CHARACTERIZATION OF UREOLYTIC ISOLATED FROM 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

ABSTRACT

The aim of this study was to isolate, identify and characterise bacteria that are capable of producing 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 substrate through urease for microbially induced precipitation (MICP) applications. Several studies have reported that the majority of ureolytic bacterial 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 ( 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 . 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 group, Pseudogracilibacillus auburnensis group, aureus group, 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 columns treated, all except the columns containing negative control (only cementation solution) had 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 . 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 .

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 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 25 1.5.4. Biodeposition on cementitious materials 27 1.5.5. Biogrout 28 1.5.6. Other essential applications of MICP 30

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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

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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

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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 64 nucleotide BLAST database 2.7 The nomenclatural taxonomy obtained using Ribosomal Database Project- 66 II database 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 76 between individual isolated urease-producing bacteria and control strain 2.11 t-test results comparing the calcite precipitate differences between 79 individual isolated urease-producing bacteria and control strain 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 116 biocemented sands 3.7 Unconfined compressive strength (UCS) of the treated sands 117

3.8 t-test results comparing the unconfined compressive strength (UCS) 118 differences between the biocemented sands 3.9 Summary of calcite content and compressive strength of selected 121 isolates and consortia

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LIST OF FIGURES

Table Page

1.1 Pathway of biominerals secretion and precipitation in a bacterial 11 1.2 Genetic organisation of urease operon in Helicobacter pylori and 13 Sporosarcina pasteurii 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 via urea 26 hydrolysis 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 samples collected from El Toro and El Zancudo limestone 35 mines located in Cordillera Central, northeast of Colombia 1.11 Map of Borneo Island showing the geographical divisions and 38 topographical features of Brunei Darussalam, Indonesia (Kalimantan) and East Malaysia (Sarawak and Sabah) 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 sequence data 68 sequence from different isolates of the current study along with sequences available in the GenBank database 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- 75 producing bacteria and the control strain 2.9 Calcite precipitation media 77 2.10 Comparison of calcite precipitated by selected UPB isolates and the control 78 strain

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2.11 Growth profile of selected ureolytic bacterial isolates and control strain 82 grown in nutrient broth containing 6% urea for 12 hr 2.12 pH profile of selected ureolytic bacterial isolates and control strain grown 83 in nutrient broth containing 6% urea for 12 hr 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, 111 positive and negative controls 3.6 Sand columns at the end of treatment using ureolytic bacteria and 112 cementation solution 3.7 Treated sand removed from their respective columns 113 3.8 Treated sand sample held after a curing period and columns were 114 successfully removed 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

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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

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RH Relative Humidity

SEM Standard Error of Mean

SE Standard Deviation

ANOVA Analysis of Variance

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Chapter 1

INTRODUCTION AND LITERATURE REVIEW

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).

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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 (Hammes, 2003). Calcium carbonate is the most reactive 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 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 of a given community (Rajendhran and Gunasekaran, 2011), with the capability of producing urease enzyme, and induce microbial calcite sufficient for MICP applications.

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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), , 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).

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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).

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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 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).

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1.3 Microbially Induced Calcite Precipitation (MICP) Natural lithification of occurs due to physical, chemical and biological processes (Gadd, 2010) which result in deposition of minerals in the , 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 2- 2+ produce metabolic products (CO3 ) that react with ions (Ca ) 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).

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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):

2+ 2- (1.1) Ca + CO3 ↔ CaCO3 2+ 2- (1.2) Ω = a (Ca ) a (CO3 ) / Kso o -9 (1.3) with Kso calcite, 25 C = 4.8 x 10

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) 2− + (1.7) HCO3− ↔ CO3 + H (pK2 = 10.329)

(1.8) With H2CO3 = CO2(aqueous) + H2CO3

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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 and , (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 - 2− (1.10) 2HCO3 ↔ CO2 + CO3 + H2O 2− - − (1.11) CO3 + H2O ↔ HCO3 +OH 2+ - − (1.12) Ca + HCO3 + OH → CaCO3 + 2H2O

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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).

2+ 2− (1.13) CaSO4·2H2O → Ca + SO4 + 2H2O 2− − - (1.14) 2(CH2O) + SO4 → 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 - 2+ (1.17) 2HCO3 + Ca → 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).

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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).

microbial urease (1.18) CO(NH2)2 + H2O NH2COOH + NH3

(1.19) NH2COOH + H2O → NH3 + H2CO3

+ 2- (1.20) H2CO3 → 2H +2CO3

+ − (1.21) NH3 + H2O → NH4 + OH

2+ 2- −9 (1.22) Ca + 2CO3 →CaCO3 (KSP = 3.8 × 10 )

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 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).

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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).

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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 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).

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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 Bure 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 Aure 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). 13

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

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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 -1 hydrolysed.min ) 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 2+ 2− Ca and CO3 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+ 3− 2− (1.26) Cl − + HCO + NH3 → NH 4Cl + CO3 2+ 2− (1.27) Cell − Ca + CO3 → Cell − CaCO3

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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, Sánchez-Navas et al., 2009), It is also suggested that the proteins of 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 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)

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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 . 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 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 lentus, Bacillus cereus, Psuedomonas antarcticus, Psuedomonas apiaries, Bacillus carboniphilus, , Psuedomonas borealis, Bacillus sporothermodrans, Bacillus lequilensis, Psuedomonas cellulositropicus, , 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 subtilis and Lysinibacillus fusiformis) from calcareous soil samples collected at Anantapur District, , . Hammes et al. (2003b) reported they isolated twelve ureolytic bacteria (closesly related to Sporosarcina pasteurii, Bacillus psychrophilus, 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.

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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 ) 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).

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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 2– 3– of CO3 from HCO 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).

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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).

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1.4.4. Dissolved inorganic carbon Inorganic carbon present in the environment plays a major role in MICP process (Soon, − 2− 2013). Dissolved inorganic carbon (H2CO3 + HCO3 + CO3 ), 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).

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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 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).

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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).

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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).

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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 . 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).

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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, (type strain DSM 8715) and Bacillus cohnii (type strain DSM 6307) (Jonkers, 2007, Jonkers and Schlangen, 2007, Jonkers et al., 2010).

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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).

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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). 28

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).

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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.

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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, 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).

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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 caves, formed from limestone rocks and cave created as a result of lava cavities (Tomczyk-Żak and Zielenkiewicz, 2015). Caves constitute oligotrophic , 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 . 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).

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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 , , 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).

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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 compounds, the water flowing directly from the 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

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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. 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).

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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 , , and (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).

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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 , 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.

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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).

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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.

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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.

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Chapter 2

ISOLATION, IDENTIFICATION AND CHARACTERISATION OF UREASE-PRODUCING BACTERIA FROM LIMESTONE CAVES OF SARAWAK

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.

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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.

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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.

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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 -1 supplemented with C2H3NaO2 (8.2 g.L , HiMedia Laboratories Pvt. Ltd), (NH4)2SO4 -1 -1 (10.0 g.L , HiMedia Laboratories Pvt. Ltd) and CH4N20 (20.0 g.L , 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).

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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).

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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 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.

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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.

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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).

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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 -1 hydrolysed.min ) 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, -1 -1 -1 Oxoid); urea (20.0 g.L , Bendosen); NaHCO3 (2.12 g.L , Sigma); NH4Cl (10.0 g.L , -1 -1 Sigma); CaCl2 · 2H2O (28.50 g.L , Sigma) and agar (20.0 g.L , 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.

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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.

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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.

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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

Sample (%) Sample ID Colour Texture oC collected RH WC1 Drapery Grey Coarse 29.2 76

WC2 White 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

WC= Wind cave; FC= Fairy cave; oC= temperature; (%)RH = relative humidity

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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.

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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.

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A B C

D E F 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.

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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

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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

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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.

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Table 2.3: Morphological characteristics of isolated bacterial colonies

Cave Isolate Form Size Optical Pigmentation Origin ID (shape) (mm) Property 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

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Table 2.4: Microscopic characteristics of bacterial isolates

Isolate Gram Cell Endospore ID stain shape 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

+ve= positive; -ve= negative.

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Table 2.5: Biochemical characteristics of bacterial isolates

Isolate Oxidase Catalase Motility ID 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 +ve= positive; -ve= negative.

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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, 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 Sporosarcina while the rest UPB were classified into the family and genus Bacillus and Staphylococcus. A phylogenetic tree shown in Figure 2.7 was constructed using a neighbour-joining method (Liang et al., 2008).

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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.

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Table 2.6: Molecular identification based on 16S rRNA sequencing data using NCBI nucleotide BLAST database

GenBank Query Isolate No Accession Closest match Base pair Cover Similarity ID number 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%

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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%

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Table 2.7: The nomenclatural taxonomy obtained using Ribosomal Database Project-II database

Isolate No Phylum Class Order Family Genus ID 1 NB33 Bacteria Firmicutes 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

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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

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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).

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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.

69

0.9

0.8

y = 0.1415x + 0.0862 0.7

0.6 ) 1 -

0.5

0.4

Conductivity (mS.cm 0.3

0.2

0.1

0.0 0 1 2 3 4 5 6 Time (min)

Figure 2.7: Relative conductivity of isolate LPB21 measured for a duration of 5 min. Error bars represent standard error of the mean.

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Table 2.8: Measurement of conductivity variation rate and SEM

Isolate Conductivity variation rate No SEM ID (mS.cm-1.min-1)

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

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Table 2.9: Conversion of changes in conductivity to urease activity

(mS.cm-1.min-1) mM urea No Isolate ID (mS.cm-1.min-1) *df 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.

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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).

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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.

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30

25

) 1 - 20

.min.OD 1 - 15

activity urease Specific 10

hydrolysed urea (mM

5

0

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.

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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.

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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.

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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.

20

18

16

14

) 1 - 12

(mg.mL 10

precipitates of weightcalcite 8

6

4

2

0 control NB30 LPB21 NB33 NB28

Figure 2.10: Comparison of calcite precipitated by selected UPB isolates and the control strain. Error bars represent standard error of the mean.

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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)

Isolate P- No M SD SE t P <* ID value 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.

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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.

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Table 2.12: Kinetics growth of ureolytic bacteria in batch cultures

Specific growth Maximum growth of Doubling time,td Isolate rate,k bacteria -1 [g] ID [h ] [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

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.

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1.2

1.0

0.8

Control

NB33 0.6 LPB21

600 NB30 OD NB28

0.4

0.2

0.0 0 2 4 6 8 10 12 14

Time (hour)

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.

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9.5

9.4

9.3

9.2

9.1 Control

NB33 pH 9.0 LPB21

NB30 8.9 NB28

8.8

8.7

8.6 0 1 2 3 4 5 6 7 8 9 10 11 12 Time (hour)

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.

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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.

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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 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).

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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.

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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).

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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 ’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: + 2- ammonium (NH4 , positively charged) and carbonates (CO3 , 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).

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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 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).

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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.

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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.

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Chapter 3

EFFECTS OF CULTURAL CONDITIONS ON UREASE ACTIVITY AND EVALUATION OF BIOCEMENTATION POTENTIALS IN SMALL SCALE TEST

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.

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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.

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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 Sporosarcina pasteurii strain DSM33 Control Comprised of four isolates; Sporosarcina pasteurii LPB21 (SUTS), Bacterial Sporosarcina pasteurii NB30 (SUTS), Sporosarcina pasteurii NB28 Consortia (SUTS) and Sporosarcina pasteurii NB33 (SUTS)

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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 1 M (CO(NH2)2) Calcium chloride 1 M (CaCl2) Sodium acetate 0.17 M (C2H3NaO2) Ammonium chloride 0.0125 M (NH4Cl) 13 g/L Nutrient broth

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.

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Table 3.3: Sand characteristics

Uniformity Coefficient of Sand D10 D30 D60 coefficient gradation Source (mm) (mm) (mm) (Cu) (Cc)

Kuching, 1.6 0.907 0.220 0.265 0.352 Sarawak

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 -1 contained cementation reagents, nutrient broth (13 g.L ), 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.

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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.

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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.

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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).

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control LPB21 NB33 NB28 NB30

50

45 *

40 1 -

.OD

1 35 -

30 * * * 25

20

Specific activity urease 15 (mM ureahydrolysed.min(mM

10

5

0 20 25 30 35 40 45

Temperature (oC)

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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Control LPB21 NB33 NB28 NB30 40

* * 35 * 30 ) 1 - .OD 1

- * 25 * 20

15 Specific activity urease

ureahydrolysed.min(mM 10

5

0 6 6.5 7 7.5 8 8.5 pH

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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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).

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Control LPB21 NB33 NB28 NB30 35

* 30 * *

) * 1 - 25 .OD 1 - *

20

15

Specific activity urease 10 ureahydrolysed.min(mM

5

0 24 48 72 96 Incubation Period (hr)

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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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).

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Control LPB21 NB33 NB28 NB30

45 * 40 * 35 * ) 1

-

.OD 30 1 - * * 25

20

15 Specific activity urease

ureahydrolysed.min(mM 10

5

0 2 4 6 8 10 urea concentration (%)

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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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.

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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

Percentage Characteristics (%)

Fine sand 6.72

Medium sand 87.95

Coarse sand 3.96

Fine gravel 1.37

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Table 3.5: Bacteria concentration and urease activity prior to biocement test

-1 mM urea Isolate OD600 CFU.mL 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

-1 OD600 = optical density; CFU.mL = colony forming unit per millilitre; mM urea hydrolysed.min-1.OD-1 = urease activity.

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A B

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.

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Consortia NB28 NB30 NB33 LPB21 Negative Positive control control

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.

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Positive Consortia LPB21 NB33 NB28 NB30 control

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.

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A B C

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.

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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.

800

700

600

500

(psi)

Strength 400

300

200

100

0 - control + control LPB21 NB33 NB30 NB28 consortia

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.

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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 M SD SE P-value t P <* ID

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.

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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

UCS test Bacteria ID Condition of Force Pressure cemented sand (kN) (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.

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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 P- M SD SE t P <* ID value

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.

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3.3.8. Calcite confirmation

A B

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.

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3.3.9. Calcite content Determination

top middle bottom 12

10

8

6

4

(w/w) % weight,Calcite 2

0

- control + control LPB21 NB30 NB28 NB33 Consortia

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.

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Table 3.9: Summary of calcite content and compressive strength of selected isolates and consortia

Weight of calcite, % (w/w) Strength Isolate conversion ID (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

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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.

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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.

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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.

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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.

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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.

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Chapter 4

GENERAL CONCLUSIONS AND RECOMMENDATIONS

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).

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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.

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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.

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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 but are surrounded by layers of thick 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 -1 urease activity (mM urea hydrolysed.min ) 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.

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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.

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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.

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