geotechnical engineering and civil engineering (Al-Thawadi,
2008, DeJong et al., 2006, Whiffin, 2004). Studies on the alternative source for known
UPB from non-pathogenic bacterial species necessary for urea hydrolysis in
biocementation application are very limited. This research gap forms the basis for, the
initiation of this study. This is the first study elucidating the isolation and identification
of ureolytic bacteria from limestone caves of Sarawak.
42
The objectives of the study in this chapter are as follows:
i. To screen and identify urease-producing bacteria.
ii. To characterise the urease activities of selected isolates.
iii. To test the ability of selected isolates to induce calcium carbonate precipitation
and bacterial growth and pH profiles.
43
2.2 Methods and materials
2.2.1. Sampling location and collection
A field sampling occurred at Fairy Cave Nature Reserve (FCNR) and Wind Cave
Nature Reserve (WCNR) and samples used in this research study were taken from this
sampling site. These caves, Fairy cave (N 01°22’53.39” E 110°07’02.70”) and Wind
cave (N 01°24’54.20” E 110°08’06.94”) are located in Bau, Kuching Division,
Sarawak, East Malaysia, on the island of Borneo. Samples taken from FCNR were
collected at depth of 5-10 cm from regions surrounded by rocks and vegetation, while
samples taken from WCNR were collected from the surfaces of speleothems inside the
cave chamber. Each sample was collected using sterile tools, placed in sterile
polystyrene containers, sealed and stored in an ice box (at the sampling site) before
being transported to Swinburne University of Technology, Sarawak campus for further
microbiological analysis. The samples were then preserved in the refrigerator (4°C)
prior to enrichment culturing.
2.2.2. Biological material
Sporosarcina pasteurii, (DSM33) type strain was purchased from the Leibniz Institute
DSMZ-German Collection of Microorganisms and Cell Cultures (Braunschweig,
Germany). This bacterial strain was used as a positive control for subsequent
experiments in this study. It was aseptically grown under aerobic batch conditions
according to the DSMZ instruction and stored on Petri plates containing nutrient agar
(HiMedia, Laboratories Pvt. Ltd) at 4oC in the fridge until when usage was necessary.
2.2.3. Growth medium and sterilisation
Nutrient broth (HiMedia) and nutrient agar (HiMedia) were used as a primary growth
medium in this study. All bacterial growth mediums, chemicals (except urea) and
glassware used in this study were sterilised by autoclaving at 121oC, 103.42 kPa for 20
min using an autoclave machine (Hirayama-HVE-50). Urea was sterile filtered through
a 0.45 µm syringe filter.
44
2.2.4. Enrichment cultures
Enrichment of the cave samples was performed as follows: 1 g or 1 mL of each sample
was inoculated into 50 mL growth media (250 mL shaking flasks). The enriched
cultures were placed in an incubation shaker (CERTOMAT® CT plus – Sartorius) under
aerobic batch conditions at 30oC for 120 hr at 130 rpm. The following growth media
were used to enrich the collected cave samples: Nutrient broth (13.0 g.L-1, HiMedia
Laboratories Pvt. Ltd)); Tryptic soy broth (30.0 g.L-1, Merck Millipore); Lactose
peptone broth (35.0 g.L-1, Becton, Dickinson and Company); Luria broth (20.0 g.L-1,
HiMedia Laboratories Pvt. Ltd) and Brain heart infusion broth (37.0 g.L-1, Oxoid
Thermo Scientific Microbiology). Each of the growth culture mediums was
supplemented with C2H3NaO2 (8.2 g.L-1, HiMedia Laboratories Pvt. Ltd), (NH4)2SO4
(10.0 g.L-1, HiMedia Laboratories Pvt. Ltd) and CH4N20 (20.0 g.L-1, Bendosen
Laboratory Chemicals). The initial pH of all media was adjusted to 8.0 using 0.1 M
NaOH or 0.1 M HCL before sterilisation (Reyes et al., 2009). Sterile Urea substrate (by
0.45 µm filter sterilisation) was added post-autoclaving to prevent chemical
decomposition under autoclave condition.
2.2.5. Isolation of urea degrading bacteria
For bacterial isolation, 1 mL of individual enriched culture samples was serially diluted
(sixfold) and plated on nutrient agar (with 6% urea). 0.1 mL aliquot of serially diluted
enrichment samples were inoculated onto Petri plates containing nutrient agar were then
spread using a sterilised L-shaped spreader until the fluid was evenly distributed. The
agar petri plates were then incubated (MMM Incucell ) under aerobic conditions at 32oC
for 42 hr. Upon the growth of isolates capable of hydrolysing 6% urea in petri plates
containing nutrient agar, subsequent sub-culturing was performed until single bacterial
colonies were obtained. Long term storage using glycerol stock was used in this study for
maintenance and preservation of the isolated bacterial isolates. Glycerol stock method
was used for long-term storage of the bacterial isolates by adopting a modified procedure
from Fortier and Moineau (2009).
45
For the maintenance of the bacterial glycerol stock, 500 µL of overnight grown cultures
were inoculated into 2.0 mL cryogenic vials containing sterilised 500 µL of 50%
glycerol to obtain a final glycerol concentration of 25% (v/v). The stocks were mixed
prudently and kept in the refrigerator at -80°C. For the case of reviving stored cells,
sterile toothpick or inoculation loop was used to scrap off the splinters of solid ice and
then onto the nutrient agar medium.
2.2.6. Screening for urease-producing bacteria
Christensen’s medium (Oxoid Thermo Scientific Microbiology Sdn Bhd) also called
urea agar base (UAB) was used to screen for urease producing bacteria (UPB). The
media components contained the following: Peptone (1.0 g.L-1); Glucose (1.0 g.L-1);
Sodium chloride (5.0 g.L-1); Disodium phosphate (1.2 g.L-1); Potassium dihydrogen
phosphate (0.8 g.L-1); Phenol red (0.012 g.L-1) and Agar (15.0 g.L-1). Urea solution, 4%,
(w/v) was separately prepared by filtration with the use of 0.45 µm syringe, and 10 mL
of the urea solution was aseptically introduced into 990 mL of the UAB medium. The
medium was carefully mixed by gentling swirling the Schott bottle containing the UAB
and 10 mL were then distributed into separate sterile test tubes. The bacterial isolates
were heavily inoculated on the surface of the UAB medium and then incubated at 37oC
for 72-120 hr. The urease production test was studied through visual observation for
colour changes. The bacterial isolate able to turn the UAB medium from pale yellow to
pink during the incubation period were selected while others were discarded.
2.2.7. Preliminary identification
Phenotypic analyses were used for a more definitive identification of bacterial isolates.
Morphological, microscopic and biochemical studies were performed under standard
protocols.
2.2.7 (a) Morphological analysis
A loopful of individual UPB cultures was serially subcultured onto Petri plates
containing nutrient agar and incubated for at 32°C for 24 hr. Colony appearance of the
overnight subcultured isolates were recorded with reference to Bergey’s Manual of
Determinative Bacteriology (Holt et al., 1994, Olufunke et al., 2014).
46
2.2.7 (b) Microscopic analysis
Gram staining and endospore staining methods were used to determine and differentiate
the cell morphology of the bacterial isolates. The standard staining protocol used to
differentiate between Gram positive and Gram negative bacteria was adapted from
Moyes, Reynolds and Breakwell (2005). The standard staining protocol used as a
differential stain to determine between the bacterial isolates capable of producing
endospores was adapted from Reynolds et al. (2009).
2.2.7 (c) Biochemical analysis
Motility, oxidase and catalase tests were performed and used for biochemical
characterization of the bacterial isolates. The procedures for these tests were adapted
from standard protocols by Shields and Cathcart (2011), Shields and Cathcart (2013),
and Bisen (2004).
2.2.8. Molecular identification
A Polymerase Chain Reaction (PCR) was used to amplify the 16S rRNA gene of the
unknown isolated urease producing bacteria. The DNA sequences of the 16S rRNA
genes were compared with the generated sequence to a database of a known sequence
which was then used to determine the molecular identification of unknown ureolytic
bacteria isolates.
2.2.8 (a) DNA extraction
A freeze and thaw method was used to lyse bacterial cell of an unknown
microorganism, to prepare Deoxyribonucleic acid (DNA) samples as templates for
DNA amplification. Colonies from 24 hr sub-cultured bacterial isolates were picked
using sterilised toothpicks. Each sample was then placed into individual wells of a
sterile 96 wells plate containing 100 µL Tris-EDTA (TE) buffer solution and then deep
frozen at -80°C for 24 hr as described by (Muramatsu et al., 2003). The 96 wells plate
was then thawed by immersing the plate in a 60°C water bath for 5 min to release DNA
from the microbial cells (Kuek et al., 2015). The lysate was used as a crude DNA
template for PCR.
47
2.2.8 (b) DNA amplification
The PCR technique comprising enzymatic amplification of nucleic acid sequence via
selected repeated denaturing, oligonucleotide annealing and DNA polymerase extension
cycles (Gibbs, 1990), was used in this study. DNA amplification was performed using
MyTaq Red Mix (Biolin) in accordance with the manufacturer’s instructions. PCR
amplification was performed using MyTaq Red Mix (Bioline) according to the
manufacturer’s instructions. The PCR master mix contained the following: template
(200 ng, 2 L); primers (1 L, 20 µm); MyTaq Red Mix (25 L) and sterile ddH2O (22
L). The forward primer, 8F: 5’-AGAGTTTGATCCTGGCTCAG-3’ (Hughes et al.,
2000) and reverse primer, 1525R: 5’-AAGGAGGTGATCCAGCC-3’ (Lane et al.,
1985) were used to amplify the 16S rRNA gene fragment. DNA amplification was
performed using a MasterCycler Gradient Thermal Cycler (Eppendorf 5331). The
cycles consisted of initial denaturation of the template DNA (95°C for 5 min),
denaturation (95°C for 60 sec), annealing (55°C for 60 sec), extension (72°C for 1 min
30 sec) and a final elongation (72°C for 7 min). The process was set to 30 cycles and
the system was held at 4°C.
2.2.8 (c) Visualisation of PCR products
Amplified DNA (PCR product) was visualised on 1% (w/v) agarose gel, stained with 1
L of Midori Green (Nippon Genetics Europe GmbH). The PCR product (5 L) was
loaded into the well of the 1% (w/v) agarose gel. MassRuler™ DNA Ladders (Thermo
Fisher Scientific) was used as a standard to determine the size of the target DNA. The
DNA was separated according to size by gel electrophoresis at 75 volts for 40 min. The
DNA bands were visualised with a gel doc XR system (Biorad) and the image was
captured.
2.2.8 (d) DNA purification and cycle sequencing
PCR purification and cycle sequencing of the products were carried out by First BASE
Laboratory Sdn. Bhd., Malaysia. DNA samples were purified using PCR Cleanup kit
(SS1012/3) with procedures following manufacturer’s instructions. The eluted solutions
(pure DNA) were then stored at -20°C. Sequencing was performed on an Applied
Biosystem 3130xl Genetic Analyzer, using BigDye® Terminator v3. Forward primer,
27F: 5'-AGAGTTTGATCMTGGCTCAG-3' (Heuer et al., 1997) was used while
1525R: 5’-AAGGAGGTGATCCAGCC-3’ was used as a reverse primer.
48
2.2.8 (e) Sequence analysis
The raw DNA chromatogram sequences were viewed using Chromas lite programme,
edited with BioEdit Programme (Hall, 1999) and stored in FASTA format. The forward
and reverse primer sequences were removed before the sequence were blasted with
existing sequences in national centre for biotechnological information (NCBI) GenBank
database (Zhang et al., 2000) using basic local alignment search tool (BLAST)
nucleotide collection database program (Ashelford et al., 2005) to search for closest best
match sequence (Tan et al., 2011). For investigation of the taxonomic composition of
the microbial strains, ribosomal database project (RDP-II) using the SeqMatch tool was
used to search the taxonomy database classification and nomenclature for all of the
organisms in the public sequence databases.
2.2.8 (f) Phylogenetic analysis
Molecular evolutionary genetic analysis (MEGA) version 6 was used to for
phylogenetic analysis (Tamura et al., 2013). Prior to phylogenetic analysis, indefinite
DNA sequences at both ends were removed and the gaps were adjusted to improve the
alignment (Zhao and Cui, 2013). Basic evolutionary distances from the MEGA
programme was used to analyse the DNA sequence (Saitou and Nei, 1987). Bootstrap
replicates (1000) were taken into account to infer the bootstrap consensus tree for the
representation of evolutionary history. The evolutionary distances were then processed
using the maximum composite likelihood method (Tamura et al., 2004, Hanif et al.,
2014).
2.2.8 (g) Nucleotide sequence accession numbers
The nucleotide sequences which were obtained in the present study have been deposited
in NCBI GenBank database (Kaverin et al., 2007). The provided GenBank accession
numbers for the submitted nucleotide sequences are KX212190 to KX212216.
2.2.9. Measurement of urease activity
The conductivity (mS.cm-1) method was used to determine the urease activity (mM urea
hydrolysed.min-1) in this study. For enzyme assay, 1.0 mL of overnight grown bacterial
cultures (0.6-1.0 OD) were inoculated into sterile individual universal bottles (20.0 mL)
containing 9.0 mL of 1.11 M urea solution (Harkes et al., 2010).
49
The changes in conductivity were monitored for a duration of 5 min at 25◦C ±1 and the
respective conductivity values were measured by immersing the probe of the
conductivity meter (Walk LAB conductivity pro meter, Trans Instruments COMPRO)
into the bacterial-urea solution. The conductivity variation rate (mS.cm-1.min-1) was
obtained from the gradient of the graph. The conductivity variation rate was then
multiplied by a dilution factor (df). The df was taken as the ratio of the stock bacteria
culture to the sampling bacteria culture before inoculating into the urea solution (Zhao
et al., 2014). These values were then used to calculate urease activity, by converting the
conductivity variation rate (mS.cm-1.min-1) to urea hydrolysis rate (mM urea
hydrolysed.min-1), based on the correlation that 1 mS.cm-1.min-1 corresponds to a
hydrolysis activity of 11 mM urea.min-1 in the measured range of activities (Paassen,
2009). The urea hydrolysis rate for the urease activity conversion was determined by
(Whiffin, 2004) as described in equation 1.23. Specific urease activity (mM urea
hydrolysed.min-1.OD-1) which reflects the urease catalytic abilities of the urea
hydrolysis (Zhao et al., 2014) was derived by dividing the urease activity (mM urea
hydrolysed.min-1) by the bacterial biomass (OD600). The specific urease activity was
also determined by (Whiffin, 2004) as described in equation 1.24. Biomass
concentration was determined by measuring the optical density of bacterial suspension
with a spectrophotometer (GENESYSTM 20, Thermo Fisher Scientific) at a wavelength
of 600 nm.
2.2.10. Evaluation of microbial calcite precipitation
2.2.10 (a) Testing calcite precipitation
A modified method of Hammes et al. (2003b) was adopted in this study and used to test
the ability of the local isolates to precipitate calcite. The Calcite precipitating media
(CPM) used in this study contains the following components: nutrient broth (3.0 g.L-1,
Oxoid); urea (20.0 g.L-1, Bendosen); NaHCO3 (2.12 g.L-1, Sigma); NH4Cl (10.0 g.L-1,
Sigma); CaCl2 · 2H2O (28.50 g.L-1, Sigma) and agar (20.0 g.L-1, HiMedia). For Calcite
precipitation screening, overnight grown bacterial broth culture were serially diluted
under the sterile condition and spread onto the CPM. The Petri dishes were then
incubated at 30°C for 6 days with the epidermal side facing upwards.
50
2.2.10 (b) Calcite estimation
A modified method of Wei et al. (2015) and Hammad et al. (2013b) were adapted for
this experiment. For a quantitative measurement of calcite precipitation in broth, the
nutrient broth was supplemented with urea 2% (w/v) and calcium chloride 2% (w/v)
solutions. The medium containing overnight grown bacterial cultures were incubated
under shaking condition (150 rpm) at 30°C for duration of 7 days. At the end of the
cultivation, the bacterial cultures were suspended through centrifugation (10,000 g for
60 sec) using centrifuge machine (Eppendorf, 5424R). The pellets which contained the
calcite precipitated and ureolytic bacteria culture were then resuspended centrifuge
tubes containing 50 mL TE buffer (10 mM Tris, 1 mM EDTA pH 8.5). Lysozyme (EC
3.2.1.17), also known as N-acetylmuramide glycanhydrolase was added to the
suspended samples, at a concentration of 1 mg.mL-1 (Wei et al., 2015). The samples
were then incubated at 37°C for 1 hr in order for the lysozyme to properly break down
the cell wall of the ureolytic bacteria. The samples were then centrifuged once more to
separate the cell debris form the calcite precipitates. The supernatants in the centrifuge
tubes were then discarded and dH2O (pH8.5) was added to the centrifuge tubes to wash
the pellets, which were then the air-dried at 37°C for 24 hr. The pellets obtained were
then weighted to estimate the amount of calcite precipitated (Walter et al., 2000).
2.2.11. Bacterial growth profile and pH profile
Ten millilitres (10 mL) of bacterial cultures were grown in universal bottles and
incubated for 24 hr at 32oC under shaking condition (150 rpm). Batch cultures were
prepared by transferring 2.5 mL of the overnight culture into 125 mL of sterile nutrient
broth medium (250 mL capacity conical flasks). The medium was then supplemented
with 6% sterile urea and the batch culture was grown for a total duration of 10 hr. Three
millilitres (3 mL) of the aliquot was sampled from the batch culture at every hour (1 hr)
and transferred into a 10 mm cuvette. A spectrophotometer (Genesys TM 20- Thermo
Scientific) was used to measure the optical density of the bacterial culture at 600 nm. A
pH meter (SevenEasyTM –Mettler Toledo) was also used to study the pH profile of the
bacterial culture by measuring the changes in pH during the incubation period.
51
2.2.12. Statistical analysis
The data were shown as mean ±SE (standard deviation) for three replicates. The results
were subjected to student’s t-test analysis, with statistical significance taken as p<0.05.
GraphPad (Quick Calc) programme was used to analyse the data.
52
2.3 Results
2.3.1. Sampling location and sample collection
Upon the authorization by Sarawak Forest Department and Sarawak Biodiversity
Centre (permit number NCCD.907.4.4 [JLD.11]-37 and SBC-RA-0102-DO)
respectively, to collect samples from nature reserves in Sarawak and conduct biological
research. A total of twelve samples (Table 2.1) were collected in March 2015 from
FCNR (Figure 2.1) and WCNR (Figure 2.2). These caves are about 5 km south-west of
Bau and 30 km from Kuching (Mohd et al., 2011). The caves are part of the nature
reserves protected by environmental laws that preserve the forest, national parks, and
nature reserve. According to Sarawak forestry department (1992), these caves covers 56
and 6.16 hectares respectively and are largely surrounded by forests. FCNR and WCNR
are also part of Bau limestone areas, covering about 150 km2 in Southwest Sarawak
(Mohd et al., 2011). The samples were collected using aseptic techniques and all
samples were stored in the refrigerator at 4oC until the enrichment and isolation
procedures were fully completed.
Table 2.1: Description of samples collected from FCNR and WCNR
WC= Wind cave; FC= Fairy cave; oC= temperature; (%)RH = relative humidity
Sample ID Sample collected Colour Texture oC (%)
RH WC1 Drapery Grey Coarse 29.2 76
WC2 Stalactite White Silk 29.7 78
WC3 Stalactite White Coarse 29.1 80
WC4 Drapery White Coarse 29.2 84
WC5 Stalactite White Fine 30.1 73
WC6 Mudbank Brown Coarse 30.5 76
WC7 Liquid nil nil 28.8 79
FC1 Soil Black Silk 27.2 84
FC2 Soil Grey Coarse 24.8 86
FC3 Soil Brown Clay 26.5 93
FC4 Soil Brown Fine 28.5 90
FC5 Soil Brown Coarse 30.4 89
53
Figure 2.1: Sampling collection site situated in FCNR, Bau, Sarawak. Samples were collected from regions surrounded by rocks and vegetation.
Figure 2.2: Sampling collection site situated in WCNR, Bau, Sarawak. Samples were collected from regions inside the cave chamber.
54
2.3.2. Enrichment culturing and bacterial isolation
In this study, using the aforementioned methods in the enrichment culture and bacterial
isolation process, a total of ninety morphologically distinct urea degrading bacteria
(UDB) colonies were successfully isolated from samples collected from FC and WC in
Sarawak, Malaysia. The samples collected from FCNR and WCNR were subsequently
cultured in different growth medium to target a variety of bacterial species capable of
hydrolyzing urea. Six percentage (6%) of urea was used in the enrichment culture
medium was used in order to screen for microorganisms capable of surviving at high
urea concentration and potentially able to produce high urease enzyme. During
incubation of the enriched samples, which occurred for a total duration of 12 hr, a
pungent smell was observed at 48 hr of incubation which suggests the release of
ammonia as a result of urea degradation by the production of urease from the
microorganisms in the enrichment culture samples. The isolates were selected from
nutrient agar plates containing a variety of microbial colonies as shown in Figure 2.3.
The selected UDB were consequently sub-cultured onto separate nutrient agar with
higher urea substrate percentage, to target isolates capable of degrading 6% urea. Pure
colonies of the UDB are shown in Figure 2.4. All ninety bacterial isolates were able to
grow on nutrient agar (6% urea). The purified bacterial colonies were then preserved as
glycerol (25%) stock at -80oC using a method adapted by Fortier and Moineau (2009).
Figure 2.3: Microorganisms grown on nutrient agar plates supplemented with 2% urea. The plates were incubated for 24-48 hr at 32°C.
55
Figure 2.4: Pure colonies of urea degrading bacteria after enrichment culture. Cave bacterial isolates (A) NB23, (B) TSB55, (C), (D) TSB 46, (E) TSB40 and (F) LB28 grown on nutrient agar plates supplemented with 6% urea and were incubated for 24 hr at 32°C to acquire pure bacterial colonies. 2.3.3. Selection of urease producing bacteria
The screening for UPB was conducted using UAB medium in test tubes as shown in
Figure 2.5. The colour changes of the test tubes from pale yellow to pink-red indicated
positive urease production. Out of the ninety bacteria isolates subcultured from the cave
samples, thirty-one bacterial isolates were selected based on the ability of the isolates to
completely turn the UAB medium pink in comparison to other isolated urease
producing bacteria and the control strain used in this study. The time taken for the
bacterial isolates and the control strain (Sporosarcina pasteurii, DSM33) to turn the
UAB medium pink was observed and noted.
A
D
C
E F
B
56
In Table 2.2, the control strain, bacterial isolates NB33, LPB21, NB28, LPB4, NB30,
NB40 and LPB4 were able to completely turn their respective UAB medium from
yellow to pink between 24-30 hr of incubation period while other UPB isolates were
able to change their respective UAB medium to pink between 36-120 hr of incubation.
The bacterial isolates which were unable to produce urease enzyme by turning the UAB
medium from pale yellow to pink were discarded.
Figure 2.5: Urease production test using UAB medium. The bacterial isolates were incubated at 37oC for 120 hr to test their ability to produce urease enzyme. Out of the 90 bacteria isolates subcultured from the cave samples, 31 bacterial isolates were able to turn the UAB medium from yellow to pink
57
Table 2.2: Hydrolysis of urea by isolates UAB medium
No Isolate ID Time (hr)
1 Control 24 2 NB33 26 3 LPB21 24 4 NB28 28 5 NB40 30 6 LPB4 30 7 TSB21 76 8 NB30 24 9 TSB4 120 10 TSB14 48 11 TSB46 38 12 BHIB17 68 13 BHIB18 70 14 NB23 70 15 TSB55 62 16 TSB31 36 17 TSB40 46 18 TSB29 40 19 TSB12 38 20 TSB8 48 21 LPB22 78 22 BHIB15 60 23 TSB20 120 24 LB6 86 25 LB48 82 26 LB1 70 27 LPB41 60 28 LB31 48 29 TSB2 72 30 A63 60 31 B53 60 32 A62 64
58
2.3.4. Phenotypic characterisation
The thirty-one isolates were initially identified using phenotypic characterizations such
as morphology, microscopic and biochemical analysis. Macroscopic morphological
analysis of the UPB colonies such as shapes, colours of the colonies, the diffusible
pigmentation of the isolates, to name a few, were observed and recorded. The colony
morphology of the bacterial isolates sub-cultured on nutrient agar media is described in
Table 2.3. It was observed that the UPB were isolated from all the samples collected
from FCNR and WCNR except enrichment sample FC1. However, the UPB isolated
from enrichment sample WC6 and WC1 showed the most number of bacterial isolates.
It was also noticeable that most of the UPB colonies had brownish-white and brown
pigmentations, circular forms and smooth surfaces. The microscopic and biochemical
analysis which were used in this study to further classify the analytic bacterial isolates
as detailed in Table 2.4 and Table 2.5 were performed under standard methods. The
majority of the bacterial isolates were Gram-positive bacteria while only three of the
isolates (A63, B53, and A62) were Gram-negative bacteria. Gram staining analysis also
showed the majority of the bacterial cells were rod-shaped except for NB23 which was
a coccus. Endospore staining test results indicate that all except NB23 were spore
forming bacteria. Oxidase and motility test indicated that all bacterial isolates except
TSB21, NB23, TSB14 and LPB41 tested positive. Catalase test showed that all bacterial
isolates except LPB41 and TSB14 tested positive.
59
Table 2.3: Morphological characteristics of isolated bacterial colonies Cave
Origin Isolate
ID Form
(shape) Size
(mm) Optical
Property Pigmentation
WC6 NB33 circular 4 transparent brownish-
white WC6 LPB21 circular 5 opaque brownish-
white WC1 NB28 circular 4 translucent brownish-
white WC1 NB40 circular 6 transparent brown WC2 LPB4 filamentous 12 transparent whitish-
yellow FC4 TSB21 circular 6 transparent white FC5 NB30 circular 3 transparent brownish-
yellow WC1 TSB4 irregular 10 transparent white WC6 TSB14 irregular 3 transparent brown WC3 TSB46 irregular 9 translucent brown FC2 BHIB17 irregular 5 translucent brownish-
yellow FC2 BHIB18 circular 7 opaque brown WC2 NB23 circular 5 transparent brown WC3 TSB55 circular 1 transparent white WC7 TSB31 circular 8 translucent brownish-
yellow FC5 TSB40 irregular 6 transparent brown WC3 TSB29 irregular 4 opaque brownish-
white FC3 TSB12 circular 6 opaque brownish-
white WC5 TSB8 circular 3 translucent brownish-
white WC5 LPB22 irregular 5 transparent brown WC5 BHIB15 circular 2 transparent white FC4 TSB20 irregular 6 transparent brown WC3 LB6 irregular 4 transparent brownish-
yellow WC6 LB48 irregular 3 opaque white WC6 LB1 circular 6 transparent brown FC3 LPB41 circular 2 translucent white FC5 LB31 circular 4 opaque brown WC1 TSB2 irregular 5 transparent brown FC2 A63 circular 1 translucent creamy WC4 B53 circular 1 opaque creamy WC4 A62 circular 1 opaque creamy
60
Table 2.4: Microscopic characteristics of bacterial isolates
+ve= positive; -ve= negative.
Isolate ID
Gram stain
Cell shape
Endospore stain
NB33 +ve, purple rod +ve LPB21 +ve, purple rod +ve NB28 +ve, purple rod +ve NB40 +ve, purple rod +ve LPB4 +ve, purple rod +ve TSB21 +ve, purple rod +ve NB30 +ve, purple rod +ve TSB4 +ve, purple rod +ve TSB14 +ve, purple rod +ve TSB46 +ve, purple rod +ve BHIB17 +ve, purple rod +ve BHIB18 +ve, purple rod +ve NB23 +ve, purple coccus -ve TSB55 +ve, purple rod +ve TSB31 +ve, purple rod +ve TSB40 +ve, purple rod +ve TSB29 +ve, purple rod +ve TSB12 +ve, purple rod +ve TSB8 +ve, purple rod +ve LPB22 +ve, purple rod +ve BHIB15 +ve, purple rod +ve TSB20 +ve, purple rod +ve LB6 +ve, purple rod +ve LB48 +ve, purple rod +ve LB1 +ve, purple rod +ve LPB41 +ve, purple rod +ve LB31 +ve, purple rod +ve TSB2 +ve, purple rod +ve A63 -ve, pink rod +ve B53 -ve, pink rod +ve A62 -ve, pink rod +ve
61
Table 2.5: Biochemical characteristics of bacterial isolates
+ve= positive; -ve= negative.
Isolate ID Oxidase Catalase Motility
NB33 +ve +ve +ve LPB21 +ve +ve +ve NB28 +ve +ve +ve NB40 +ve +ve +ve LPB4 +ve +ve +ve TSB21 -ve -ve -ve NB30 +ve +ve +ve TSB4 +ve +ve +ve TSB14 -ve -ve -ve TSB46 +ve +ve +ve BHIB17 +ve +ve +ve BHIB18 +ve +ve +ve NB23 -ve +ve -ve TSB55 +ve +ve +ve TSB31 +ve +ve +ve TSB40 +ve +ve +ve TSB29 +ve +ve +ve TSB12 +ve +ve +ve TSB8 +ve +ve +ve LPB22 +ve +ve +ve BHIB15 +ve +ve +ve TSB20 +ve +ve +ve LB6 +ve +ve +ve LB48 +ve +ve +ve LB1 +ve +ve +ve LPB41 -ve -ve -ve LB31 +ve +ve +ve TSB2 +ve +ve +ve A63 +ve +ve +ve B53 +ve +ve +ve A62 +ve +ve +ve
62
2.3.5. Molecular characterization
The DNA templates from the thirty-one UPB were successfully extracted using freeze
and thaw method adapted from Muramatsu et al.(2003). In this study, using the
aforementioned method in PCR amplification process, the DNA bands were visualised
on 1% agarose gel containing Midori Green (Nippon Genetics Europe GmbH) using gel
doc XR system (Biorad). The forward and reverse primer sequences were removed
before the sequence were blasted with existing sequences in NCBI GenBank database
(Zhang et al., 2000) using BLAST nucleotide collection database program to search for
closest best match sequences (Tan et al., 2011, Ashelford et al., 2005).
The nucleotide BLAST analysis results of the 16S rRNA region displayed a rational
level of correlation with the physiological characterization, especially the
morphological descriptions of species within the genus (Achal et al., 2011). All
bacterial isolates from limestone caves of Sarawak showed high degrees of similarity
(91-99%) to their respective closest bacterial species as shown in Table 2.6. The
BLAST results suggested that the UPB were closely related to bacteria from the
Sporosarcina pasteurii group, Pseudogracilibacillus auburnensis group,
Staphylococcus aureus group, Bacillus lentus group, Sporosarcina luteola group and
Bacillus fortis group. The result in Table 2.6 also suggest that the isolated ureolytic
bacteria can be classified into their closest relative groups as Sporosarcina pasteurii
(NB33, LPB21, NB28, NB40, LPB4, NB30, TSB4, TSB46, BHIB17, BHIB18, TSB31,
TSB40, TSB29, TSB12, TSB8, LPB22, BHIB15, TSB20, LB6, LB1 and TSB2),
Pseudogracilibacillus auburnensis (TSB21, TSB14 and LPB41), Staphylococcus
aureus (NB23), Bacillus lentus (TSB55), Sporosarcina luteola (LB48 and LB31) and
Bacillus fortis (A63, B53, and A62).
Results from the taxonomic composition of the UPB using ribosomal database project
(RDP-II), specifically with the aid of the SeqMatch tool confirmed the taxonomy
database classification and nomenclature for the UPB in the public sequence databases
as shown in Table 2.7. The findings suggest the majority of the UPB were classified
into the family and genus of Planococcaceae Sporosarcina while the rest UPB were
classified into the family and genus Bacillaceae Bacillus and Staphylococcaceae
Staphylococcus. A phylogenetic tree shown in Figure 2.7 was constructed using a
neighbour-joining method (Liang et al., 2008).
63
The phylogenetic tree suggests that the majority of the isolated UPB were related to
Sporosarcina pasteurii group. However, there was still satisfactory diversity in the 16
rDNA to differentiate the ureolytic bacterial isolates into distinctive clusters (Devos et
al., 2005) which were noticeably acknowledged in Figure 2.6. The largest clusters
observed from the phylogenetic tree suggest that majority of the ureolytic isolates are
closely related to Sporosarcina pasteurii. However, LPB22 and TSB2 were grouped in
a cluster together while TSB20 was grouped as an independent cluster, but they were
derived from a common ancestor member of Sporosarcina pasteurii. Isolate A63, B53,
and A62 were grouped in one cluster while TSB21, TSB14, and LPB41 were grouped
in another cluster. However, isolates TSB55 and NB23 were as independent clusters
and were noticeably distant from others.
64
Table 2.6: Molecular identification based on 16S rRNA sequencing data using NCBI nucleotide BLAST database
No Isolate ID
GenBank Accession number
Closest match Base pair Query Cover
Similarity
1 NB33 KX212190 Sporosarcina pasteurii strain WJ-4 [KC211296] 1198 100% 97%
2 LPB21 KX212191 Sporosarcina pasteurii strain fwzy14 [KF208477] 1385 95% 97%
3 NB28 KX212192 Sporosarcina pasteurii strain WJ-5[KC211297] 1280 90% 96% 4 NB40 KX212193 Sporosarcina pasteurii strain WJ-5 [KC211297] 1200 99% 97%
5 LPB4 KX212194 Sporosarcina pasteurii strain WJ-4 [KC211296] 1298 98% 97%
6 TSB21 KX212195 Pseudogracilibacillus auburnensis [KR153879] 1050 99% 93%
7 NB30 KX212196 Sporosarcina pasteurii strain fwzy14 [KF208477] 1279 99% 98%
8 TSB4 KX212197 Sporosarcina pasteurii strain WJ-4 [KC211296] 599 99% 99%
9 TSB14 - Pseudogracilibacillus auburnensis [KR153879] 1119 93% 94%
10 TSB46 KX212198 Sporosarcina pasteurii strain WJ-4 [KC211296] 1219 98% 96%
11 BHIB17 KX212199 Sporosarcina pasteurii strain WJ-4 [KC211296] 1200 93% 97%
12 BHIB18 - Sporosarcina pasteurii strain WJ-4 [KC211296] 1147 90% 96%
13 NB23 - Staphylococcus aureus strain CICC [KJ643929] 1275 95% 95%
14 TSB55 KX212200 Bacillus lentus strain NBRC 16444 [NR112631] 920 99% 91%
15 TSB31 KX212201 Sporosarcina pasteurii strain WJ-5 [KC211297] 1219 99% 97%
16 TSB40 KX212202 Sporosarcina pasteurii strain WJ-5 [KC211297] 1159 100% 98% 17 TSB29 KX212203 Sporosarcina pasteurii strain WJ-4 [KC211296] 1250 99% 98%
18 TSB12 KX212204 Sporosarcina pasteurii strain fwzy14 [KF208477] 1200 100% 99%
19 TSB8 - Sporosarcina pasteurii strain fwzy14 [KF208477] 1150 81% 97%
65
20 LPB22 KX212205 Sporosarcina pasteurii strain WJ-5 [KC211297] 1151 99% 96%
21 BHIB15 KX212206 Sporosarcina pasteurii strain fwzy14 [KF208477] 1198 100% 99%
22 TSB20 KX212207 Sporosarcina pasteurii strain WJ-4 [KC211296] 1250 99% 95%
23 LB6 KX212208 Sporosarcina pasteurii strain WJ-4 [KC211296] 1110 100% 99%
24 LB48 KX212209 Sporosarcina luteola strain WJ-1 [KF208477] 1269 92% 98%
25 LB1 KX212210 Sporosarcina pasteurii strain WJ-5 [KC211293] 1374 93% 97%
26 LPB41 KX212211 Pseudogracilibacillus auburnensis [KR153879] 1298 99% 95%
27 LB31 KX212212 Sporosarcina luteola strain WJ-1 [KF208477] 1149 99% 99%
28 TSB2 KX212213 Sporosarcina pasteurii strain WJ-3 [KC211295] 1267 98% 97%
29 A63 KX212214 Bacillus fortis strain R-6514 [NR042905] 1250 98% 96%
30 B53 KX212215 Bacillus fortis strain R-6514 [NR042905] 1325 99% 97%
31 A62 KX212216 Bacillus fortis strain R-6514 [NR042905] 1248 100% 97%
66
Table 2.7: The nomenclatural taxonomy obtained using Ribosomal Database Project-II database
No Isolate ID Domain Phylum Class Order Family Genus
1 NB33 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
2 LPB21 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
3 NB28 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
4 NB40 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
5 LPB4 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
6 TSB21 Bacteria Firmicutes Bacilli Bacillales Bacillaceae Bacillus
7 NB30 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
8 TSB4 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
9 TSB14 Bacteria Firmicutes Bacilli Bacillales Bacillaceae Bacillus
10 TSB46 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
11 BHIB17 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
12 BHIB18 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
13 NB23 Bacteria Firmicutes Bacilli Bacillales Staphylococcaceae Staphylococcus
14 TSB55 Bacteria Firmicutes Bacilli Bacillales Bacillaceae Bacillus
15 TSB31 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
16 TSB40 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
17 TSB29 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
18 TSB12 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
67
19 TSB8 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
20 LPB22 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
21 BHIB15 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
22 TSB20 Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina Firmicutes
23 LB6 Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina Firmicutes
24 LB48 Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina Firmicutes
25 LB1 Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina Firmicutes
26 LPB41 Firmicutes Bacilli Bacillales Bacillaceae Bacillus Firmicutes
27 LB31 Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina Firmicutes
28 TSB2 Bacteria Firmicutes Bacilli Bacillales Planococcaceae Sporosarcina
29 A63 Bacteria Firmicutes Bacilli Bacillales Bacillaceae Bacillus
30 B53 Bacteria Firmicutes Bacilli Bacillales Bacillaceae Bacillus
31 A62 Bacteria Firmicutes Bacilli Bacillales Bacillaceae Bacillus
68
Figure 2.6: Phylogenetic tree based on the bacterial 16S rRNA gene sequence data sequence from different isolates of the current study along with sequences available in the GenBank database (Kang et al., 2014a). The results show that the bacterial isolates were identified as Sporosarcina pasteurii, Pseudogracilibacillus auburnensis, Staphylococcus aureus, Bacillus lentus, Sporosarcina luteola and Bacillus fortis. The tree was constructed using Molecular Evolutionary Genetic Analysis (MEGA) version 6 (Tamura et al., 2013, Tan et al., 2011). Numerical values indicate bootstrap percentile from 1,000 replicates. Bar, 0.005 substitutions per nucleotides (Kang et al., 2014c).
69
2.3.6. Measurement of conductivity
Urease activity was measured through changes in conductivity (mS.cm-1) in the absence of
calcium ions (Whiffin, 2004). The conductivity (mS.cm-1) of each locally isolated
ureolytic bacteria and the control strain were measured for duration of 5 min and the
gradient was obtained from the curve of conductivity (mS.cm-1) against time (hr). Each of
the conductivity measured for individual UPB was performed in three trials, each having
three replicates to obtain data presented as mean ±SE (standard deviation). Figure 2.7
showed a curve of relative conductivity for bacterial culture from LPB21 where the
conductivity measured at 0 min was 6.98 mS.cm-1. The conductivity variation rate for
isolate LPB21 was 0.142 mS.cm-1.min-1 as shown in Figure 2.7. The conductivity variation
rate for the rest of UPB isolates and the control strain are respectively shown in Table 2.8.
The result from table 2.8 showed bacterial isolates NB33, LPB21, NB28, NB30 and
control strain had 0.194, 0.169, 0.132, 0.140 and 0.127 mS.cm-1.min-1 respectively,
suggesting they had the highest conductivity variation rate when compared to the rest
isolates. However, in comparison to all the isolates including the control strain, NB33
showed the highest conductivity variation rate which is 0.194 mS.cm-1.min-1, while TSB4
showed the lowest conductivity variation rate which is 0.010 mS.cm-1.min-1.
2.3.7. Urease Activity Assay
The urease activity of the locally isolated ureolytic bacteria was calculated and compared
to that of the control strain. Table 2.9 showed the conductivity variation rate (mS.cm-
1.min-1) to urease activity (mM urea hydrolysed.min-1)The conductivity was multiplied by
the dilution factor (df) and the constant (11.11) derived by Whiffin (2004), based on the
correlation that 1 mS.cm-1.min-1 corresponds to a hydrolysis activity of 11 mM urea.min-1
in the measured range of activities (Paassen, 2009). The urease activity shown in Table 2.9
showed bacterial isolates NB33, LPB21, NB28, NB30, and control strain had 21.513,
18.768, 14.636, 15.587 and 14.087 mM urea hydrolysed.min-1 respectively, suggesting
they had the highest urease activities when compared to the rest isolates. However, in
comparison to all the isolates including the control strain, NB33 showed the highest urease
activity which is 21.513 mM urea hydrolysed.min-1, while TSB4 showed the lowest urease
activity which is 1.130 mM urea hydrolysed.min-1.
70
Figure 2.7: Relative conductivity of isolate LPB21 measured for a duration of 5 min. Error bars represent standard error of the mean.
y = 0.1415x + 0.0862
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
0 1 2 3 4 5 6
Co
nd
uct
ivit
y (
mS
.cm
-1)
Time (min)
71
Table 2.8: Measurement of conductivity variation rate and SEM
No Isolate ID
Conductivity variation rate (mS.cm-1.min-1) SEM
1 Control 0.127 0.027 2 NB33 0.194 0.057 3 LPB21 0.169 0.038 4 NB28 0.132 0.058 5 NB40 0.083 0.019 6 LPB4 0.103 0.023 7 TSB21 0.059 0.022 8 NB30 0.14 0.018 9 TSB4 0.01 0.006 10 TSB14 0.069 0.015 11 TSB46 0.091 0.010 12 BHIB17 0.089 0.021 13 BHIB18 0.085 0.008 14 NB23 0.065 0.038 15 TSB55 0.067 0.039 16 TSB31 0.103 0.024 17 TSB40 0.078 0.021 18 TSB29 0.087 0.014 19 TSB12 0.112 0.008 20 TSB8 0.094 0.043 21 LPB22 0.056 0.004 22 BHIB15 0.071 0.027 23 TSB20 0.016 0.001 24 LB6 0.053 0.016 25 LB48 0.052 0.026 26 LB1 0.06 0.011 27 LPB41 0.075 0.011 28 LB31 0.114 0.017 29 TSB2 0.058 0.015 30 A63 0.113 0.033 31 B53 0.089 0.005
72
Table 2.9: Conversion of changes in conductivity to urease activity
No Isolate ID (mS.cm-1.min-1) (mS.cm-1.min-1) *df
mM urea hydrolysed.min-1
1 Control 0.127 1.268 14.087 2 NB33 0.194 1.936 21.513 3 LPB21 0.169 1.689 18.768 4 NB28 0.132 1.317 14.636 5 NB40 0.083 0.826 9.181 6 LPB4 0.103 1.027 11.414 7 TSB21 0.059 0.588 6.529 8 NB30 0.140 1.403 15.587 9 TSB4 0.010 0.102 1.130 10 TSB14 0.069 0.695 7.718 11 TSB46 0.091 0.914 10.158 12 BHIB17 0.089 0.890 9.892 13 BHIB18 0.085 0.848 9.425 14 NB23 0.065 0.654 7.270 15 TSB55 0.067 0.666 7.399 16 TSB31 0.103 1.027 11.406 17 TSB40 0.078 0.783 8.703 18 TSB29 0.087 0.871 9.677 19 TSB12 0.112 1.121 12.451 20 TSB8 0.094 0.941 10.458 21 LPB22 0.056 0.562 6.240 22 BHIB15 0.071 0.706 7.847 23 TSB20 0.016 0.159 1.763 24 LB6 0.053 0.531 5.903 25 LB48 0.052 0.523 5.811 26 LB1 0.060 0.600 6.662 27 LPB41 0.075 0.747 8.299 28 LB31 0.114 1.138 12.647 29 TSB2 0.058 0.576 6.396 30 A63 0.113 1.130 12.554 31 B53 0.089 0.890 9.888 32 A62 0.084 0.840 9.332
df = dilution factor; mS.cm-1.min-1= conductivity variation rate; mM urea hydrolysed.min-1= urease activity.
73
2.3.8. Determination of specific enzyme activity
The SUA of the locally isolated ureolytic bacteria were individually calculated and
compared to that of the control strain. The SUA shown in Figure 2.8 was determined as
the amount of urease activity per unit biomass (Whiffin, 2004). The biomass (OD600)
was determined by measuring the optical density of overnight ureolytic bacterial
cultures. The SUA shown in Figure 2.8 showed bacterial isolates NB33, LPB21, NB28,
NB30 and control strain had 19.975, 23.968, 19.275, 20.091 and 17.751 mM urea
hydrolysed.min-1.OD-1 respectively, suggesting they had the highest specific urease
activities when compared to the rest isolates. However, in comparison to all the isolates
including the control strain, LPB21 showed the highest SUA which is 23.968 mM urea
hydrolysed.min-1.OD-1, while TSB4 showed the lowest urease activity which is 1.594
mM urea hydrolysed.min-1.OD-1. Each of these isolates (NB33, LPB21, NB28, NB30,
and control strain) had biomass OD of 1.072, 0.785, 0.738, 0.775 and 0.783 when
measured at a wavelength of 600 nm.
An independent-samples t-test was conducted to compare the SUA of the UPB isolated
from limestone caves of Sarawak against the SUA of the control strain used in this
study. GraphPad program was used to determine if there is a significant difference
between the mean values. The results in Table 2.10 showed out of thirty-one UPB, there
were no significant differences between the SUA of twenty-two UPB when compared
with the control strain. However, bacterial isolate A62 (M=12.4111, SD=0.979), A63
(12.311, SD=3.947), TSB14 (M=12.052, SD=1.527), LPB41 (M=11.480, SD=0.919),
LPB22 (M=9.227, SD=0.0242), TSB2 (M=9.171, SD=2.096), TSB20 (M=2.497,
SD=0.341) and TSB4 (M=1.594, SD=0.768) showed there were significant differences
between their respective SUA when compared with the control strain (M=17.751,
SD=2.345). In addition, four bacterial isolates with the highest SUA as shown in Figure
2.8 were also compared with the control strain to test for statistical analysis. The result
in Table 2.10 showed that SUA of LPB21 (M=23.968, SD=5.722), NB30 (M=20.091,
SD=1.849), NB (M=19.975, SD=5.227) and NB (M=19.275, SD=5.512) were not
significantly different from the SUA of the control strain (M=17.751, SD=2.345).
74
The rationale of this study was to isolate and screen for UPB with comparable urease
production or SUA with the control strain, Sporosarcina pasteurii, (DSM33) type strain
purchased from the Leibniz Institute DSMZ-German Collection of Microorganisms and
Cell Cultures (Braunschweig, Germany). The results from Figure 2.8 and Table 2.10
supports the suggestion that the urease production of ureolytic bacteria LPB21, NB30,
NB33, and NB33 are comparable with the control strain as their respective SUA are
higher than that of the control strain and the t-test analysis also confirms there were no
significant differences between their independently mean values (SUA), thus
confirming suggestion that SUA of the aforementioned bacterial isolates is comparable
with the control strain used in this study.
The effectiveness of these isolates to show comparatively high SUA justify the decision
to confine the selection of these ureolytic bacteria for the subsequent studies in this
chapter and in chapter four. The decision to choose only four ureolytic bacteria (LPB21,
NB30, NB33, and NB28), out of the thirty-one ureolytic bacteria isolated from
limestone caves of Sarawak is because of the SUA these bacteria showed in comparison
to other bacterial isolates and control strain.
75
Figure 2.8: Specific urease activity (mM urea hydrolysed.min-1.OD-1) of urease-producing bacteria and the control strain. Error bars represent standard error of the mean.
0
5
10
15
20
25
30
Spe
cifi
c u
reas
e a
ctiv
ity
(mM
ure
a h
ydro
lyse
d-1
.min
.OD
-1)
76
Table 2.10: t-test results comparing the specific urease activity differences between individual isolated urease-producing bacteria and control strain. (N=3; df=2)
No Isolate ID M SD SE P-value t P <*
1 control 17.751 2.345 nil nil nil nil 2 LPB21 23.968 5.722 4.407 0.294 1.4107 - 3 NB30 20.091 1.849 4.217 0.339 1.2471 - 4 NB33 19.975 5.227 2.667 0.651 0.5272 - 5 NB28 19.275 5.512 2.904 0.625 0.5714 - 6 TSB12 17.400 2.839 1.774 0.9149 0.1207 - 7 TSB31 16.397 2.963 3.174 0.525 0.7634 - 8 LPB4 14.063 3.564 2.823 0.365 1.1624 - 9 TSB29 13.618 2.623 2.190 0.281 1.4641 - 10 B53 13.562 1.989 1.343 0.196 1.9128 - 11 LB31 13.052 0.605 1.555 0.073 3.4989 - 12 NB40 13.003 2.430 3.880 0.093 3.0542 - 13 TSB8 12.714 5.285 4.566 0.324 1.2983 - 14 TSB55 12.715 7.231 2.265 0.3850 1.103 - 15 BHIB15 12.473 2.172 0.880 0.145 2.3299 - 16 A62 12.411 0.979 1.123 0.026 6.0649 + 17 A63 12.311 3.947 2.931 0.040 4.8438 + 18 TSB21 12.089 5.985 0.558 0.193 1.9323 - 19 TSB14 12.052 1.527 2.931 0.010 10.2095 + 20 TSB40 11.994 2.743 0.934 0.1930 1.9323 - 21 LPB41 11.480 0.919 1.745 0.022 6.7172 + 22 BHIB17 11.324 1.921 2.060 0.066 3.6838 - 23 TSB46 10.799 2.047 1.983 0.078 3.3742 - 24 LB1 9.956 1.214 3.755 0.059 3.9304 - 25 NB23 9.813 5.070 1.214 0.1688 2.1142 - 26 LBP22 9.227 0.242 1.593 0.020 7.0212 + 27 TSB2 9.171 2.096 2.296 0.033 5.3851 + 28 BHIB18 8.822 1.646 3.513 0.060 3.8881 - 29 LB48 8.652 4.073 2.450 0.122 2.59 - 30 LB6 7.590 3.133 1.267 0.054 4.1468 - 31 TSB20 2.497 0.341 1.594 0.007 12.0372 + 32 TSB4 1.594 0.768 4.152 0.010 10.1363 +
N= number of sample size; df= degree of freedom; M=mean; SE= standard error; SD= standard deviation; P-value= calculated probability; t= test statistic; += significant; -= not significant; *= P-value is significant at 0.05 level.
77
2.3.9. Microbial calcite precipitates
The CPM was used to test the ability of the UPB to induce calcium carbonate. Bacterial
isolates LPB21, NB28, NB33, NB30, and control strain were selected for this
experiment and the remaining subsequent experiments in this study because they
showed highest enzyme activities in Figure 2.8 and Table 2.9. The bacterial isolates
were cultivated for 24 hr and the serially diluted before being spread on the CPM. The
CPM was studied through visual observation for the formation of precipitates on the
CPM upon addition of bacterial cultures. When tested on CPM, UPB isolates, and
control strain was able to induce precipitate calcite after being incubated at 30°C for 6
days. Milky-white crystal was observed covering the colonies grown on the CPM and
appeared at the 4th day of incubation seen in Figure 2.9. All the precipitates grown on
the CPM appeared as a distinct circular zone around the growth area of the bacterial
colonies. Based on the morphology of the precipitation there was no difference in
crystal formation on the agar plates, all isolates induced the same morphological sizes,
shape and colour of the precipitates.
Figure 2.9: Calcite precipitation media. The appearance of calcite precipitates on bacterial colonies on the 4th day of incubation at 30°C.
78
2.3.10. Calcite estimation
The calcite precipitation induced by LPB21, NB28, NB33, NB30, and control strain
were studied by using a modified method of Wei et al. (2015). Calcite precipitates were
quantified after nutrient broth supplemented with 2% urea and 2% calcium chloride
solutions inoculated with overnight grown cultures under shaking condition (150rpm) at
30°C for incubation period of 7 days.
Figure 2.10: Comparison of calcite precipitated by selected UPB isolates and the control strain. Error bars represent standard error of the mean.
0
2
4
6
8
10
12
14
16
18
20
control NB30 LPB21 NB33 NB28
we
igh
t o
f ca
lcit
e p
reci
pit
ate
s(m
g.m
L-1
)
79
The moment the bacterial cultures were inoculated into the broth media, white
precipitates appeared instantly at the bottom of the conical flasks and its density
increased with incubation. At the end of 7 days incubation, the precipitates were
collected and weighed. All five isolate (including control strain) produced a similar
amount of calcite precipitates at the end of the incubation period. Figure 2.10 showed
the bacterial NB33, LPB21, NB28, NB30, and control strain induced 12.44, 15.82,
10.51, 17.00 and 10.51 mg.mL-1 of calcite precipitates, respectively. This finding
suggests that UPB strain NB30 showed the highest productivity of calcite which was
17.0 mg.mL-1.
Table 2.11: t-test results comparing the calcite precipitate differences between individual isolated urease-producing bacteria and control strain. (N=3; df=2)
No Isolate ID M SD SE P-
value t P <*
1 control 10.511 0.834 nil nil nil nil
2 LPB21 15.822 0.731 0.633 0.014 8.392 +
3 NB30 17.000 0.581 0.774 0.014 8.384 +
4 NB33 12.444 0.269 0.329 0.028 5.879 +
5 NB28 10.511 2.672 1.084 1.000 0.000 -
N= number of sample size; df= degree of freedom; M=mean; SE= standard error; SD= standard deviation; P-value= calculated probability; t= test statistic; += significant; -= not significant; *= P-value is significant at 0.05 level.
80
An independent-samples t-test was conducted to compare the calcite precipitated by the
UPB isolates against that of the control strain used in this study. The results in Table
2.11 showed that there were significant differences between the calcite precipitated by
LPB21 (M=15.822, SD=0.731), NB30 (M=17.000, SD=0.581) and NB33 (M=12.444,
SD=0.269) against the control strain (M=10.511, SD=0.0834). On the other hand, there
was no significant difference between the calcite precipitated by NB28 (M=10.5111,
SD=2.672) against the control strain.
2.3.11. Bacterial growth and pH profiles
Optical density (OD) at a wavelength of 600 nm, indicative of bacterial growth, is
presented in Figure 2.11 which was studied up to 12 hr in a batch culture containing
nutrient broth and 6% urea. It was observed from the graph that the growth of the UPB
cells increased in response to time and all the ureolytic bacteria tested had similar
growth patterns for the total duration of the incubation period. Table 2.12 summarises
the results of the kinetic growth of the ureolytic bacteria during the batch culturing. The
specific growth rate (k), from the experimental result, showed the highest value was
0.398 h-1 from bacteria strain NB30 while the control strain showed the lowest value for
specific growth rate which is 0.254 h-1. Doubling time (td), which refers to the time the
bacterial cells doubles, with shorter times implies more rapid growth. By referring to the
result of td in Table 2.12, isolate NB30 showed the shortest td of 1.741 g to double its
cells, while the control strain showed the longest td of 2.726 g to double its cells. The
result for maximum growth (OD600) of each bacterial culture after being studied for 12
hr showed that the OD values for the ureolytic bacterial cultures ranged between 0.882
to 1.009.
81
Table 2.12: Kinetics growth of ureolytic bacteria in batch cultures
The growth profile shown in Figure 2.11 showed that all the bacterial cultures continued
to have a progressive cell growth, hence, a prolonged stationary phase or death phase
was not observed. It was also observed that the cultures showed similar growth pattern.
The figure suggests all the bacterial isolates showed their maximum growth at the 12 hr
of their incubation period. In table 2.12, LPB21 showed the highest value in the
maximum growth yield [OD600] of 1.076, while the lowest value in the maximum
growth yield [OD600] was observed to be 0.882 by isolate NB28.
Isolate
ID
Specific growth rate,k [h-1]
Doubling time,td [g]
Maximum growth of bacteria [OD600]
Control 0.254 2.726 0.914
NB33 0.274 2.535 0.976
LPB21 0.261 2.653 1.079
NB28 0.319 2.172 0.882
NB30 0.398 1.741 1.009
82
Figure 2.11: Growth profile of selected ureolytic bacterial isolates and control strain grown in nutrient broth containing 6% urea for 12 hr. Error bars represent standard error of the mean.
0.0
0.2
0.4
0.6
0.8
1.0
1.2
0 2 4 6 8 10 12 14
OD
600
Time (hour)
Control
NB33
LPB21
NB30
NB28
83
Figure 2.12: pH profile of selected ureolytic bacterial isolates and control strain grown in nutrient broth containing 6% urea for 12 hr. Error bars represent standard error of the mean.
8.6
8.7
8.8
8.9
9.0
9.1
9.2
9.3
9.4
9.5
0 1 2 3 4 5 6 7 8 9 10 11 12
pH
Time (hour)
Control
NB33
LPB21
NB30
NB28
84
The pH of the growth medium containing the bacterial culture was also studied.
Figure 2.12 shows that the pH of the medium significantly increased in correspondence
with the growth of the bacterial curve. An increase in the pH medium corresponds to
urea hydrolysis as a result of urea degradation by the ureolytic bacteria. The pH profile
in Figure 2.12 showed similar profiles among the UPB isolates and control strain.
NB33, LPB21, NB28, NB30, and control strain had final maximum pH values of 9.30,
9.32, 9.31, 9.31 and 9.34 respectively. However, all the isolates experienced a
fluctuation in their respective curves during the incubation.
85
2.4 Discussion The study in this chapter aimed at isolating, screening, identifying UPB from limestone
cave (FCNR and WCNR) samples of Sarawak, and to determine their respective urease
activity in comparison to Sporosarcina pasteurii (DSM33), which served as a
representative strain. In order to screen for ureolytic bacteria from cave regions, was
necessary to select appropriate conditions at which desired microorganism would
survive (Al-Thawadi and Cord-Ruwisch, 2012). Hence, samples were collected from
limestone cave samples of Sarawak, Malaysia. The extreme environmental
characteristics cave regions it possesses, make it able to accommodate the unexpected
diversity of microbial communities. (Tomczyk-Żak and Zielenkiewicz, 2015). These
make Sarawak limestone caves suitable sampling locations as microbial communities in
these environments are enriched and exposed to alkaline and limestone conditions. To
screen for highly active urease producing bacteria, enrichment culture technique was
used to instigate a competitive ecosystem among the microorganism for the availability
of growth nutrients (Gorski, 2012b). Enrichment culture technique is widely used to
isolate bacteria in clinical, biotechnological and environmental studies because it brings
about competition among microbiota for available nutrients and against growth
inhibitors by favouring specific bacterial type isolates or subgroups (Gorski, 2012a,
Maite Muniesa et al., 2005).
The present study showed those microorganisms indigenous to cave regions are
promising sources for urease production with the capability of inducing calcite
minerals. This finding is supported by Banks et al. (2010) who previously isolated fifty-
one bacteria from an unnamed cave region in Kentucky, USA. Their results showed that
majority of these microbial species were capable of inducing calcite precipitates.
Stabnikov et al. (2013) suggested that UPB is common inhabitants of soils with the
consistent provision of urea substrate, a final production of amino metabolism (nitrogen
metabolism) of mammals. Hence, enrichment culture designed to select UPB suitable
for MICP ought to be supplemented with an adequate amount of urea substrate
(Burbank et al., 2012, Chu et al., 2011, Hammes et al., 2003a). It was observed that
during the incubation of the enrichment culture samples, there was a unique pungent
smell, indicating the release of ammonia gas.
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The breakdown of urea by the urease enzyme allows the release of ammonium gas to
the bacteria’s environment. This gas can be poisonous to humans if inhaled and can
cause serious respiratory (Gueye et al., 2001, Woto-Gaye et al., 1999). Hence, it is
recommended to work using a facial mask when handling urease producing bacteria
inside the incubation room. Another precaution which should be taken is to incubate
these bacteria in a small incubator (MMM Incucell 55, MMM Medcenter Einrichtungen
Gmbh) and place the incubator inside a fume hood (BASIX 52, LABCRAFT) to
prevent the discharge and spread of ammonia gas in the laboratory. The isolated UDB
capable of growing on nutrient agar supplemented with 6% urea were screened using
UAB medium to test their respective capability of urease production. The incubation
temperature 32oC was chosen because it is the average temperature of Kuching,
Sarawak. Hence it would aid the growth of a wider variety of mesophilic bacteria in the
enrichment cultures.
Typically, aerobic bacteria are often incubated at 30-35oC for a maximum incubation of
72 hr, which is appropriate for cultivation of bacteria from microbiological growth
media (Moldenhauer, 2014). Kielpinski et al. (2005) reported that incubation
temperature of 32oC provided an improvement detection of a microorganism in a sterile
microbiological growth media. Gordon et al. (2014) suggested that this incubation
temperature is appropriate for the recovery of total aerobic microorganism counts from
sample collection using general microbiological growth medium.
UAB is a differentiation medium that tests the ability of a variety of microorganism to
produce urease, an extracellular enzyme which is secreted outside the cells of
microorganisms (Atlas, 2010). Urease test media often contains 2-4% of urea and
phenol red as a pH indicator, which detects an increase in the pH of the medium due to
ammonia production resulting in colour changes from yellow (pH 6.8) to a bright pink
(8.2) (Brink, 2010). Previously Staurt’s urea broth medium was often used to
distinguish urease producers, however only Proteus species were detected as urease-
positive because this medium only contained essential nutrients that facilitated the
growth of only Proteus species and the medium is a highly buffered medium requiring a
large quantity of ammonia production to raise the pH of the medium above o for colour
change to occur(MacFaddin, 2000, Winn et al., 2006).
87
On the other hand, Christensen’s urea agar (UAB) contains peptone and glucose which
supports growths of a wider variety of urease-producing microorganisms and it also has
a reduced content of buffer which allows a quicker detection of urea degradation (Brink,
2010). Several studies have reported using urea agar base media as a preferred
qualitative urease assay for isolation and differentiation of ureolytic microorganisms
(Hammad et al., 2013b, Elmanama and Alhour, 2013, Dhami et al., 2013c). When urea
is hydrolysed by the bacteria, ammonia is released and becomes accumulated in the
medium which increases the pH of the environment making it alkaline (Hammad et al.,
2013a). This is the first study to show the presence of at least one cultivable bacterium
from FCNR and WCNR with the production of urease capabilities in the presence of
high concentration of urea. The isolation of a few urease producing bacteria isolates
from the collected samples suggests that a small percentage of environmental bacteria
are capable of participating in the precipitation of calcite through urea hydrolysis
(Burbank et al., 2012).
There were noticeable morphological differences among the isolated urease- producing
bacteria. The limited diversity of the bacterial community in limestone environment is
not surprising because of its extreme alkaline condition, only organisms capable of
growing in these conditions can survive in. such an environment (Achal et al.,
2010b).The close morphology of bacterial isolates was observed among the isolates and
it might be as a result of the dominance species which might occur during enrichment
culturing period since Bacillus species are usually selected by the isolation and
cultivation methods (Shannon Stocks-Fischer et al., 1999). Physiological properties of
the majority of the bacterial isolates resemble those of Bacillus and Sporosarcina
species previously reported (Tominaga et al., 2009, Stocks-Fischer et al., 1999, Gordon
and Hyde, 1982, Gordon et al., 1973) which were then verified via 16S rRNA gene
analysis. The genus bacillus and Sporosarcina appear to be inhabitants of extreme
environments (Achal and Pan, 2011). Achal and Pan (2011) reported in their study that
one of the reasons ureolytic bacteria are capable of enduring alkaline pH might be as a
result of their alkaline habitat of which the aforementioned bacterial isolates were also
isolated from.
88
A study from Aono et al. (1999) on contribution of the cell wall component
teichuronopeptide to pH Homeostasis and alkaliphilic in the alkaliphilic Bacillus lentus
C-125 reported that some of the cell walls of some alkaliphiles such as teichurono-
peptide may play a role in the pH homeostasis at alkaline pH and support the bacteria
isolates to survive in extreme environments.
Based on sequence data of the 16S of the rRNA region, all bacterial isolates from
Sarawak limestone caves showed a high degree of similarity (91-99%) to their closest
species. The results in the phylogenetic tree suggest that the ureolytic bacteria isolated
from samples collected from Sarawak cave were identified as Sporosarcina pasteurii,
Pseudogracilibacillus auburnensis, Staphylococcus aureus, Bacillus lentus,
Sporosarcina luteola and Bacillus fortis. It is noteworthy that the BLAST analysis
showed that most of the local isolates revealed less than 98% similarity to their closest
species. The Sporosarcina comprised 65% of the cultivable ureolytic bacteria isolated
from the cave samples. This isn’t surprising as various studies have reported the
identifying majority of their locally bacterial isolates as Sporosarcina pasteurii. In
addition, a recent study by Wei et al. (2015) reported isolating ureolytic bacteria from
marine sediments of which majority of the identified bacterial isolates were
Sporosarcina sp. However, a few other studies reported isolating bacillus sp.
(Stabnikov et al., 2013, Burbank et al., 2012, Banks et al., 2010).
Sporosarcina pasteurii, majority of the bacteria isolated from FC and WC samples, has
been reported to be non-pathogenic, however sometimes isolated from human faeces with
no pathological reaction happening from its association (Ranganathan et al., 2006), but
it's been suggested it may comprise the immune system of infected patients because
Sporosarcina pasteurii possess abilities to significantly reduce blood urea nitrogen levels
(Arpita et al., 2013, Alhour, 2013, Yoon et al., 2001). Sporosarcina luteola, from the
same genus as Sporosarcina pasteurii, is also non-pathogenic and commonly isolated
from soil samples (Tominaga et al., 2009).
Pseudogracilibacillus auburnensis, which is among the bacteria isolated from FC and
WC samples, has been reported to be a bacterial abundantly available and often isolated
from lakes, desert soil, saline soil and has been reported to be application in controlling
plant pathogens (Mandic-Mulec et al., 2015, Glaeser et al., 2014, Waino et al., 1999).
89
Staphylococcus aureus, one of the bacteria isolated from FC and WC samples, has been
reported to be a virulent pathogen, presently the most common cause of bacterial
infections in hospitalised patients. It's frequently isolated from urine samples obtained
from long-term care patients this pathogenic bacteria’s infection can involve any organ
system has an increasing resistance to antibacterial agents (Rasigade and Vandenesch,
2014, Bien et al., 2011, Archer, 1998). Due to its pathogenicity, it is recommended not to
use this microorganism for biocement application.
According to Stabnikov et al. (2013) to some ureolytic bacteria can be pathogenic,
especially isolates such as Helicobacter pylori, Proteus vulgaris, Staphylococcus aureus,
and Pseudomonas aeruginosa. Due to their level of their pathogenicity, they are not
suitable for biocementation applications. Bacillus lentus and bacillus fortis, which is
among the bacteria isolated from FC and WC samples, are commonly isolated from the
soil, marine waters, and extreme environments. They are considered nonpathogenic and
study has shown the isolation of these microbes have been isolated from dairy farms and
cave environments (Banks et al., 2010, Scheldeman et al., 2004). All the bacterial isolates
except Staphylococcus aureus (NB23) are capable of forming endospore, which has
special resistant dormant structures formed within a cell making the bacteria able to
survive harsh or hostile environments (Krishnapriya et al., 2015). This makes these
isolates suitable for various biocementation applications.
According to Al-Thawadi (2008), conductivity can be used to determine the enzymatic
rate of reaction because the device is robust, easy to operate and an inexpensive assay
system. Conductivity measurement is a suitable method to measure urease activity,
because urease turns the urea molecule (non-conductive) into two charged ions:
ammonium (NH4+, positively charged) and carbonates (CO3
2-, negatively charged)
(Cuzman et al., 2015b).
The release of ammonia as a result of urea hydrolysis can be toxic and detrimental to
most bacterial cells especially when the concentration is high (Cheng and Cord-Ruwisch,
2013). This production of ammonia is advantageous to specific bacteria, such as ureolytic
bacteria which uses the ammonia production for the generation of ATP (Cheng and Cord-
Ruwisch, 2013).
90
The relative changes in conductivity of the urea-bacterial solutions were measured at an
ambient temperature (25◦C ±1) for duration of 5 minutes. These measurement conditions
were also performed by Li et al. (2013), Cheng and Cord-Ruwisch (2013) and Cheng and
Cord-Ruwisch (2012). The use of conductivity to measure bacterial urease activity has
been extensively studied and reported (Krishnapriya et al., 2015, Cuzman et al., 2015b,
Cuzman et al., 2015a). The conductivity variation rate (mS.cm-1.min-1) for Sporosarcina
pasteurii (DSM 33) reported in other studies was in a range of 0.083-0.23 mS.cm-1.min-1
(Cuzman et al., 2015b, Whiffin et al., 2007). A study by Hammad et al. (2013a) reported
using Sporosarcina pasteurii (NCIMB 8841) which had an average change in
conductivity of 0.05 mS.cm-1.min-1. However, another study conducted by (Chu et al.,
2012) reported having an average change in conductivity of 0.06 mS.cm-1.min-1 for
halotolerant and Alkaliphilic urease-producing bacteria which were isolated from tropical
beach sand and later identified as Bacillus sp. The finding by these researchers supports
the results presented in this study on conductivity variation rate of ureolytic bacteria.
Urease activity for Sporosarcina pasteurii (DSM33) reported by Harkes et al. (2010) was
between the range of 5 to 20 mM urea hydrolysed.min-1. Another report from Whiffin
(2004) on the urease activity of Sporosarcina pasteurii (ATCC11859) was between 2.2 to
13.3 mM urea hydrolysed.min-1. However, other studies reported locally isolated Bacillus
strains have urease activity between 3.3 to 8.8 mM urea hydrolysed.min-1 (Stabnikov et
al., 2013, Al-Thawadi and Cord-Ruwisch, 2012). The ability of ureolytic bacteria to
induce calcite precipitation after being incubated for duration of 120 hours was reported
(Hammad et al., 2013b, Hammes et al., 2003b). It is suggested that this biomineralization
is not entirely associated with any specific group of microorganisms, however, it is
relatively associated with a wide variety of microorganism (Boquet et al., 1973). The
capability of bacterial isolates to be able to induce calcite precipitates has been widely
studied and reported, hence proving that ureolytic bacteria are capable of inducing
calcium carbonate (Wei et al., 2015, Krishnapriya et al., 2015, Gat et al., 2014).
Bacterial growth curve of ureolytic isolates has been previously studied and reported. It
shows similar growth pathway to that of the local isolates isolated from samples
collected from Sarawak (Krishnapriya et al., 2015, Achal and Pan, 2014, Stabnikov et
al., 2013, Cheng and Cord-Ruwisch, 2013, Chahal et al., 2011, Achal et al., 2009a).
91
The growth and pH profiles of the local cultures were studied up to 12 hours in nutrient
broth culture supplemented with 6% urea. Figure 2.11 and 2.12 showed that the local
cultures had similar growth profile which confirms they are all of the similar genera
(Sporosarcina pasteurii). The maximum pH observed from the local cultures was
around pH9. Other reports showed that maximum pH profile of Sporosarcina pasteurii
(NCIM 2477, type strain) and Bacillus sp. were at pH11 when grown in nutrient broth
supplement with urea. However, 2% urea was used in their study which suggests the
reason why the growth profile and pH profile were slightly different (Achal and Pan,
2014, Cheng and Cord-Ruwisch, 2013).
Studies by Cheng and Cord-Ruwisch (2013) have shown culturing ureolytic bacteria
using chemostat can maximum their enzyme production and working in non-sterile
conditions would not have a significant impact on the enzyme production or the
bacterial culture. Another study by Achal, Mukherjee and Reddy (2010b), suggested the
use of alternative media such as lactose mother liquor for biocementation applications
but reported there were no significant differences in bacterial growth, urease production
and compressive strength among all media used, however, can serve as a better media
for bacterial growth, support calcite precipitation and reduce the cost in biocementation.
92
2.5 Conclusion The study in this chapter reports the isolation of urease-producing bacteria from
samples collected from limestone caves of Sarawak. Ninety urea degrading bacteria
were successfully isolated from Fairy and Wind cave samples. These bacterial isolates
were tested for their abilities to produce urease enzyme. The experiments performed in
this study indicate the presence of urease-producing bacteria in the cave sample. Thirty-
one bacterial isolates were selected based on their abilities to produce urease. DNA
sequence identification classified the thirty-one urease-producing bacteria isolates as
belonging to the genus of Sporosarcina, Pseudogracilibacillus, Staphylococcus, and
Bacillus. However, the majority of the isolates were similar to Sporosarcina pasteurii
when compared to the 16S rRNA sequencing data in NCBI nucleotide BLAST
database. Conductivity method was used to measure the urease activity of the isolates
and the control. The urease activity detected suggest the potential use of these bacterial
isolates in biocementation. However, results from the specific urease activity, indicates
bacterial isolates LPB21, NB30, NB28 and NB33 produced the highest enzyme activity,
thus were selected as the preferred isolates for the rest subsequent experiments
performed due to their high specific urease activities when compared to other isolates
and also the control strain. The selected aforementioned isolates were then used in the
next chapter. Further studies on LPB21, NB30, NB28 and NB33 were performed in
chapter three which involved studying various cultural conditions that affect the
production of urease and Evaluating these isolates efficiency in biocementation.
Chapter
3 EFFECTS OF CULTURAL CONDITIONS ON UREASE
ACTIVITY AND EVALUATION OF BIOCEMENTATION
POTENTIALS IN SMALL SCALE TEST
93
3.1 Introduction Biocementation is a new ground improvement technique that can be used to improve the
geotechnical properties of soil in a way similar to ordinary cement (Chu et al., 2009).
The use of chemicals agents such as lime, asphalt, sodium silicate, and Portland cement
for soil enhancement has been proven successful (Peethamparan et al., 2009, Basha et
al., 2005, Anagnostopoulo and Hadjispyrou, 2004), however these artificial injection
formulas often alter the pH level of soil, contaminates the soils and groundwater,
attributing to their toxic and hazardous characteristics (DeJong et al., 2006, Karol,
2003). The advantage MICP, a type of biocementation technique has over conventional
ground improvement methods is that it requires a range of ambient conditions with the
diminutive usage of fuel or carbon footprint during production, unlike conventional
cement (Dhami et al., 2016). Studies have also shown that MICP process is able to
significantly improve soil’s shear strength and reduce permeability by filling the pores
of the soil with minerals precipitated (Zhang et al., 2015, Feng and Montoya, 2015,
Bundur et al., 2015).The implementation of MICP as an established ground
improvement method has been partially limited by the need for cultivation and injection
of specific bacteria (Gomez et al., 2014). Although various forms of MICP forms are
available with the use of different bacterial and precursor, however, this chapter
divulges a biological approach for manufacturing biocement using a selected number of
locally isolated urease producing bacteria from limestone caves of Sarawak.
The objectives of the study in this chapter are as follows:
i. To optimise various cultural conditions for maximum urease activity. ii. To study in vitro biocementation potential using single and consortia of
ureolytic bacterial isolates. iii. To determine the calcite contents precipitated in the treated sand specimens.
94
3.2 Methods and Materials 3.2.1. The Effect of Cultural Conditions On Urease Activity 3.2.1 (a) Incubation temperature (oC)
The influence of different temperatures ranging from 20 to 45oC 2 with an interval of
5oC were carried out by incubating the ureolytic bacteria cultures for 24 hr, under
aerobic batch conditions at 32oC with agitation at 130 rpm. The bacterial cultures were
grown in nutrient broth media (13.0 g.L-1, HiMedia Laboratories Pvt. Ltd),
supplemented with 4% urea. The overnight grown bacteria were inoculated (2% v/v)
into separate sterile conical flasks (containing 125 mL nutrient broth). The initial pH of
the growth medium used was attuned to pH 7.5 with the use of 1 N NaOH and 1 N HCl.
The conductivity and OD600 were measured and used to determine the specific urease
activity at the end of the cultivation period.
3.2.1 (b) Initial medium pH
The effect of distinctive pH on the ureolytic activity from the selected isolates was
determined by examining urease activity at different pH ranging from 6.0 to 8.5 with an
interval of 0.5. The bacterial cultures were grown in nutrient broth media (13.0 g.L-1,
HiMedia Laboratories Pvt. Ltd), supplemented with 4% urea. The initial pH of the
growth medium used was attuned with the use of 1 N NaOH and 1 N HCl. The bacterial
cultures were incubated at the optimised incubation temperature for the duration of 24
hr, with agitation at 130 rpm. The conductivity and OD600 were measured and used to
determine the specific urease activity at the end of the cultivation period.
3.2.1 (c) Incubation period (hr)
The optimal incubation period was determined by incubating the ureolytic bacteria
culture at different incubation periods ranging from 24 to 96 hr with an interval of 24 hr,
with agitation at 130 rpm and optimised temperature. The bacterial cultures were grown
in nutrient broth media (13.0 g.L-1, HiMedia Laboratories Pvt. Ltd), supplemented with
4% urea. The initial pH of the growth medium used was attuned with the use of
1 N NaOH and 1 N HCl to maintain the optimised pH medium. The conductivity and
OD600 were measured and used to determine the specific urease activity at the end of the
cultivation period.
95
3.2.1 (d) Urea concentration (%)
The influence of urea substrates with varied concentration for enzyme production was
studied. Different urea concentration ranging from 2 to 10% (w/v) with an interval of
2% was selected. The bacterial cultures were grown in nutrient broth media (13.0 g.L-1,
HiMedia Laboratories Pvt. Ltd) at an incubation temperature, initial pH medium and
incubation period previously studied. The conductivity and OD600 were measured and
used to determine the specific urease activity at the end of the cultivation period.
3.2.1 (e) Statistical analysis
The data were presented as mean ±SE (standard deviation) for three replicates. The
optimisation results for different parameters (Incubation temperature, initial medium
pH, incubation period and urea concentration) were analysed using Microsoft Excel
(version 2016) and StatPlus programmes. The analysis of variance (ANOVA) with
Tukey’s procedure was used to compare the variance between different groups with the
variability within each of the groups. The level of significance was set at 0.05.
3.2.2. Small Scale Biocementation Test 3.2.2 (a) Bacteria culture
The selected ureolytic bacteria used in this experiment are shown in Table 3.1. The
ureolytic bacteria were grown under sterile aerobic batch conditions. After incubation,
the bacteria cultures were stored in their growth medium in the fridge at 4oC prior to
use.
Table 3.1: Selected ureolytic bacteria for biocement test Isolate Closest Match
NB33 Sporosarcina pasteurii strain WJ-4 [KC211296]
LPB21 Sporosarcina pasteurii strain fwzy14 [KF208477]
NB28 Sporosarcina pasteurii strain WJ-5[KC211297]
NB30 Sporosarcina pasteurii strain fwzy14 [KF208477] Reference Control Sporosarcina pasteurii strain DSM33
Bacterial Consortia
Comprised of four isolates; Sporosarcina pasteurii LPB21 (SUTS), Sporosarcina pasteurii NB30 (SUTS), Sporosarcina pasteurii NB28 (SUTS) and Sporosarcina pasteurii NB33 (SUTS)
96
3.2.2 (b) Cementation solution
The cementation solutions used to treat the sand columns were modified from (Cheng et
al., 2014, Weaver et al., 2011). The constituents and concentration of the cementation
solution are listed in Table 3.2. All the cementation solution components were
autoclaved except urea and CaCl2, which were added after the solution was autoclaved.
Table 3.2: Biocement treatment components
Constituents Concentration Urea
(CO(NH2)2) 1 M
Calcium chloride (CaCl2) 1 M
Sodium acetate (C2H3NaO2)
0.17 M
Ammonium chloride (NH4Cl) 0.0125 M
Nutrient broth 13 g/L
3.2.2 (c) Preparation of sand columns
The characteristics of the sand used in this experiment are summarised in Table 3.3.
Re-informed paper tubes served as the moulds used in this experiment. The moulds had
an internal diameter of 75 mm and length of 49 mm. Each of the column (mould) was
autoclaved and then packed with 294.73 g of sand. All columns were placed on flat
surfaced polypropylene sheet; five holes were drilled on the surfaces of the
polypropylene sheets to allow the effluents of the cementation solution to pass through.
The polypropylene sheets containing drilled holes were later covered with Whatman
filter papers. A plastic container was placed below the polypropylene sheet to
accumulate the effluents. The top of each column was covered with a layer of scouring
pads (Scotch-BriteTM) as filters to prevent disturbance on the surfaces of the sands
during biocement treatments.
97
Table 3.3: Sand characteristics
Sand Source
Uniformity coefficient
(Cu)
Coefficient of gradation
(Cc)
D10
(mm) D30
(mm) D60
(mm)
Kuching, Sarawak 1.6 0.907 0.220 0.265 0.352
3.2.2 (d) Biocementation treatment method
Prior to the beginning of the treatment, each of the sand was pre-mixed with bacteria
culture, calcium chloride (1M) and urea (1M) solution before being compacted into
their respective columns. The sand columns were treated with the bacteria and
cementation solutions by percolation (i.e. unrestrained flushing of fluid from top to
bottom). The columns were treated twice daily with the 80 mL ureolytic bacteria culture
(Table 3.1) and 80 mL cementation solution (Table 3.2). However, the treatment was
split into two series of treatment and added twice daily. The Sporosarcina pasteurii
isolates, consortia and control strain were grown in nutrient broth media under aerobic
condition (Table 3.5). The grown cultures, Isolate LPB21 (4.8 X 107), isolate NB30 (4.0
X 107), isolate NB33 (1.5 X 107), isolate NB28 (4.1 X 107), consortia (5.0 X 107) and
control strain (4.7 X 107) were harvested at their respective late exponential phases
before being mixed with the air dried sand specimens. The cementation solution
contained cementation reagents, nutrient broth (13 g.L-1), C2H3NaO (0.17 M), NH4Cl
(0.0125 M). The cementation solution used in this study were urea (CO(NH2)2) and
calcium chloride (CaCl2) which were prepared at a concentration of 1.0 M. The MICP
treatment was performed by introducing 80 mL of bacterial culture and 80 mL of
cementation solution into the sand specimens at an interval of 12 hr for a duration of 96
hrs. The treatments of the sand columns were performed inside a fume hood. Upon
completion of the treatments, all the sand columns were cured at room temperature for a
duration of 14 days before the treated sand were being removed from their respective
mould. Besides the soils being treated with bacteria culture and cementation solution,
another set of control sand specimen was prepared, i.e. sand specimen treated with
cementation solution only.
98
3.2.2 (e) Monitoring methods
During the course of treatment and curing time, the environment where the samples
were placed, was monitored by recording its temperature and relative humidity. The
MICP sand treatment was performed inside a fume hood (LabCraft, BASIX 52). The
biomass concentration and urease activity of the bacteria cultures were also measured
via optical density, bacterial viability, and conductivity methods.
3.2.2 (f) Strength measurement The sand specimens that underwent different
treatment conditions (i.e MICP and sand specimen treated with cementation solution
only) were tested for their respective surface strength and shear strength. The surface
strength measurements of the treated sand were obtained by using a pocket
penetrometer (ELE International, 38-2695) as suggested by Al-Thawadi (2008) and
unconfirmed compression strength (UCS) test in reference to American Society for
Testing and Materials (ASTM) C67-07a for conventional bricks and structural clay tile
test (ASTM, 2007). The penetrometer tests were performed by placing the tip of the
instrument on the surface of the cemented sand. Two different penetrometers with
different reading scales were selected for this test. One of the penetrometers had a
reading scale from 0 to 400 kg/cm2 (0 to 441.229 kPa) while the other had a reading
scale from 0 to 700 psi (0 to 4.826 MPa). The pocket penetrometers were used to
measure the surface strength by pushing the tip of the penetrometer into the soil to a
depth of approximately 0.25 inches and three selected surface regions were tested on
each of the cemented sand. The readings of the loaded weight were recorded when the
samples were completely penetrated (breakage). Test for UCS was performed on an
automatic mortar compression / flexural & concrete flexural machine (NL® Scientific
Instruments Sdn. Bhd., NL 3027 X / 002). All the surfaces of the testing apparatus were
cleaned and the sand specimens were placed on it. The tests were performed until the
sand column reached its failure and maximum stress level.
99
3.2.2 (g) Acid quick test
The acid quick test to confirm the presence of calcite precipitate was performed by
using modified procedures from Cordua (2010). A few amount of precipitates found on
the surface of the sand column after the treatment period were collected, weighed and
kept inside sterile test tubes. Each of the test tubes was filled with 10 mL sterile dH20.
The test tubes containing the precipitates were then added with 2 mL of 10% diluted
HCl. The presence of calcite was visually determined by observing for bubble
formation.
3.2.2 (h) Calcite (CaCO3) content measurement
Calcite content measurements were performed and adapted from methods described by
Weaver et al. (2011) and Bernardi et al. (2014). Samples were obtained from the top,
middle and bottom parts of each cemented sands after strength test. The dry weight of
each sample was taken, then washed with 2M HCl, dried and weighed again after
washed with acid to determine the relative amount of calcite present. The samples were
dried for 3 hr at 90°C in an oven before being weighed. The differences in weight
between the dry sands samples prior and after washing with HCl were divided by the
dry weight after washing to determine the percentage of the calcite precipitation by
weight.
3.2.2 (i) Statistical analysis
For statistical analysis, a standard deviation (SE) for each experimental result was
calculated using Excel Spreadsheets available in the Microsoft Excel (version 2016).
The results obtained from the penetrometer tests were analysed with GraphPad (Quick
Calc) program. The data were subjected to student’s t-test analysis, with statistical
significance taken as p<0.05.
100
3.3 Results
3.3.1. Temperature (oC)
The bacteria were allowed to grow in broth media at temperatures ranging from 20 to
45oC 2 with an interval of 5oC. The optimal incubation temperature supporting urease
activity for the UPB is shown in Figure 3.1. Maximum specific urease activity was
observed at 30oC for isolates NB33 (25.32 mM urea hydrolysed.min-1.OD-1), NB30
(41.98 mM urea hydrolysed.min-1.OD-1) and control strain (23.03 mM urea
hydrolysed.min-1.OD-1), while 25oC was observed to be the maximum specific urease
activity for isolates LPB21 (26.96 mM urea hydrolysed.min-1.OD-1) and NB28 (26.26
mM urea hydrolysed.min-1.OD-1). However, isolates NB30 showed the highest specific
urease activity as 41.98 mM urea hydrolysed.min-1.OD-1 when compared to other
isolates and the control strain. A one-way between groups analysis of variance
(ANOVA) was conducted using Statplus program to compare the effects of different
incubation temperature (ranging from 20 to 45oC) on specific urease activity of
individual ureolytic isolates. The ANOVA for the data on specific urease activity as a
function of variation due to different incubation temperature were statistically
significant for isolate LPB21 (F (5,12) = 12.93, P-value = 1.74E-04); NB33 (F (5,12) =
17.30, P-value = 4.06E-05); and isolate NB30 (F (5,12) = 135.35, P-value = 8.91E-07).
On the other hand, the analysis of variance for control strain (F (5,12) = 5.42, P-value =
0.008) and isolate NB28 (F (5,12) = 9.06, P-value = 9.21E-04 were not statistically
significant. A post hoc analysis using the Tukey’s procedure (α=0.05) further revealed
that the effect of different incubation temperature for isolate LPB21, the mean of 25oC
(M= 29.82; SD= 2.98), was significantly higher than the mean of 35oC (M= 15.97; SD=
2.90), 40oC (M= 8.25; SD= 3.75) and 45oC (M= 19.36; SD= 6.60). The post hoc
analysis revealed that for isolate NB33, the mean of 30oC (M= 25.32; SD= 6.88), was
significantly higher than the mean of 25oC (M= 14.60; SD= 1.33), 40oC (M= 2.72; SD=
0.31) and 45oC (M= 6.57; SD= 0.54). The Tukey’s procedure analysis for isolate NB30
revealed that the mean of 30oC (M= 41.98; SD= 2.88), was significantly higher than the
mean of 20oC (M= 31.69; SD= 2.17), 35oC (M= 22.85; SD= 0.72), 35oC (M= 26.42;
SD= 3.74), 40oC (M= 10.31; SD= 3.87) and 45oC (M= 16.86; SD= 4.56).
101
*
Figure 3.1: The effect of different temperature on urease activity. Cultivation of ureolytic bacteria in NB-medium in 250 mL conical flasks incubated at 20 to 45oC for 24 hr. Vertical error bars indicate standard deviation. The analysis of variance (ANOVA) with Tukey’s procedure was used to compare the variance between different groups with the variability within each of the groups. The level of significance was set at 0.05 (*).
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3.3.2. Initial medium pH
The optimum initial medium pH enhancing the activity of urease was performed by
incubating the bacteria cultures in growth medium with varied pH values ranging from
6.0 to 8.5 with an interval of 0.5 as illustrated in Figure 3.2. Maximum specific urease
activity was observed in the medium of pH 6.5 for isolate NB33 (23.71 mM urea
hydrolysed.min-1.OD-1), whereas isolate LPB21 and control strain showed their
respective maximum specific activity at pH 7.5 with 33.74 and 21.43 mM urea
hydrolysed.min-1.OD-1. Isolates NB30 and NB 28 showed their individual maximum
enzyme activities at pH 8.0 with 30.72 and 34.51 mM urea hydrolysed.min-1.OD-1.
However, isolates NB28 showed the highest specific urease activity as 34.51 mM urea
hydrolysed.min-1.OD-1 when compared to other isolates and the control strain. The
ANOVA analysis showed that there were statistical significances in different initial pH
medium (6.5 to 8.5) for the control strain (F (5,12) = 6.35, P-value = 0.004); isolate
LPB21 (F (5,12) = 39.88, P-value = 4.56E-04); NB33 (F (5,12) = 30.59, P-value = 1.97E-
05); isolate NB30 (F (5,12) = 67.80, P-value = 2.24E-07) and isolate NB28 (F (5,12) =
30.99, P-value = 1.84E-04. The post hoc analysis using the Tukey’s procedure (α=0.05)
further revealed that the effect of different initial pH medium for control strain, the
mean of pH 7.5 (M= 21.43; SD= 0.79), was significantly higher than the mean of pH
6.0 (M= 12.52; SD= 4.23), pH 8.0 (M= 12.20; SD= 0.95) and pH 8.5 (M= 12.7; SD=
1.90), for isolate LPB21, the mean of pH 7.5 (M= 33.74; SD= 3.17), was significantly
higher than the mean of pH 6.0 (M=17.88; SD= 1.93), pH 6.5 (M= 15.06; SD= 1.29),
pH 8.0 (M= 16.95; SD= 2.02) and pH 8.5 (M= 14.91; SD= 1.82). The post hoc analysis
revealed that for isolate NB33, the mean of pH 6.5 (M= 23.71; SD= 0.47), having the
maximum specific urease activity, was significantly higher than the mean of pH 6.0
(M= 9.62; SD= 0.54), pH 7.0 (M= 9.75; SD= 3.91), pH 7.5 (M= 4.22; SD= 0.39), pH 8.0
(M= 10.45; SD= 1.10) and pH 8.5 (M= 10.36; SD= 2.77). The Tukey’s procedure analysis
for isolate NB28 revealed that the mean of pH 8.0 (M= 34.51; SD= 3.98), was
significantly higher than the mean of pH 6.0 (M= 19.68; SD=4.37), pH 6.5 (M= 13.24;
SD= 1.92), pH 7.0 (M= 10.19; SD=2.85), pH 7.5 (M= 11.65; SD= 0.39) and pH 8.5 (M=
15.46; SD= 0.20). The analysis for isolate NB30 revealed that the mean of pH 8.0 (M=
30.92; SD= 1.19, was significantly higher than the mean of pH 6.0 (M= 16.41; SD=1.65),
pH 6.5 (M= 20.37; SD= 1.35), pH 7.0 (M= 14.71; SD=0.40), pH 7.5 (M= 23.65; SD= 1.56)
and pH 8.5 (M= 21.22; SD= 0.51).
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Figure 3.2: The effect of different pH on urease activity. Cultivation of ureolytic bacteria in NB-medium in 250 mL conical flasks incubated for 24 hr. Vertical error bars indicate standard deviation. The analysis of variance (ANOVA) with Tukey’s procedure was used to compare the variance between different groups with the variability within each of the groups. The level of significance was set at 0.05 (*).
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Control LPB21 NB33 NB28 NB30
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3.3.3. Incubation period (hr)
The optimum incubation period for the UPB was performed in growth medium with
varied incubation duration ranging from 24 to 96 hr with an interval of 24 hr as
exemplified in Figure 3.3. Maximum specific urease activity was observed at 24 hr
incubation period for isolates LPB21 (25.98 mM urea hydrolysed.min-1.OD-1), NB33
(27.93 mM urea hydrolysed.min-1.OD-1), isolates NB28 (25.54 mM urea
hydrolysed.min-1.OD-1), NB30 (29.70 mM urea hydrolysed.min-1.OD-1) and control
strain (22.08 mM urea hydrolysed.min-1.OD-1). However, isolates NB30 showed the
highest specific urease activity as 29.70 mM urea hydrolysed.min-1.OD-1 when
compared to other isolates and the control strain. The ANOVA for the data on different
incubation period suggested that there were statistical significances in the effect of
different incubation period for the control strain (F (3,8) = 106.43, P-value = 8.70E-07);
isolate LPB21 (F (3,8) = 20.84, P-value = 3.88E-04); NB33 (F (3,8) = 106.14, P-value =
8.79E-07); isolate NB30 (F (3,8) = 138.04, P-value= 3.15E-04) and isolate NB28 (F (3,8)
= 7.32, P-value = 1.10E-02. Tukey’s procedure (α=0.05) on the effect of different
incubation period showed that for control strain, the mean of 24 hr (M= 22.08; SD=
2.21), was significantly higher than the mean of 48 hr (M= 8.77; SD= 0.92), 72 hr (M=
0.92; SD= 1.17) and 96 hr (M= 4.64; SD= 0.86). The result for isolate LPB21 showed
that the mean of 24 hr (M= 25.98; SD= 3.34), was significantly higher than the mean of
48 hr (M= 6.89; SD= 1.27), 72 hr (M= 9.36; SD= 6.00) and 96 hr (M= 5.34; SD= 1.90).
In addition, the analysis for NB33 that the mean of 24 hr (M= 27.93; SD= 2.03), was
significantly higher than the mean of 48 hr (M= 27.65; SD= 0.70), 72 hr (M=8.86; SD=
2.20) and 96 hr (M= 3.41; SD= 2.00). the Tukey’s test result for Isolate NB28 indicated
that the mean of 24 hr (M= 25.54; SD= 6.09), was significantly higher than the mean of
48 hr (M= 9.24; SD= 1.76), 72 hr (M= 6.69; SD= 3.29) and 96 hr (M= 18.34; SD=
8.47). the test result for Isolate NB30 suggested that the mean of 24 hr (M= 29.70; SD=
2.49), was significantly higher than the mean of 48 hr (M= 7.79; SD= 1.26), 72 hr (M=
65.82; SD= 0.23) and 96 hr (M= 5.86; SD= 1.99).
105
Figure 3.3: The effect of different incubation period on urease activity. Cultivation of ureolytic bacteria in NB-medium in 250 mL conical flasks incubated for 24 hr. Vertical error bars indicate standard deviation. The analysis of variance (ANOVA) with Tukey’s procedure was used to compare the variance between different groups with the variability within each of the groups. The level of significance was set at 0.05 (*).
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Control LPB21 NB33 NB28 NB30
*
*
*
*
*
106
3.3.4. Effect of urea concentration (%)
Experimental results showing the enzyme activity at a varying substrate (urea)
concentration ranging from 2 to 10% with an interval of 2% is presented in Figure 3.4.
Maximum specific urease activity was observed at 6% of urea concentration for isolates
LPB21 (32.36 mM urea hydrolysed.min-1.OD-1) and NB28 (25.98 mM urea
hydrolysed.min-1.OD-1), while 8% urea concentration was observed to show the
maximum specific activity for isolates NB33 (33.95 mM urea hydrolysed.min-1.OD-1),
NB30 (39.21 mM urea hydrolysed.min-1.OD-1) and control strain (24.66 mM urea
hydrolysed.min-1.OD-1). However, isolates NB30 showed the highest specific urease
activity as 39.21 mM urea hydrolysed.min-1.OD-1 when compared to other isolates and
the control strain. The ANOVA for the data on different incubation period suggested
that there were statistical significances in the effect of different urea concentration for
the control strain (F (4,10) = 7.91, P-value = 0.00); isolate LPB21 (F (4,10) = 12.60, P-
value = 6.43E-04); isolate NB30 (F (4,10) = 15.67, P-value= 2.62E-04) and isolate NB28
(F (34,10) = 4.176, P-value = 3.00E-02. On the other hand, there was no statistical
significance. isolate NB33 (F (4,10) = 6.65, P-value = 0.007). Tukey’s procedure
(α=0.05) on the effect of different urea concentration showed that for control strain, the
mean of 8% (M= 24.66; SD= 8.91), was significantly higher than the mean of 2% (M=
5.18; SD= 1.48), 4% (M= 9.26; SD= 2.40) and 10% (M= 11.71; SD= 4.18). The result
for isolate LPB21 showed that the mean of 6% (M= 32.36; SD= 6.62), was significantly
higher than the mean of 2% (M= 6.48; SD= 1.51), 4% (M= 15.96; SD= 4.42), 8% (M=
19.81; SD= 12.69) and 10% (M= 18.88; SD= 5.45). The test result for Isolate NB30
suggested that the mean of 8% (M= 39.21; SD= 9.33), was significantly higher than the
mean of 2% (M= 5.11; SD= 1.18), 4% (M= 18.54; SD= 4.27), 6% (M= 20.09; SD=
1.85) and 10% (M= 21.76; SD= 5.62). The Tukey’s test result for Isolate NB28
indicated that the mean of 6% (M= 25.98; SD= 11.40), was significantly higher than the
mean of 2% (M= 5.92; SD= 2.66).
107
Figure 3.4: The effect of different urea concentration on urease activity. Cultivation of ureolytic bacteria in NB-medium in 250 mL conical flasks incubated for 24 hr. Vertical error bars indicate standard deviation. The analysis of variance (ANOVA) with Tukey’s procedure was used to compare the variance between different groups with the variability within each of the groups. The level of significance was set at 0.05 (*).
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Control LPB21 NB33 NB28 NB30
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108
3.3.5. Biocementation treatment test
The sand used for biocement test was classified as poorly graded medium sand in
accordance with British Standards, BS5930. The percentage of the particle size
distribution of the type of sands selected is shown in Table 3.4. The particle sizes
selected ranged from fine sand (0.075 mm) to fine gravel (4.75 mm). The sand samples
were selected by sieving for designated particles sizes ranges and the sand that could
pass through sieve number 10 (2 mm) were used for the homogeneity. The selected
samples were later oven dried in 105oC overnight and then allowed to cool to room
temperature. The samples were later autoclaved to eliminate any presence of the
microorganism. In order to immobilise bacteria in the columns for use in subsequent
biocement treatment, 80 mL the ureolytic bacteria were premixed with 294.73 g of sand
and 40 mL of 1M urea and 1M CaCl2. The sands were then immersed in columns and
allowed to sit in a fume hood for 8 hr before subsequent addition of bacteria and
cementation solution. Measurements for optical density, viable cells and enzyme
activity of the bacterial cultures were monitored during the treatments (Table 3.5). The
temperature and relative humidity of the environment where the sand columns were
placed ranged between 23 to 29oC and 74 to 85 %.
In Figure.5 (A), there was no visual observation of calcite on the top layer of the
columns during the initial period of immersion of the bacterial culture and cementation.
Whereas, during the third day of inoculation, white precipitates were seen on all
triplicate samples of the columns containing bacterial cultures as shown in Figure 3.5
(B). On the other hand, none of the columns containing the negative control displayed
any visible precipitation on their respective top layers. Upon completion of the
treatment, the sand columns were then allowed to cure for a total duration of 14 days at
room temperature as presented in Figure 3.6. During the curing period, it was observed
that there was an excessive amount of white precipitates on the surfaces of columns
belonging to Consortia, NB33, NB30, LPB21, and control strain. However, column
belonging to NB28 showed it had a lesser amount of precipitates on the surfaces of its
columns when compared to other columns. On the other hand, the columns containing
negative control showed no white precipitates, despite the continual addition of
cementation solution during the subsequent treatment which occurred for 96 hrs.
109
The columns holding the biocemented sands were carefully removed at the end of the
curing period as shown in Figure 3.7. All the biocemented sand samples appeared to
remain intact after removal from the columns. It was also observed that the scouring
pads (Scotch-BriteTM) which were used to prevent any disturbance of the column’s top
surfaces was not very productive during injection of the cementation solution. However,
the hardness of the biocemented sands was not affected. After the columns were fully
removed from the biocemented sands, any other parts of the columns which remained
on the biocemented sand were then carefully removed (Figure 3.8). The sands were then
kept in an incubator at 37oC for 24 hr to minimise the effect of any differences in water
content remaining in the biocemented sands before their mechanical properties were
evaluated.
Table 3.4: Sand grain size characteristics
Characteristics Percentage (%)
Fine sand 6.72
Medium sand 87.95
Coarse sand 3.96
Fine gravel 1.37
110
Table 3.5: Bacteria concentration and urease activity prior to biocement test
Isolate OD600 CFU.mL-1 mM urea hydrolysed.min-1.OD-1
LPB21 0.79 4.8 X 107 16.6
NB30 0.52 4.0 X 107 17.26
NB33 0.69 1.5 X 107 20.96
NB28 0.76 4.1 X 107 23.49
control 0.64 5.0 X 107 13.65
consortia 0.56 4.7 X 107 12.51
OD600 = optical density; CFU.mL-1 = colony forming unit per millilitre; mM urea hydrolysed.min-1.OD-1 = urease activity.
111
Figure 3.5: Treatment of sand column using locally isolated bacteria, consortia, positive and negative controls. [A] setup of sand columns before treated with ureolytic bacteria and cementation solution (Left). [B] sand columns during treatment with bacteria and cementation solution (right). The environment (fume hood) where the MICP treatment occurred had a temperature of 23-29oC and relative humidity of 74-85% during the course of biocement test.
A B
112
Figure 3.6: Sand columns at the end of treatment using ureolytic bacteria and cementation solution. The MICP treatment occurred in a fume hood for a duration of 96 hr with an interval of 12 hr.
Consortia NB28 NB30 NB33 LPB21 Negative control
Positive control
113
Figure 3.7: Treated sand removed from their respective columns. The biocement specimens were allowed to cure for 14 days before being removed from their respective moulds.
Positive control
Consortia LPB21 NB33 NB28 NB30
114
Figure 3.8: Treated sand sample held after a curing period and columns were successfully removed. (A) side view [left], (B) top view [middle] and (C) bottom view [right]. The biocemented specimens were incubated at 37oC for 24 hr to remove any remaining water content before the mechanical properties of the biocement specimens were evaluated.
A C B
115
3.3.6. Soil surface strength
Surface strengths using penetrometer were measured for all the biocemented sand after
curing of the samples. In Figure 3.9, the strength measured for the biocemented sand
treated with different ureolytic bacteria are 582.33 psi for isolate LPB21, 626.67 psi for
isolate NB33, 573.33 psi for isolate NB30, 700 psi for isolate NB28, 533.33 psi for
bacterial consortia and 563.33 for the positive control strain. However, the negative
control was too soft to measure and could not yield any result.
Figure 3.9: Surface strength of the biocemented sand samples. A pocket penetrometer (ELE International, 38-2695) was used to test the surface strength. Vertical error bars indicate standard deviation.
0
100
200
300
400
500
600
700
800
- control + control LPB21 NB33 NB30 NB28 consortia
Str
en
gth
(psi
)
116
The highest strength measured was 700 psi for biocemented sand treated with isolate
NB28 while the lowest strength measured was 533.33 psi for consortia. Among all the
biocemented sand, the sample treated with isolate NB28 reached the maximum reading
of the penetrometer and none of its samples cracked during this surface strength test,
unlike other samples. It was also observed that the sand treated with bacterial cultures
and cementation solutions were slightly more cemented in areas closest to the point of
injection regions. Visual observation after the strength test also indicated that there were
much more precipitates on the surface of the biocemented sands than other areas.
Table 3.6: t-test results comparing the strength (psi) differences between the biocemented sands. (N=3; df=2)
Isolate ID M SD SE P-value t P <*
control 563.33 42.10 nil nil nil nil
LPB21 582.33 67.35 21.221 0.4651 0.8953 -
NB33 626.67 99.81 57.097 0.3829 1.1092 -
NB30 573.33 6.51 26.312 0.7405 0.3801 -
NB28 700.00 0.00 24.306 0.0302 5.6228 +
consortia 533.33 116.93 76.374 0.7324 0.3928 -
(N) number of sample size; (df) degree of freedom; (M) mean; (SE) standard error; (SD) standard deviation; (P-value) calculated probability; (t) test statistic; (+) significant; (-) not significant ;(*<) P-value is significant at 0.05 level.
117
The independent t-test was conducted to compare the surface strength measurement
obtained from biocemented sands for the local isolates and bacterial consortia against
those of the control strain. As illustrated in Table 3.6, there was a significant difference
between the strength of biocemented sand treated with isolate NB28 (M= 700.00; SD=
0.00) against that of the control strain (M= 563.33; SD= 42.10). However, there were no
noticeable significant different between the test results when compared against that of
the control.
3.3.7. Compressive strength
None of the sands treated with the negative control was tested for strength measurement
using the automatic mortar compression machine as the sands were too soft and not
amenable to unconfined compressive testing. During the UCS testing, it was visually
observed that the failure points for all biocemented sands started at their respective
bottom layers. The results from Table 3.7 indicate that the biocemented sands with the
highest test were treated with isolate NB28 (0.219 N/mm2), sustaining a force of 1.020
kN, while sands treated with the lowest strength was treated with the control strain
(0.143 N/mm2), sustain a force of 0.697 kN.
Table 3.7: Unconfined compressive strength (UCS) of the treated sands
Bacteria ID
UCS test
Condition of cemented sand
Force (kN)
Pressure (N/mm2)
- control - - - + control + 0.647 0.143 LPB21 + 0.697 0.152
NB33 + 0.833 0.176
NB30 + 0.647 0.143
NB28 + 1.020 0.219
consortia + 0.623 0.147
(-) the column was not cemented; it was extremely soft and unable to be measured. (+) the cemented column was broken when the maximum strength was applied.
118
The independent t-test was also conducted to compare the UCS test results obtained
from biocemented sands for the local isolates and bacterial consortia against those of the
control strain. The results in Table 3.8 showed that out of all the strength results from
biocemented sands of the isolates and bacterial consortia, there were noticeable
significant differences between the strength results for biocemented sands treated with
isolates LPB21 (M= 0.152; SD= 0.006), NB33 (M=0.176; SD= 0.025) and NB28 (M=
0.219; SD= 0.013) against the control strain (M= 0.143; SD= 0.006).
Table 3.8: t-test results comparing the unconfined compressive strength (UCS) differences between the biocemented sands (N=3; df=2)
Isolate ID M SD SE P-
value t P <*
control 0.143 0.006 nil nil nil nil
LPB21 0.152 0.006 0.005 0.011 9.526 +
NB33 0.176 0.025 0.012 0.072 3.534 +
NB30 0.143 0.002 0.001 0.840 0.229 -
NB28 0.219 0.013 0.000 0.004 16.174 +
consortia 0.147 0.009 0.005 0.374 1.134 -
(N) number of sample size; (df) degree of freedom; (M) mean; (SE) standard error; (SD) standard deviation; (P-value) calculated probability; (t) test statistic; (+) significant; (-) not significant;(*<) P-value is significant at 0.05 level.
119
3.3.8. Calcite confirmation
Figure 3.10: Confirming calcite precipitates. The calcite contents precipitates found on the surfaces of the biocement moulds were tested using quick acid test. (A) before addition of HCl [left]. (B) after addition of HCl [right]. The white precipitates which were seen on the top layer of sand columns were presumed to be calcite precipitates. In order to confirm
this precipitates that were induced by the ureolytic bacteria during the treatment period, some amount of the excess precipitates was
taken and kept in sterile test tubes as shown in Figure 3.10 (A). After the addition of 10% HCl solution, the continual formation of
bubbles was visually observed. The addition of acid (HC)l onto the calcite resulted in bubbles of carbon dioxide gas to be released as
indicated in Figure 3.10 (B). This bubble formation signals the presence of calcite.
A B
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3.3.9. Calcite content Determination
Figure 3.11: Comparison of the relative quantity of calcites in the biocemented sands. The calcite contents were dried for 3 hr at 90°C in an oven before being weighed.
0
2
4
6
8
10
12C
alc
ite
we
igh
t, %
(w
/w
)top middle bottom
+ control LPB21 ConsortiaNB33NB28NB30 - control
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Table 3.9: Summary of calcite content and compressive strength of selected isolates and consortia
Isolate ID
Weight of calcite, % (w/w) Strength conversion
(MPa)* Top Middle Bottom
- control 0.00 0.00 0.00 0.00
+ control 5.28 3.95 3.19 3.88
LPB21 9.20 2.01 5.59 4.02
NB33 5.86 4.65 7.19 4.32
NB30 6.12 3.09 6.69 3.95
NB28 10.08 7.14 7.09 4.83
consortia 4.70 1.72 1.73 3.68
(*) converted strength (psi) into MPa by knowing that 1 psi = 0.00689476 MPa. The content of the calcite precipitated in the sand specimens as shown in Figure 3.11
were determined by using acid wash method. The average calcite content of the
biocemented sands was determined from samples collected at the top, middle and
bottoms sections of the treated sand as presented in Table 3.9. The top layers of the
biocemented sands treated with the control strain (5.28%), isolates LPB21 (9.20%),
NB28 (10.08%) and bacterial consortia (4.70%) had the highest average calcite
contents, while the sand samples treated with isolates NB33 (5.86%) and NB30 (6.12%)
showed that the bottom layer had the highest average calcite content. The differences in
calcite content on top–middle layers and top-bottom layers for the biocemented sand
samples treated with control strain (3.88 MPa), isolates LPB21 (4.02 MPa), NB28 (4.83
122
MPa) and bacterial consortia (3.68 MPa) were 1.33% and 2.09%; 7.19% and 3.61%;
2.92% and 2.99%; 2.98% and 2.97%, respectively. On the other hand, the differences in
calcite content on bottom–middle layers and bottom-top layers for the biocemented sand
samples treated with NB33 and NB30 were 1.33% and 2.54%; 0.57% and 3.60%,
respectively.
There was not any homogeneity of calcite contents within any layer of the biocemented
sand samples. However, results in Table 3.9 indicated that they were similar calcite
contents between the middle and bottom layers of biocemented sands treated with the
control strain, isolate NB28 and bacterial consortia. This result suggests there was
reasonable precipitation uniformity from middle to bottom layers of these
aforementioned sand samples. In Figure 3.11, among all the biocemented sands, the
highest average calcite content for the top, middle and bottom were determined to be
10.08% (NB28), 7.19% (NB33) and 4.83% (NB28), respectively.
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3.4 Discussion A study by Hamzah et al. (2012) indicated that physical parameters such as initial
medium pH and incubation temperature play important roles which promote microbial
biomass production. Among all parameters to be studied, the temperature was
considered the most important factor and it is a critical parameter which needs to be
controlled as it usually varies from one organism to another (Delilie et al., 2004, Kumar
and Takagi, 1999). Hence, this physical parameter was first selected among others to
determine the optimum temperature which can facilitate an appropriate production of
urease for the ureolytic bacteria. The result presented in Figure 3.1 suggests that 25oC
and 30oC are the optimum temperature for the selected ureolytic bacteria and the control
strain which produced the most favourable activity of urease enzyme. These
temperatures correspond to the standard temperature of Kuching, Malaysia. It was
perceptible that these bacterial cultures would yield substantial urease activities at a
temperature between 20 to 35oC.
Earlier studies on optimisation of temperatures on urea hydrolysis and calcium
precipitation showed the preferred temperature for bacterial growth was between 30 to
35oC (Helmi et al., 2016, Seshabala and Mukkanti, 2013). Higher temperature
conditions did not have a favourable outcome of enzyme activity when the UPB were
cultured at 40oC and 45oC. However, it was noteworthy that isolate LPB21 showed a
reasonable enzyme activity when cultured at 45oC. Microbial urease strongly relies on
temperature as the rate of urea hydrolysis changes with changes in temperature due to
kinetic energy inducing the collisions between an enzyme and its substrate, and is
capable of stimulating modifying to the cellular membrane of bacteria. (Akgöl et al.,
2002, Rahman et al., 2005). It is well known that proteins conformation changes or
degrades at a higher temperature as the structure of the enzyme may become altered,
thus making the enzyme’s catalytic become eventually destroyed (Seshabala and
Mukkanti, 2013, Abusham et al., 2009).
The pH of a microorganism’s growth medium plays an important role by inducing
morphological changes in microbes, enzyme secretion and affecting the microbe’s
stability in the growth medium (Sethi and Gupta, 2014). Hence, the effect of the initial
medium pH on urease activity for the ureolytic bacteria was studied.
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The results illustrated in Figure.2 indicate that pH 6.5, 7.5 and 8.0 are the optimum pH
values for the selected ureolytic bacteria and the control strain. pH 7.5 and 8.0 produced
the most noticeable activity of urease enzyme when compared to others. On the other
hand, at an acidic level of pH 6.0 and an alkaline level of 8.5, the activities of urease for
the ureolytic bacteria were found to be low and unfavourable. Urea hydrolysis in a
growth medium is expected to lead to an increase in the pH value due to the production
of ammonium (Gat et al., 2014). Previous studies have shown that urease enzyme is
more active at alkaline pH (Prah et al., 2011, Anne et al., 2010, Stocks-Fischer et al.,
1999).
Urea hydrolysis at different alkaline pH was investigated and confirmed by Helmi et al.
(2016)for the previous research outcomes. The primary step of the chemical reaction is
the hydrolysis of urease enzyme as it leads to an induction of calcium carbonate (Millo
et al., 2012, Okwadha and Li, 2010). A study by Helmi et al. (2016) showed that the
hydrolysis of urease enzyme by Bacillus licheniformis and formation of ammonium
increases the pH value up to 8.0, sufficient enough to induce calcite precipitation. A
study by Gat et al. (2014) suggested that the pH values of Sporosarcina pasteurii
(DSM 33) for urea hydrolysis during incubation is at pH 7.39 after being incubation at
28 hr. A study on variation of solution pH for purified urease enzyme from jack bean
meal and microbial urease from Bacillus megaterium by Jiang et al. (2016) showed that
regardless of oxic or anoxic conditions, pH values increases sharply within the initial 1
hr which is ascribed to immediate hydrolysis of urea. Ferris et al. (2004) found that
bacteria cell growth during ureolysis process in anoxic conditions contributes to extra
acidic substances which can reduce the pH values and increase electric conductivity
values.
The incubation period is an essential parameter for enzyme production (Gautam et al.,
2011). The result of the optimum incubation period is presented in Figure 3.3 which
shows that maximum urease activity for all the bacterial isolates and control strain was
found to be at 24 hr incubation period. On the other hand, at an incubation period of 48
to 96 hr, the enzyme activity decreased and was not favourable. This decline is as a
result of saturation of actives sites of the microbial enzyme by the substrate molecules
which is not longer to be involved in the breakdown of it (Fisher, 2001).
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A report by Gat et al. (2014) showed that Sporosarcina pasteurii (DSM 33) grown for
10 days with agitation (100 rpm) at 30oC using nutrient broth supplemented with 2%
w/v urea (333 mM) showed a steady increase in bacterial and enzyme activity its first
80 hr but remained at lag phase at 17 hr of incubation which could be as a result of low
concentration of urea supplemented. The decline in urease activity shown in Figure 3.3
stands in agreements with the finding of previous studies by (Ferris et al., 2004, Dupraz
et al., 2009) that Sporosarcina pasteurii can still hydrolyze urea in the absence of
sufficient organic carbon source, however the number of viable cells and enzyme
activity is likely to decrease significantly with time under these conditions. Another
study by (Achal et al., 2010a) on biocalcification by Sporosarcina pasteurii (NCIM
2477) using corn steep liquor as a nutrient source has grown at 37 oC for a duration of
160 hr with corn steep liquor, nutrient broth and yeast extract, each supplemented with
2% urea. This resulted showed that urease activity and biomass from corn steep liquor
medium was significantly higher than those observed in nutrient broth and yeast extract.
Thus suggesting corn steep liquor is noteworthy a preferred growth medium for enzyme
production and a low-cost nutrient compared to the aforementioned nutrients.
Bacteria needs a source of nitrogen to support their maximal growth because nitrogen is
a key building block of protein, enzymes and nucleic acids (Hamzah et al., 2013).
Hence, this parameter was also studied to determine the optimum urea concentration
(w/v %) since ureolytic bacteria primary requires urea substrate as their source of
nitrogen. The result illustrated in Figure 3.4 suggests that 6% and 8% are the optimum
urea concentration for the selected ureolytic bacteria and the control strain which
produced the most favourable activity of urease enzyme. A study by Mortensen et al.
(2011) on the subject of environmental factors such as ammonium ions and free oxygen
affecting urease activity. Their results showed that since ureolytic activity depends on
the available substrate (urea) and concentration of urease enzyme, production of
ammonium during urea hydrolysis would not alter urease activity. Ammonia, a nitrogen
source for most bacteria can be detrimental or toxic when present in high concentration
due to cytotoxic effect (Hess et al., 2006). On the other hand, a high concentration of
ammonia can be advantageous to particular ureolytic bacteria such as Sporosarcina
pasteurii as it can assist their ATP generation but with an increasing concentration of
urea, a decrease in biomass and specific urease activity can be met (Cheng and Cord-
Ruwisch, 2013).
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In addition, a study by Cuzman et al. (2015b) suggested that fertilizer urea served as an
effective and cost urea substrate when compared to expensive pure grade urea (Sigma-
Aldrich) as it showed similar behaviour in the presence of commercial urease but there
were no significant differences was observed as regards to microbial urease activity.
The maximum urease ureolytic rate measured for pure grade urea was 0.0892 mS.cm-
1.min-1 and 0.0866 mS.cm-1.min-1 for fertilizer urea. Hence suggesting the use of
fertiliser urea could reduce cost and reduce the environmental impact of urea production
from the release of ammonia by ureolytic bacteria, possible urea substitute such as
poultry manure could be considered (Cuzman et al., 2015b).
An in situ laboratory biocement experiment was conducted in columns containing sand
samples. Three column samples were prepared for each bacterial culture and
cementation solution treatments. The experiment was made in triplicates as presented in
Figure 3.5 to check for repeatability and to quantify the changes in the sand properties
statistically. The negative control contained only the cementation solution and was used
to treat its respective sand columns. This was done to rule out the possibility that
precipitates found in the sand columns were only as results microbial urea hydrolysis
and not any other process. However, during the premixing of sand, bacteria, urea and
calcium chloride solution, the pungent smell was perceived indicating the breakdown of
urea and release of ammonium gas. This treatment method was applied to ensure
bacteria culture and premix urea and calcium chloride solution attach at particle
contacts within the permeable sand matrix. The white precipitates on top layers of the
biocemented sand shown in Figure 3.6 and Figure 3.7 were also reported by Zhao et al.
(2014) and Chu et al. (2012) which Indicates the presence of nucleation sites for MICP
as a result of addition of more bacterial solution in order to promote more urease
enzyme. The precipitates were confirmed to be calcite as shown in Figure 3.10 by
addition diluted HCl onto the precipitates. To confirm the presence of calcite, a quick
acid test can be used by adding drops of HCl on calcite mineral, the reactions allow
bubbles formation and a vigorous effervescence which last for some minutes or seconds
Cordua (2010).
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The results of the biocemented sands showed that all the samples were found to be
tightly packed except the negative control. The use of penetrometer to test for surface
strength showed that samples treated with NB28 (Figure 3.9) were found to be the most
compact sample, requiring more force to be applied to break the samples. Calcite
precipitation was detected over the entire length of the treated sand samples, which
indicated that the ureolytic bacteria and reactants were present at all locations of the
samples (Whiffin et al., 2007). However, it was observed that the precipitation of the
calcite contents in the biocemented sand samples was not homogeneous and more
calcite contents were found in the top layers of the samples (Figure 3.11). In other
biocementation studies, formation of calcite contents dominantly around surfaces of
their respective columns and with no homogeneity have been reported (Rowshanbakht
et al., 2016, Dhami et al., 2016, Neupane et al., 2015, Feng and Montoya, 2015, Zhao et
al., 2014, Achal et al., 2009b, Achal et al., 2009a). On the other hand, in other similar
studies, the formation of calcite precipitates on the layer of their respective columns was
not mentioned (Harkes et al., 2010, Whiffin et al., 2007, van Paassen et al., 2010).
The reason there was predominantly more calcite formation at the top layers of the sand
samples is mainly that Sporosarcina pasteurii is a facultative anaerobic bacterium,
which grows at a higher rate in the environment containing oxygen and consequently
leading to higher rates of calcites precipitated around the top surface areas (Whiffin et
al., 2007). Additionally, the influence of microbial cementation on granular behaviour is
dependent on the ability of the bacteria to move freely throughout the pore spaces of the
sand and on sufficient particle-particle contact per unit volumes at which cementation
will occur. Hence, this quicker formation of calcite precipitation at injection points of
the bacteria and cementation solution prevents more precipitates from flowing freely
downward the columns and causing a non-uniformity of calcite precipitates (Dhami et
al., 2016, Achal et al., 2009b, Achal et al., 2009a). According to ATSM (D2166-00)
standards, to test for unconfirmed compressive strength of cohesive soil, specimen sizes
are required to have a minimum diameter of 30 mm with a length of one-tenth of the
specimen diameter or 72 mm with a length of one-sixth of the specimen diameter.
However, the diameter and length of column samples used in this experiment were75
mm and length of 49 mm, hence it did not follow the standard of ATSM.
128
The biocement test was primary designed to test the ability of the locally isolated
ureolytic bacteria in treating loose soils by filling their pores and testing the surface
strength of the samples without being in their respective columns. However, the UCS
test was later performed to get an estimated unconfined compressive strength and
understand what state of force will the samples reach their respective failed points.
The results presented in Table 3.7, Table 3.9 and Figure 3.9 suggested that there was a
correlation between the surface strength using penetrometer test, calcite contents, and
compressive strength. Park et al. (2014) stated that increase in calcite precipitation due
to increasing microbial injections may weaken the existing cementation in the cemented
soil. This was supported by Park et al. (2010) and Ghosh et al. (2009) suggesting that
microbes do not always increase the biocement strength, instead, might decrease the
strength. However, other studies on MICP reported an increase in strength as a result of
calcite precipitation with the addition of more bacteria (van Paassen et al., 2010,
DeJong et al., 2006, Ramachandran et al., 2001). Paassen (2009) reported that bacterial
urease activity dropped after 20 days of injected but improved after another injection
batch of bacterial culture was added to the column. It was suggested that the enzyme
activity could have been as a result of a hydraulic construct such as the bacteria being
trapped inside the pores of the sands during precipitation and interruption of chemical
transport which could prevent the flow of required nutrients for growth and more calcite
precipitation from reaching the bacteria. Hence, it was necessary to maintain sufficient
amount of repeated additional bacterial culture to the columns so as to prevent possible
accumulation of metabolic waste which could result in a decrease of urease activity,
cell death and poor precipitation (Stocks-Fischer et al., 1999).
Synergistic microbial communities are abundant in nature, with metabolic capabilities
and robustness. This has inspired fast-growing interest in engineering synthetic
microbial consortia for biotechnology development (Minty et al., 2013). Hence, a
bacterial consortium was included in the biocement test as a comparison with the
individual isolates and control strain and to determine which will provide the best
strength and calcite content. The results in Table 3.9 indicated that the ureolytic bacteria
performed better individually and the consortia showed the lowest performance in terms
of strength and calcite content.
129
The soils samples containing the negative control were unable to bind together, thus, the
solution media flowed freely and the samples fell apart when the columns holding the
samples were removed. This observation was also reported by (Kavazanjian and
Hamdan, 2015). However (Whiffin et al., 2007) reported having the strength of 167 kPa
for samples untreated with bacteria but it was not reported if the samples used were
autoclaved to prevent false results. To rule out the possibility of having false
precipitations such as chemically induced calcite precipitation on any of the samples,
the sand samples used were autoclaved before treated with the ureolytic bacteria. This
suggestion was adopted from Burbank et al. (2011). Their finding showed that the
autoclaved soil samples not treated with ureolytic bacteria did not precipitate calcite
when observed by X-ray powder diffraction analysis. Li et al. (2011a) suggested the use
of soybean meal as it can serve as an alternative source of nutrients to support bacterial
growth and increase urease activity for enhanced supply of calcite precipitation.
The biocement application of bacterial consortia that is well adapted to the
environmental conditions in Sarawak was studied. The advantage of a mixed bacterial
consortium which comprised of Sporosarcina pasteurii LPB21 (SUTS), Sporosarcina
pasteurii NB30 (SUTS), Sporosarcina pasteurii NB28 (SUTS) and Sporosarcina
pasteurii NB33 (SUTS) had the lowest urease activity (12.51 mM urea hydrolysed.min-
1.OD-1) when compared to the single isolates. The low production of urease was seen to
have affected the potential of inducing a high amount of calcium carbonate precipitates
during the in vitro biocement test. The bacterial consortia might have performed better
than the single isolates as a result of a low number of bacteria cell not culture was low.
One of the factors which may have affected the biomass synergy of the bacterial
consortia could be attributed to insufficient oxygen in the microenvironment (Hamzah
et al., 2013). The oxygen demand of an aerobic bacterial culture is influenced by the
bacterial concentration and growth rate (Elsworth et al., 1957). If the oxygen demand by
the bacteria consortia exceeded the oxygen supply, the biomass production will be
affection. Another factor which could affect the ability of the constructed bacterial
consortia from having a performance during the biocement test is the urea (nitrogen
source) concentration.
130
The optimised urea concentration of the four bacterial isolates which were used for the
design of the bacterial consortia were seen to be between 6-8%. sufficient urea can
promote necessary biomass production and urease activity required for binding of soil
particles. The determination of urea concentration for bacterial consortia might help to
enhance the synergistic effectiveness. Findings from this study suggest that the use of
single urease-producing bacteria were more effective than the designed bacterial
consortia for the in vitro biocementation.
131
3.5 Conclusion A series of laboratory test were carried out to determine the cultural conditions required
in improving urease activities for the ureolytic bacteria. It was observed that when
incubated these conditions: at 25 to 30oC; pH 6.5 to 8.0; incubation period at 24 hr; and
urea concentration of 6 to 8%, maximum specific urease activities for the selected
ureolytic bacteria isolates and control strain were obtained. The in situ laboratory
biocement test proved that cement precipitation was observed in all sand columns
except columns treated with negative control. The results presented in this chapter
demonstrated that biocementation using the selected ureolytic bacteria can significantly
improve the engineering properties of poorly graded soils. However, the efficiency of
the MICP process in improving the soil strength varied among the samples which were
treated with different isolates, the bacteria consortia, and the control strain. The results
also showed there was higher cementation level at positions close to the injection points
and more calcite contents were obtained from the top layers of the biocemented sand.
Based on the surface strength using penetrometer test and compressive strength using
UCS test, samples treated with isolate LPB21 and isolate NB28 showed significant
strengths when compared to other isolates, consortia, and the control strain. However,
the rest isolates showed similar performance with the control strain. This comparative
study has shown that the ureolytic bacteria isolated from limestone samples of Sarawak
are capable of improving a poorly graded soil when compared to the control strain. The
results in this chapter also give a basis for further study with large scale fermentation of
the ureolytic bacterial (LPB21, NB30, NB28, and NB33) for application in civil and
geotechnical engineering.
Chapter
4 GENERAL CONCLUSIONS AND RECOMMENDATIONS
132
4.1 General Conclusion 4.1.1. Aim of the thesis
This thesis presents data from our work exploring the biodiversity of Sarawak, Malaysia
for urease producing bacteria. Numerous researchers have selected Sporosarcina
pasteurii as their ideal ureolytic bacteria for biocement applications because of its high
urease activity and inability to cause harmful diseases (Wei et al., 2015, Cuzman et al.,
2015b, Kang et al., 2014b). In addition, many studies have reported purchasing different
type strains of Sporosarcina pasteurii from microorganism culture collection centres
such as National collection of industrial and marine bacteria (Sidik et al., 2014, Raut et
al., 2014, Hammad et al., 2013b), German collection of microorganisms and cell
cultures (Gat et al., 2014, Harkes et al., 2010, Paassen et al., 2009) and American type
culture collection (Zhang et al., 2015, Bundur et al., 2015, Onal and Frigi, 2014) for
their respective MICP investigative studies and field applications.
Some common Sporosarcina pasteurii type stains which have been reported as ideal
MICP agents are ATCC 6453, ATCC 11859, DSMZ 33, MTCC 1761, ATCC 14581,
NCIMB 8841, NCIMB 8221 and NCIM 2477 (Sidik et al., 2014, Abo-El-Enein et al.,
2013, Lee et al., 2012). These strains were selected as bio-agents of biocement
applications because they are able to survive in high alkaline (above pH 8.5)
environments and also a high concentration of calcium ions (i.e. 0.75 M) (Stabnikov
and Ivanov, 2016). On the other hand, apart from the use of Sporosarcina pasteurii for
MICP process, some species of the genus of Bacillus capable of producing urease and
biocement capabilities are Bacillus cereus, Bacillus sphaericus, Bacillus subtilis,
Bacillus licheniformis, Bacillus Cohnii, and Bacillus lentus (da Silva et al., 2015,
Vahabi et al., 2014, Sierra-Beltran et al., 2014).
The current exploitations of isolating alternative ureolytic bacteria species for effective
biocement applications are still very limited (Soon et al., 2014, Dhami et al., 2013d,
Cheng and Cord-Ruwisch, 2012). This drawback spurred the interest of studying
Sarawak’s environment for the possibility to isolate highly active urease-producing
bacteria. Prior to this study, there were no records of any urease producing bacteria
capable of showing high specific urease activity in Sarawak when compared with
representative microbial urease strain, Sporosarcina pasteurii (DSM33).
133
This thesis is thus, a first report on the strategy to screen and characterise urease
producing bacteria isolated from limestone cave samples of Sarawak. This study also
presents the effects of cultural conditions on urease activity for the local ureolytic
isolates, and an evaluation of biocementation potentials in small scale test. This chapter
summarises the findings of chapters two and three presented in this thesis. This chapter
also offers recommendations for imminent works in this study area.
4.1.2. Limestone area as source of ureolytic bacteria
Caves are extreme environments which are unique, unexploited and poorly studied,
making it an ideal region to screen for microorganism capable of producing novel
bioactive compounds (Gabriel and Northup, 2013). Various environmental factors in
caves such as limited sunlight and nutrients influences microorganisms to adapt to such
location, forcing them to search for other carbon and nitrogen sources (Cheeptham,
2013, de Lurdes N. Enes Dapkevicius, 2013, Northup et al., 2011). Energy is essential
for the production of bacterial metabolites, inorganic compounds such as nitrogen,
sulphur and carbon dioxide (Northup et al., 2011). The ability for microorganisms to
survive in as caves, suggests that they are ideal environments that provide diverse
habitats for microbial survival (Alnahdi, 2014). Limestone is a type of carbonate
mineral composed of pure calcite or pure calcite (CaCO3) or dolomite (CaMg(CO3)2) or
a mixture of both (James et al., 2008). Limestone formation can be discovered regions
such as seawater, caves, and coral reefs, these formations usually contain high
concentrations of calcium and bicarbonate ions (James et al., 2008). The possibility of
certain cave microorganisms capable of inducing calcite precipitates on their cell’s
surfaces contributing to the formation of limestone caves initiated the concept of
exploring and isolating indigenous microbial species from extreme environments with
highly active urease producing bacteria. Hence, Fairy and Wind limestone caves of
Sarawak were selected to explore the possibility to attain novel ureolytic bacteria which
are indigenous to Sarawak’s environments.
134
4.1.3. Enrichment culture and isolation
Enrichment and isolation methods were used in this thesis to target microorganisms
capable of surviving on urea as a main source of nitrogen. Nutrient broth (HiMedia),
Tryptic soy broth (Merck), Lactose peptone broth (BD Difco™), Luria broth (HiMedia)
and Brain heart infusion broth (Oxoid) growth medium were used in this thesis because
they contain various compositions of ingredients suitable for growing a substantial
number of bacteria. Each of the enrichment cultures was supplemented with Sodium
acetate (8.2 g.L-1), Ammonium sulphate (10.0 g.L-1) and Urea (40 g.L-1). This selective
enrichment technique facilitated in successfully isolating ninety urea degrading bacteria.
Urea (CO(NH2)2) is a naturally-occurring form of nitrogen present in both aquatic and
terrestrial environments (Fisher, 2014). Urea is part of dissolved organic nitrogen pool
which microorganisms depend on as a valuable nitrogen substrate for survival (Berman
and Bronk, 2003, Altman and Paerl, 2012, Tyler et al., 2003). Urease enzyme raises the
pH of a bacteria’s environment, by allowing it to depend solely on urea as nitrogen
source (Williams et al., 1996). For the purpose of targeting highly active ureolytic
bacteria, 6% urea substrate was selected and used in enrichment culturing of the
samples collected from FCNR and WCNR. Upon successful isolation of urea degrading
bacteria, 6% urea substrate was subsequently used to confirm the ability of the isolated
microorganisms to degrade high urea concentration when subcultured on nutrient agar.
4.1.4. Screening and identification
Christensen’s medium (urea agar base) was used to screen for urease producing
bacteria. Various studies have suggested the use of Christensen’s medium to screen and
detect urease-producing bacteria (Hammad et al., 2013b, Elmanama and Alhour, 2013,
Dhami et al., 2013d). Out of the ninety isolates, thirty-one isolates were capable of
producing urease. Molecular identification showed that these urease producing bacteria
belonged to Sporosarcina, Pseudogracilibacillus, Staphylococcus and Bacillus groups
with 91 to 99% sequence similarity to existing sequences of their respective closest
bacterial species in the GenBank database. However, the majority of the urease-
producing bacteria were similar to Sporosarcina pasteurii when compared to the 16S
rRNA sequencing data in NCBI nucleotide BLAST database.
135
The majority of the bacterial isolates were Gram-positive bacteria while only three of
the isolates (A63, B53, and A62) were Gram-negative bacteria. Gram-positive bacteria
lack an outer cell membrane but are surrounded by layers of thick peptidoglycan cell
wall, while Gram-negative bacteria has a thin peptidoglycan cell wall with an outer
membrane containing lipopolysaccharide (Andrew, 2013, Silhavy et al., 2010). This
membrane in Gram-negative bacteria is responsible for many antigenic properties of
Gram-negative bacterial species, making the majority of species of Gram-negative
bacteria be pathogenic. Hence, for MICP process, Gram-positive bacteria are often
preferred (Wong, 2015).
4.1.5. Measurement of urease activity
Conductivity method was used to determine urease activity of the urease producing
bacteria. This method was suggested as the most preferred urease enzyme assay because
it is easy to use and inexpensive (Al-Thawadi, 2008, Whiffin, 2004). The changes in
conductivity of bacterial-urea solutions were monitored for a duration of 5 min at 25◦C
and the respective conductivity values were measured using a Walk LAB conductivity
pro meter, Trans Instruments COMPRO. The conductivity variation rate (mS.cm-1.min-1)
is obtained from the gradient of the graph. The urea hydrolysis rate for the urease activity
conversion was determined by (Whiffin, 2004) as described in equation 1.23, while the
Specific urease activity (mM urea hydrolysed.min-1.OD-1) which reflects the urease
catalytic abilities of the urea hydrolysis (Zhao et al., 2014) was derived by dividing the
urease activity (mM urea hydrolysed.min-1) by the bacterial biomass (OD600). The
specific urease activity was also determined by (Whiffin, 2004) as described in equation
1.24. The results determined from the enzyme assay showed that isolates NB33 (19.975
mM urea hydrolysed.min-1.OD-1), LPB21 (23.968 mM urea hydrolysed.min-1.OD-1),
NB28 (19.275 mM urea hydrolysed.min-1.OD-1), and NB30 (20.091 mM urea
hydrolysed.min-1.OD-1) had the highest specific urease activities when compared to other
isolates and the representative strain (17.751 mM urea hydrolysed.min-1.OD-1). Due to
their high enzyme activities, the aforementioned isolates were selected and used for rest
subsequent experiments in this thesis.
136
4.1.6. Biocementation competency of local isolates
The effectiveness of MICP treatment on poorly graded sand specimens used in this
thesis was performed by using locally isolated ureolytic bacteria. The implementation of
MICP treatment using these isolates was compared with sand specimens treated with a
bacterial consortium and a representative strain (Sporosarcina pasteurii). The
biocement test results showed the MICP agents were able to induce calcite precipitates
capable of filling the pore particles of the poorly graded sand samples, hence improving
the strength and stiffness of loose sands. The surface strength and compressive strength
test results showed the local ureolytic bacteria had comparative strengths to that of the
representative strain. However, the highest surface and compressive strength results
obtained were 700 psi and 0.231 N/mm2, which was from samples treated with NB28
bacterial culture. The findings from the calcite content for all the samples treated with
microbes showed the distribution of the calcite contents were not uniform. The top layer
of the specimens contained the highest calcite content. The results in this thesis showed
that biocementation treatment using locally isolated ureolytic bacteria were successfully
able to improve the mechanical properties of poorly graded sands comparable with the
representative strain utilised in this thesis.
4.2 Future Directions and Recommendations The findings in this thesis suggest that the isolated ureolytic bacteria (NB28, LPB21,
NB33, and NB30) have the potential to be used as alternative microbial MICP agents
for biocement applications. Future work involving these four isolates may involve
large-scale bacterial production using computerised bioreactor. The large scale bacterial
production can be utilised for MICP treatment involving field application. The use of
alternative growth medium as a carbon source for large-scale bacteria production can be
studied in order to minimise the cost of purchasing nutrient source. A study on how this
alternative medium enhances the production of bacterial growth, urease activity and
calcite precipitation can be investigated. A comparison between the lab grade urea
substrate and industrial grade urea or alternative nitrogen sources can also be
investigated for future work. This will also be essential for field applications and
reduction of cost for MICP treatments. The effect of cementation reagents MICP
process was not investigated in this study.
137
The urea and calcium chloride concentration used were 1 M (w/v%), this might have
limited the precipitation of calcite during biocement treatment. Hence, the effect of the
concentration of these reagents should be further investigated. Additionally, alternative
calcium ions such as eggshells, pearl shells, snail shells, calcium sulphate or calcium
acetate can also be examined in comparison to calcium chloride and determine which
might result in the most amount of calcite content.
The evidence of microbial involvement in calcite precipitation has brought a revolution
in the discipline of biotechnology, geotechnical and civil engineering. However, some
MICP applications involving some expensive materials have resulted in the successful
commercialization of biocement, but it has been at a high cost (Dhami et al., 2013a).
Hence, it is recommended to explore the use of cheap industrial by-products such as fly
ash which can serve a supplementary cementitious material and capable of significantly
reduce cement and concrete carbon footprint (Thomas, 2007). The investigation of
using the mixture of fly ash, sand, and locally isolated ureolytic bacteria could improve
the existing findings which support the use of MICP process as an alternative
economically friendly construction material.
Screening of urease-producing bacteria was from Fairy and Wind Caves Reserves from
Sarawak. A collection of speleothem, calcareous and Guano (from the bat) samples
from other cave regions such as Gunung Mulu National Park, Deer Cave, Lang Cave
and Clearwater Cave situated in other parts of Sarawak should be performed. Screening
of novel highly active ureolytic isolates of various genuses might be attainable. The
discovery of the ureolytic bacteria isolate mentioned in this thesis could be applied in
other MICP applications to solve problems relating to environmental biotechnology,
civil engineering, and geotechnical engineering. The isolated ureolytic bacteria
described in this thesis may hold additional potential in the field MICP and this thesis
can serve as a useful reference resource for researchers in microbial biotechnology and
construction microbial biotechnology sub-disciplines.
138
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