Microbial cell attachment, colonisation and degradation of coal

Nur Hazlin Hazrin Chong

A thesis in fulfilment of the requirements for the degree of Doctor of Philosophy

School of Biotechnology and Biomolecular Sciences Faculty of Science The University of New South Wales Sydney, Australia September 2013

Abstract

Discoveries of aerobic coal degrading microorganisms have led to their utilisation in various biotechnological coal processes. One promising application of these microbes is the acceleration of coal to methane, which provides an avenue for more sustainable coal usage. However, despite the various findings of coal degraders and related mechanisms, a key aspect in coal degradation, which is cell attachment and colonisation, has been largely neglected.

This study is among the first to describe in detail microbial cell attachment and colonisation on coal. Using coal-degrading bacteria and fungi, the initial cell attachment and biofilm formation on coal were investigated across different coal types and conditions. Physico-chemical analyses based on contact angle measurements revealed that hydrophobicity, surface free energy and adhesion thermodynamics, as well as secondary biological and environmental factors, played a crucial role in governing the first form of cell interaction with coal. Direct observation and electron microscopy highlighted different colonisation mechanisms on coal based on cell morphology, surface topography and environmental conditions. Correlations were found between colonisation and degradation of coal, which stressed the importance of cell attachment in coal degradation, although exceptions were present.

Another interest of this study was to isolate native coal-degrading fungi for potential field applications. Through multiple coal degradation screenings, Fusarium oxysporum G9o was discovered as a promising bituminous coal-degrading fungus. The isolate showed softening of raw bituminous coal, and infrared analyses revealed oxidation and cleaving mechanisms of coal components. Further, the colonisation of coal by soil communities was monitored through microbial community analyses using Terminal- Restriction Fragment Length Polymorphism (T-RFLP) and pyrosequencing analyses. Unique communities from soil were identified as dominant colonisers on coal, which have not been previously revealed through conventional cultural techniques.

Overall, the findings in this study provide valuable insights into the mechanisms of cell attachment and colonisation on coal. This serves as a foundation for a new research area in coal microbiology, which will increase our currently limited understanding on coal-cell interactions.

ii

Dedication

This thesis is dedicated to my four pillars of life,

God, my parents, and husband

God, for creating my existence in life, for providing me sustenance, for showing me strength and patience, and for showering your Love and Mercy

Mum and Dad, for being the reason I am here today, for your unconditional and constant love and support, and for my valued and healthy upbringing

My husband, for being my best friend, for sharing my every happiness and sadness, and for your endless love and support

iii Acknowledgements
I am eternally grateful to have met and worked with an amazing group of people, of whom without their help, this thesis could not have been written.

Thank you to my supervisor Dr Mike Manefield for his excellent guidance, support and encouragement throughout my whole candidature. Mike, you never cease to inspire me to become a better scientist, thinker and writer. Your words and gestures have made a positive impact to me as a young researcher and I am forever thankful to have embarked on this PhD journey with you as my guide. Thank you for believing in me. I hope I have made you proud and will continue to do so in my future career.

Thank you to Dr Maria Luisa Gutierrez-Zamora for her dedicated support, guidance and friendship. I truly appreciate all your help and advice on so many aspects of this PhD project, right from when I first started until I handed in my thesis. I owe you a lot for teaching me the bulk of the molecular work done in this study. Your positive vibe in the lab and our meetings were always infectious to me and everyone else. I am so thankful to know you as an excellent researcher and friend!

Special thanks to Dr Chris Marjo and Dr Anne Rich from the Solid State Elemental Analysis Unit at UNSW for their kind help and support in teaching me everything about FTIR. Thanks to Anne for showing me how to prepare the coal samples, and operate and troubleshoot the FTIR machine. Thanks to Chris for his thorough explanations on interpreting FTIR spectra and the complexity (and discovered beauty) behind it.

Thank you to Jenny Norman from the Electron Microscopy unit at UNSW for the training and technical advice to produce high quality SEM micrographs, which were necessary for fundamental observations on the bacterial and fungal attachment on coal.

I am thankful to Dr Theerthankar Das for his expert help and excellent support in the thermodynamics of adhesion works in this study, which played an important role in the discovery on the physico-chemical relationship between microbes and coal.

Thank you to Professor Colin Ward for his geological advice on coal. Especially thank you for providing me with the lignite coal, which gave very useful results in this study. Also thanks to Joanne Wilde from the BEES Rock Preparatory lab at UNSW for her coal polishing services.

iv Thanks very much to Shaun Nielsen for teaching me how T-RFLP and pyrosequencing work, and Dr Ezequiel Marzinelli for teaching me statistical analyses using the PRIMER-E software. Thanks to John Webster for his advice on sub-sampling my pyrosequencing data.

Thank you to Dr Sohail Siddiqui and Dr Haluk Ertan from the Cavicchioli lab for kindly showing me how to perform the enzymes assay and calculations.

To the Manefield group, I am grateful to have you as my research mates. Mona, Iman, Onder, Das, Sharma, Leena, Matt, Valentina, John and Sabrina- you guys are a terrific bunch. Thank you for all your help in the lab and office and for the nice and/or funny conversations we’ve had. Special thanks to Dr Adrian Low for all your help, and Mona El Hassan and Iman Taleb for your great friendship.

I want to thank Professor Staffan Kjelleberg and Professor Peter Steinberg from the Centre for Marine Bioinnovation for allowing me to be a part of their fantastic research group. Special thanks to Kirsty, Leena, Adam, Chau and Amy for their helpful administrative support. Thanks Gee, Raymond, Vipra and the whole 304/616 lab for your help and support whenever I needed them. Not forgetting Arthur Bauer, Josh Glass and Sophie Holland as wonderful and helpful interns!

Thank you to Peter Francis and Glen Cunningham from Biogas Energy Pty. Ltd. for funding my research. A special thanks to the coal to methane (C2M) group led by Mike and Dr Torsten Thomas for giving me helpful feedback in the meetings.

Thank you to the Malaysian Ministry of Higher Education and National University of Malaysia for financially supporting my study abroad and ensuring my well being as an international student.

Special thanks to James Hayton for giving me helpful advice and support throughout the final months of my PhD. It’s been great working with you!

Thank you to all my lovely akhawat: Kak Mel, Kak Hani, Afida, Ainul, Dayah R, Ain, Elyna, Nadiah and the rest (it’s a long list!) for being there for me and supporting through my PhD journey. Uhibukkunna fillah abadan abada.

Last, but definitely not least, a HUGE thank you and appreciation to Arif for being the most loving, understanding, supportive and patient husband I can ever have. Your constant push and motivation for me to do my best are deeply appreciated. You are truly a kind person and I am grateful to have the best partner in life. Thank you Mama and Baba, for raising Arif to become a wonderful person he is today.

v Table of contents

Abstract ...... ii Dedication ...... iii Acknowledgements ...... iv Table of contents ...... vi List of publications ...... x List of figures ...... xi List of tables ...... xv Abbreviations ...... xvi

1 A literature review on the microbial cell attachment and degradation of coal ...... 1 1.1 Overview ...... 1 1.2 Coal ...... 4 1.2.1 Origin of coal ...... 4 1.2.2 Coal ranks ...... 4 1.2.3 Physical and chemical properties of coal ...... 5 1.2.4 Coal biodegradation ...... 7 1.2.5 Aerobic coal biodegradation ...... 10 1.3 Biofilms – Cell adhesion and microbial surface colonisation ...... 19 1.3.1 Biofilm formation ...... 20 1.3.2 Fungal biofilms ...... 23 1.3.3 Environmental biofilms ...... 25 1.3.4 Factors that influence cell attachment and biofilm formation ...... 26 1.3.5 Methods in studying cell attachment and biofilm formation ...... 29 1.4 Microbial adhesion and colonisation on coal ...... 33 1.5 Aims of this study ...... 36

2 Pseudomonas fluorescens attachment and oxidation of bituminous coal ...... 38 2.1 Introduction ...... 38 2.2 Material and Methods ...... 39

vi 2.2.1 Medium preparation ...... 39 2.2.2 Culture preparation ...... 40 2.2.3 Coal preparation ...... 40 2.2.4 Control surfaces preparation ...... 43 2.2.5 Attenuated Total Reflectance- Fourier Transform Infra Red (ATR-FTIR) Spectroscopy of coal ...... 43 2.2.6 Scanning Electron Microscopy of P. fluorescens attachment on bituminous coal


...... 44 2.2.7 Imaging of P. fluorescens cells in supernatant in the presence of coal and without glucose and CAA ...... 45 2.2.8 Contact angle measurements and surface thermodynamics of adhesion between P. fluorescens and coal ...... 45 2.2.9 Enumeration of P. fluorescens cells on different types of coal ...... 48 2.2.10 Live and dead P. fluorescens attachment on coal ...... 48 2.3 Results ...... 49 2.3.1 ATR-FTIR of coal oxidation by P. fluorescens ...... 49 2.3.2 Scanning Electron Microscopy (SEM) of P. fluorescens attachment on bituminous coal in different media ...... 52 2.3.3 Contact angle (CA) measurements ...... 59 2.3.4 Thermodynamics of adhesion between P. fluorescens and coal ...... 64 2.3.5 Live and dead P. fluorescens cell attachment to bituminous coal ...... 68 2.4 Discussion ...... 69

3 Isolation of coal-degrading fungi from an abandoned shale mine in The Wolgan Valley and Newnes forest, New South Wales ...... 84 3.1 Introduction ...... 84 3.2 Material and Methods ...... 87 3.2.1 Coal preparation ...... 87 3.2.2 Elemental analysis of coal ...... 87 3.2.3 Sample collection ...... 88 3.2.4 Inocula preparation ...... 89 3.2.5 Initial screening for coal-degrading microbes on coal agar ...... 89 3.2.6 Further screening of coal-degrading isolates on coal silica ...... 89 3.2.7 Molecular identification of coal-degrading isolates ...... 90 3.2.8 Coal colonisation and degradation assay ...... 93 3.2.9 Oxidative enzyme assay ...... 94

vii 3.3 Results ...... 95 3.3.1 Coal elemental analysis ...... 95 3.3.2 Screening of isolates through coal agar and coal silica ...... 96 3.3.3 Identification of fungal isolates ...... 98 3.3.4 Coal colonisation and degradation assay ...... 101 3.3.5 Oxidative enzyme assay ...... 111 3.4 Discussion ...... 114

4 Characterisation of the fungus Fusarium oxysporum G9o on the attachment and degradation of coal ...... 124 4.1 Introduction ...... 124 4.2 Material and Methods ...... 125 4.2.1 Molecular investigation on species identification of Fusarium sp. G9o .. 125 4.2.2 Direct observation and Scanning Electron Microscopy (SEM) of bituminous coal colonised by F. oxysporum G9o ...... 127 4.2.3 Contact angle measurements and surface thermodynamics of adhesion of F. oxysporum G9o on coal ...... 127 4.2.4 Fourier Transform Infrared (FTIR) analysis of coal incubated with F. oxysporum G9o ...... 128 4.2.5 Analysis of F. oxysporum G9o coal degradation in liquid culture ...... 128 4.2.5 Extracellular oxidative enzyme activity analysis of F. oxysporum G9o .. 129 4.2.6 Statistical analyses ...... 131 4.3 Results ...... 131 4.3.1 Species identification of Fusarium sp. through 18S, ITS and LSU sequencing ...... 131 4.3.2 Direct observation and Scanning Electron Microscopy (SEM) of bituminous coal colonised by F. oxysporum G9o ...... 132 4.3.3 Contact angle measurements and surface thermodynamics of adhesion between F. oxysporum G9o and coal ...... 135 4.3.4 Fourier Transform Infrared (FTIR) analysis of different coal types colonised and/or degraded by F. oxysporum G9o ...... 138 4.3.5 Coal degradation by F. oxysporum G9o in submerged conditions ...... 142 4.3.6 Extracellular oxidative enzyme activity of F. oxysporum G9o in the presence of coal ...... 146 4.4 Discussion ...... 148

viii 5 A microbial community analysis of bituminous coal buried in soil from the Lithgow State Coal Mine, New South Wales ...... 160 5.1 Introduction ...... 160 5.2 Material and methods ...... 161 5.2.1 Experimental set up ...... 161 5.2.2 Coal sampling and DNA extraction ...... 162 5.2.3 Bacterial 16S rRNA gene amplification of the scraped biomass on coal and LSCM soil ...... 163 5.2.4 Terminal restriction fragment length polymorphism (T-RFLP) of 16S rRNA genes extracted from biomass in soil and on coal ...... 164 5.2.5 454 pyrosequencing and quality filtering ...... 165 5.2.6 Statistical analyses for T-RFLP and pyrosequencing data ...... 166 5.2.7 Scanning Electron Microscopy of coal pieces exposed to soil ...... 166 5.3 Results ...... 166 5.3.1 DNA extraction and amplification of biomass on coal ...... 166 5.3.2 Terminal restriction fragment length polymorphism (T-RFLP) of DNA extracted from soil and biomass on coal ...... 167 5.3.3 Pyrosequencing results ...... 172 5.3.4 Scanning Electron Microscopy of biomass on coal ...... 180 5.4 Discussion ...... 183

6 General discussion ...... 191 6.1 Coal as substratum for cell attachment and colonisation ...... 192 6.2 Linking cell attachment and colonisation with degradation of coal ...... 197 6.3 Fusarium oxysporum G9o: A suitable candidate in the application of coal bioconversion to methane? ...... 198 6.4 Future directions ...... 200 6.5 Concluding remarks ...... 201
References ...... 202
Appendices ...... 231

ix List of publications

Hazrin-Chong, N.H., Marjo, C.E., Das, T., Rich, A.M. and Manefield, M. (2013). Surface analysis reveals biogenic oxidation of bituminous coal by Pseudomonas fluorescens (in preparation).

Hazrin-Chong, N.H. and Manefield, M. (2012). An alternative SEM drying method using hexamethyldisilazane (HMDS) for microbial cell attachment studies on sub-bituminous coal. Journal of Microbiological Methods, 90: 96-99

Conference proceeding

Beckmann, S., Hazrin-Chong, N.H., Webster, J., Thomas, T., Manefield, M. (2012). Acetoclastic methanogenesis in a microbial community associated with a sub-bituminous coal seam in New South Wales, Australia. Poster presentation. Proceedings of the 14th International Symposium on Microbial Ecology (ISME 14), 19-24 August 2012. Copenhagen, Denmark.

x List of figures

Figure 1.1 The coalification process that transforms peat into different bands of coal ______4 Figure 1.2 Physical and chemical representations of coal ______6 Figure 1.3 Proposed stepwise degradation of coal ______8 Figure 1.4 Example of coal (lignite) solubilisation by Alternaria sp. pre-grown on an agar plate ______17 Figure 1.5 Stages in biofilm formation ______20 Figure 1.6 Biofilm formation stages for filamentous fungi ______24 Figure 2.1 (A) Representative FTIR stackplot of: (i) P. fluorescens on a glass slide; (ii) P. fluorescens on coal; and (iii) coal sample with no biological material. (B) Expanded region of spectrum ______50 Figure 2.2 Representative micrographs of P. fluorescens attachment on coal in M9 medium with glucose and CAA after 24 h of incubation ______53 Figure 2.3 Representative micrographs of P. fluorescens attachment on coal in glucose-only M9 medium after 24 h of incubation ______54 Figure 2.4 Representative micrographs of P. fluorescens attachment on coal with coal as the sole carbon source after 24 h of incubation ______55 Figure 2.5 Microcolony of P. fluorescens in M9 medium without external carbon source present. ______56 Figure 2.6 A microcolony of P. fluorescens consisting of short and extended cells (square) colonising a coal microparticle in the absence of an additional carbon source. ______56

Figure 2.7 Optical density (OD600) of supernatants of P. fluorescens used in the coal adhesion experiment in the presence and absence of glucose ______57 Figure 2.8 Representative image of free-living P. fluorescens in M9 medium without glucose and CAA ______58 Figure 2.9 Representative micrographs of P. fluorescens on broken glass (A) and acid-washed pebble (B) in M9 medium without external carbon source after 24 h of incubation ______59

xi Figure 2.10 Average water contact angles of three types of P. fluorescens: Glucose-fed, starved and killed ______60 Figure 2.11 Water contact angles of surfaces of P. fluorescens (glucose-fed), various coal types and glass slides ______61 Figure 2.12 The number of starved P. fluorescens cells (cells/mm2) attached to various types of coal surfaces. ______68 Figure 2.13 Representative SEM micrographs of killed and live P. fluorescens on bituminous coal. ______69 Figure 2.14 Proposed chemical groups based of FTIR spectra in this study _ 71 Figure 3.1 Examples of isolates from a non-diluted sample grown on coal agar in different media types ______97 Figure 3.3 Representative images of preferential and non-preferential colonisation on coal by isolates ______102 Figure 3.4 Second test of coal colonisation and solubilisation assay _____ 103 Figure 3.5 Representative images of non-filamentous isolates showing no colonisation on coal after several weeks of incubation ______104 Figure 3.6 A summary of the proportion of isolates colonising coal ______104 Figure 3.7 Fusarium sp. G9o colonisation and degradation of the untreated coal in comparison to the treated coal ______106 Figure 3.8 P. chrysosporium degradation of untreated bituminous coal after 6 months of incubation at 22 oC ______107 Figure 3.9 Lignite coal solubilisation by isolate Penicillium sp. Y7e before and after 3 months of incubation. ______108 Figure 3.10 Lignite coal solubilisation by isolate Penicillium sp. G5o before and after 3 months of incubation ______108 Figure 3.11 Lignite coal solubilisation of P. chrysosporium after approximately 3 months of incubation ______109 Figure 3.12 G9o colonisation and solubilisation test of four different types of coal______110

Figure 3.13 Bottom view of agar plates containing MnCl2 grown with isolates G9o, G5o and Y7e ______112 Figure 3.14 Bottom view of agar plates containing ABTS grown with isolates G9o, G5o and Y7e ______113 Figure 3.15 Reactive Violet 5 plate grown with G5o on 0, 2 and 10 days __ 113
xii Figure 3.16 Bottom view of Remazol Brilliant Blue R plate grown with Y7e at the start, middle and end of incubation ______114 Figure 4.1 Phylogenetic tree generated by BioNJ method based on ITS rDNA sequences of Fusarium oxysporum G9o and sequences from GenBank ___ 132 Figure 4.2 Untreated bituminous coal pieces incubated with G9o on Day 25 and 52 ______132 Figure 4.3 Representative SEM micrographs of F. oxysporum G9o cell morphology ______133 Figure 4.4 Representative micrographs of coal surfaces exposed to G9o colonisation ______134 Figure 4.5 Water contact angle of G9o surface on Days 2, 7 and 14 in comparison to those of untreated bituminous coal and P. fluorescens _____ 135 Figure 4.6 Representative FTIR spectra of untreated bituminous and nitric acid- treated bituminous coal modified by G9o against the unmodified control ___ 140 Figure 4.7 Representative FTIR spectra of Loy Yang lignite and Kalimantan lignite modified by G9o against the unmodified coal ______141 Figure 4.8 A) Representative F. oxysporum G9o liquid cultures in glutamate medium with 1% w/v glucose and no coal or 0.25% w/v lignite coal or 0.25 % w/v untreated bituminous coal ______142 Figure 4.9 Supernatants of G9o with coal (lignite or untreated bituminous) and controls (G9o only and coal only) ______143 Figure 4.10 Absorbances at 300, 350, 400, 450 and 650 nm of G9o supernatants with and without coal for 37 days ______145 Figure 4.11 Manganese peroxidase, lignin peroxidase and laccase extracellular enzyme activity (mU/ml) by G9o, with and without coal (lignite or bituminous coal) during 14 days of incubation ______147 Figure 4.12 A possible pathway for oxidation of a hypothetical coal residue 155 Figure 5.1 Representative agarose gel illustrating amplified DNA from the scraped biomass of the untreated coal and treated coal against their reference soil samples over the sampling time (week) ______167 Figure 5.2 Non-metric multidimensional scaling (nMDS) plots of the T-RFs of scraped biomass of untreated coal and treated coal samples against their respective soil reference samples ______169

xiii Figure 5.3 Average similarity (%) between the community composition among replicates for each sample (Untreated and treated coal and their respective soil samples) over 27 weeks of incubation. ______170 Figure 5.4 Representative 16S rDNA T-RFLP electropherograms of soil and coal at week 27 ______171 Figure 5.5 Overview of 101 individual OTUs (y-axis) across individual coal and soil samples (x-axis) at weeks 17 and 27 ______175 Figure 5.6 A non-metric multidimensional scaling (nMDS) plot of OTUs of coal and soil samples ______176 Figure 5.7 Abundance of OTUs (%) in the treated coal replicates (Treated 1, 2 and 3) at week 27 ______177 Figure 5.8 Proportion of the most abundant OTUs in ‘Untreated 1 Week 17’ coal sample, where Methanococcoides sp. and Porphorybacter sp. (in bold) dominated the sample population ______178 Figure 5.9 Fungal OTUs of coal and soil samples at week 17 and 27 _____ 179 Figure 5.10 Soil aggregate attachment on the untreated coal. ______180 Figure 5.11 A) Soil aggregate attachment on the treated coal. B) A blown up image of a soil aggregate in (A), which consisted of complex and heterogeneous material. ______181 Figure 5.12 Microbial-looking entities observed on coal and across the soil aggregates ______182

xiv List of tables

Table 1.1 Examples of brown (low-rank) and black (high-rank) coal-degrading microorganisms and associated mechanisms of degradation. ______15 Table 2.1 Test liquids and their surface energy components. ______46 Table 2.2 Peak fitting statistics of a region rich in P. fluorescens on coal ___ 51 Table 2.3 Peak fitting results for influence of peroxidase on coal ______52 Table 2.4 Cell length and width (μm) of P. fluorescens on coal in different M9 medium. ______58 Table 2.5 A summary of all contact angle work, i.e. water (θ W), formamide (θ F) and diiodomethane (θ Di) on various P. fluorescens and coal surfaces and glass as control ______63

Table 2.6 Total Gibbs free energy of adhesion (∆GAdh) between differently treated P. fluorescens cell and coal surfaces with glass as reference ______66 Table 3.1 A list of samples obtained from Wolgan Valley, NSW, as inocula for the isolation of coal-degrading microorganisms. The specific description and location of the samples are included. ______88 Table 3.2 Elemental analysis data of coal samples used: 1) Untreated bituminous, 2) Nitric acid-treated bituminous, 3) Loy Yang lignite, and 4) Kalimantan lignite. All data presented were calculated to a dry, ash-free basis (% daf) ______96 Table 3.3 Molecular identification of several fungal isolates obtained in this study ______100 Table 3.4 Oxidative enzyme assay results with isolates G9o, G5o and Y7e 112 Table 4.1 Contact angle and calculated surface energy components of G9o on days 2, 7 and 14 ______137

Table 4.2 Total free energy of adhesion (∆GAdh) for F. oxysporum G9o against untreated bituminous, nitric acid-treated bituminous and lignite coals across days 2, 7 and 14 of incubation ______138

xv Abbreviations

γ Surface free energy

γ + Electron accepting sub-component of acid-base surface energy component

γ - Electron donating sub-component of acid-base surface energy component

∆GAdh Surface free energy of adhesion

θ Contact angle

μL microlitre

μm micrometre

μM micromolar AB Acid-base ATR-FTIR Attenuated total reflectance – Fourier transform infrared

CH4 Methane

CO2 Carbon dioxide DGGE Denaturing gradient gel electrophoresis dH20 Deionised water DNA Deoxyribonucleic acid EDTA Ethylenediaminetetraacetic acid EPS Exopolymeric substance FAM 6-carboxy-fluorescein FTIR Fourier transform infrared f. sp. Forma specialis g Gravity, in relation to relative centrifugal force KBr Potassium bromide LiP Lignin peroxidase
xvi LW Lifshitz-Van der Waals m metre M9 Minimal 9 salts min s-1 Minute per second min Minute ml millilitre Mn Manganese MnP Manganese peroxidase nMDS Non-metric multidimensional scaling PBS Phosphate buffered saline PCR Polymerase chain reaction rDNA Ribosomal deoxyribonucleic acid rRNA Ribosomal ribonucleic acid SEM Scanning electron microscopy s Second sp. species TAE Tris-Acetate-EDTA TES Trace element solution T-RFLP Terminal-restriction fragment length polymorphism T-RFs Terminal-restriction fragments W Watt

xvii

1 A literature review on the microbial cell attachment and degradation of coal

1.1 Overview

Throughout history, coal has been an important source of energy worldwide. Its high abundance and energy density has made coal one of the most utilised and economically preferred fossil fuels for many decades. This continues to be the case despite a growing interest in renewable energy technologies. In 2012, approximately 6.6 billion tonnes of black coal and 1 billion tonnes of brown coal were used globally, mainly for electricity generation and steel production (International Energy Agency 2012; World Coal Association 2012b; World Coal Association 2012c). This accounts for almost half the total global energy demand, where the other half is a combination of other fuels, including renewables (World Coal Association 2012a). A continuous surge of the global energy demand is anticipated due to an increasing access to modern technologies, not only in developed countries, but more drastically in developing nations today (World Coal Association 2012b; World Coal Association 2012a). Based on the current production, consumption and availability of coal, it is likely that the hydrocarbon will remain relevant in meeting the energy demand for another century (BP P.L.C. 2010).

As the demand for coal involves energy-intensive processing i.e. high- temperature and pressure coal combustion, the latter inevitably contributes to an increase in global warming. Therefore, there is a need to utilise coal in a more environmentally sustainable manner. More recently, the discovery of methane production from coal as a viable energy resource claims to be the ‘greener’ alternative of coal utilisation. More interestingly from a microbiological perspective, it has been found that methane generated from coal is largely driven by microorganisms living within many different coal seams across the
1 globe (Ahmed and Smith 2001; Aravena et al. 2003; Montgomery 1999; Strąpoć et al. 2007; Zhou et al. 2005). Whilst methanogenic archaea are responsible for the actual production of methane gas, other coal-degrading microbes act initially to degrade coal into simpler molecules and eventually H2, CO2 and/or acetate that feed methanogenesis (Strąpoć et al. 2008). Although mostly an anaerobic process, the first stages of the transformation of coal to methane can be carried out aerobically. A biotechnological approach could allow the utilisation of aerobic coal-degrading microorganisms to accelerate the biotransformation of coal to methanogenic substrates. Applying aerobic microbes could allow for an enhanced initial rate-limiting step, the fragmentation of the recalcitrant components of coal, to accelerate coal transformation to methane. This has only been shown very recently by a study that utilised an aerobic fungal isolate, Penicillium chrysogenum, for the initial coal degradation as a precursor for methanogenesis (Haider et al. 2013).

Discoveries of aerobic coal-degrading microorganisms and their mechanisms of degradation have spanned a century (for a full review on the history refer to Hofrichter and Fakoussa 2001). The first reported isolation of bacteria grown on coal was made in 1910 (Galle 1910). Several years after, similar observations were made on fungi (Fischer and Fuchs 1927a; Fischer and Fuchs 1927b). However, it was not until the early 1980’s that Fakoussa (1981) and Cohen and Gabriele (1982) first discovered that certain bacteria and filamentous fungi were not only able to utilise coal as a sole carbon source but also to solubilise it. This led to other worldwide discoveries of bacteria and fungi that showed coal- degrading ability through extracellular enzyme production (Hofrichter et al. 1997a; Hofrichter and Fritsche 1997; Hofrichter et al. 1997b; Holker et al. 1995; Holker et al. 1999; Igbinigie et al. 2008; Laborda et al. 1997; Monistrol and Laborda 1994; Silva-Stenico et al. 2007; Wilmann and Fakoussa 1997b; Wondrack et al. 1989). Although these studies provided breakthrough results in the discovery of coal-degrading microorganisms and their associated degradation mechanisms, a fundamental and important investigation, namely the initial cell attachment to and biofilm formation on coal, was often overlooked.

2 Many of the above mentioned studies have observed coal surface colonisation by the isolated microbes occurring prior to coal degradation. However, these observations were often briefly discussed and no thorough investigation on the role of cell attachment and biofilm formation on coal was conducted, leaving a knowledge gap that is critical to fill towards a better understanding of microbe- coal interactions. Studies on cell attachment and/or biofilm formation have been extensively conducted in other fields, e.g. infection and diseases, cell-to-cell communication, biofouling, steel corrosion, microbial fuel cells and many others, which have shown the importance and prevalence of this subject across various disciplines. Therefore, it seems fitting that this aspect is also investigated in coal microbiology, which further broadens biofilm research and adds valuable insights to the existing knowledge on microbial adhesion to hydrocarbons.

This review is divided into three main sections, each with a specific aim:

1) Coal. To appreciate coal as a substratum for attachment and substrate for microbial degradation. General information on coal, including its origin, ranks and physical and chemical properties are included. Existing knowledge on primarily aerobic coal biodegradation is discussed. 2) Biofilms. To summarise established theories and studies on cell attachment and biofilm formation. Different stages of biofilm growth and biofilm types are highlighted. A section on the analysis of microbial communities in environmental biofilms is also discussed. Several factors influencing cell attachment and biofilm formation, which focus on the physico-chemical components, are reviewed. Current techniques in analysing cell attachment and colonisation are also considered. 3) Microbial adhesion to coal. To highlight previous observations on cell attachment to coal. A critical summary of the existing literature on microbial adhesion to coal is given.

3 1.2 Coal

1.2.1 Origin of coal

Coal is a combustible, sedimentary rock, which consists of plant debris that originated over hundreds of millions of years. The plant debris was once contained in a swampy depositional environment to create a soft, spongy material called peat (Ward 2001). Through time, physical and chemical processes that were caused by elevated heat and prolonged burial (i.e. pressure) transformed peat into coal of different ranks. This transformation process is called ‘coalification’ or ‘rank advance’ as illustrated in Figure 1.1.

Figure 1.1 The coalification process that transforms peat into different bands of coal. Image was obtained from Greb (n.d.) from Kentucky Geological Survey, University of Kentucky.

1.2.2 Coal ranks

Depending on the degree of coalification, different coal ranks are formed. The lowest order is lignite, followed by sub-bituminous, bituminous and anthracite. The two former coals are also known as ‘brown coal’ or ‘soft coal’, and the latter
4 three as ‘black coal’ or ‘hard coal’. Various properties in coal are analysed to determine its rank. These include the total amount of carbon, oxygen, mineral matter, ash and volatile material in coal (Ward 1984). Recognisable morphological features in coal are called macerals representing certain components of the original plants (e.g. root, bark, spores) (Stach et al. 1982). Vitrinites, inertinites and liptinites are the main classification of macerals in coal, where varying proportions of each can be observed (through microscopy) for different coal ranks.

1.2.3 Physical and chemical properties of coal

The degree of the coal rank reflects its physical and chemical properties. Although solid, coal is naturally porous due to the parallel sets of planar fractures called cleats. Cleats are naturally found in coal beds as a result of various stresses (e.g. compaction and desiccation) to the coal bed during the coalification process (Masterlerz et al. 1999). They are usually analysed as a measure of permeability (and thus difficulty) of the coal bed in releasing methane (Laubach et al. 1998; Masterlerz et al. 1999). Although permeability appears to be characteristic in coal, it tends to decrease as the coal rank increases, thereby making lignite the most porous among all coal types (Masterlerz et al. 1999).

Coal is mainly comprised of carbon, with a lower proportion of hydrogen, oxygen and nitrogen and even lower of sulfur, phosphorus, chlorine and trace elements. Depending on the coal rank, the relative proportion of each element differs in each coal. This is also translated into different overall molecular structures found at each coal rank. Due to the molecular complexity and heterogeneity found in coal, it is difficult to elucidate one definite structure for each rank. However through various model construction strategies (e.g. Nuclear Magnetic Resonance (NMR), flash-pyrolysis, X-ray scattering; for a comprehensive review, refer to Mathews and Chaffee 2012), model representations for each coal rank have been determined, as exemplified in Figure 1.2.

5 A B C

D

E

F

Figure 1.2
Physical (A-C) and chemical (D-F) representations of coal in increasing order: Lignite (A and D), bituminous coal (B and E) and anthracite (C and F). Chemical structures D, E and F were adopted from Mallya and Zingaro (1984), Wiser (1984) (with adaptation by Mathews and Chaffee 2012) and Wender (1976), respectively. Images were obtained from the United States Geological Survey. M represents possible metal ion carboxylate interactions in lignite molecules.

Of all coal ranks, lignite has the most variability in its molecular structure. This is due to the presence of an array of aromatic rings linked and cross-linked with various lengths of aliphatic chains (Wender 1976). Lignin, a common compound derived from wood, acts as the precursor to lignite, as indicated by the presence of C3-C6-aliphatic side chains (Hatcher 1990; Hatcher and Clifford 1997; Wender 1976). Oxygen is present in different forms, e.g. carboxyl, alcohol, ether, ester, phenol, ketone groups, that reflect the relatively higher oxygen content in lignite than higher-ranked coals (Wender 1976). Nitrogen- containing groups, e.g. amide, amine, hydroxylamine and indole (Philip et al. 1984), and also those of sulfur e.g. thiol and trisulfides (Gryglewicz and

6 Rutkowski 2001) can also be present in lignite. Further, metal cations may also be included, which can link between phenol and carboxylic acid sites
(Mallya and Zingaro 1984).

As the coal rank increases, however, the various functional groups and aliphatic side chains found in lignite begin to decrease in quantity, and are replaced with aromatic ring structures. This creates a more ‘compact’ molecular structure in bituminous coal compared to the more ‘loosely’ structured lignite. Nevertheless, oxygen is still present in functional groups and/or linking between different aromatic groups, albeit at a lower frequency than in lignite (Given 1960; Shinn 1984). Anthracite is the most aromatically dense among all coals, containing large sheets of aromatic structures with very little to no oxygen-links and functional groups. The increased amount of aromatic structures in bituminous coal and anthracite is also reflected on their physically hard structures, the latter being the hardest among all coals.

Based on the molecular structures of coal, it is expected that lignite is the easiest (i.e. least energy intensive) to biologically degrade compared to the higher-ranked coals. This is due to a higher number of sites, e.g. aliphatic side- chains and functional groups, vulnerable for ‘attacks’ that result in fragmentation of the macromolecule (Hofrichter and Fakoussa 2001). In contrast, the more aromatically dense structure in bituminous and higher ranked coals make them much more impenetrable to biological attack, thereby requiring much more energy and/or time to degrade compared to lignite.

1.2.4 Coal biodegradation

Even at its lowest rank coal consists of complex organic molecules that require either an enormous amount of energy or time to degrade. In the context of microbial degradation of coal, it is highly improbable to find one single microorganism that could completely transform coal to CO2 or CH4, due to the coal molecule complexity and recalcitrance. It is thus very likely that total coal biodegradation involves many different groups of microorganisms, which specifically target a ‘niche’ in certain coal molecules to suit their growth and/or
7 metabolic needs. In light of this, Strąpoć et al. (2008) illustrated a simplified model that shows the different stages of coal biodegradation alongside potential microbes that may be associated with each stage (Figure 1.3). Similar models were also shown by Jones et al. (2010) and Strąpoć et al. (2011b). These proposed schemes were originally created to better understand the processes governing the coal bioconversion to methane, which is seen as the last (desirable) step in coal degradation.

Figure 1.3 Proposed stepwise degradation of coal by Strapoc et al. (2008). Three main stages include 1) aerobic and/or anaerobic fragmentation of coal macromolecule into smaller fragments consisting of aliphatics and aromatics; 2) degradation of the fragmented molecules through fermentation (F) and anaerobic oxidation (AO), and 3) utilisation of the by-products generated from (2), which include acetate, H2 and CO2 by methanogens to generate methane. Labelled numbers in the diagram correspond to the microorganisms thought to be responsible for the reactions.

Based on the decomposition models, coal degradation begins with the fragmentation of the coal macromolecule into smaller entities consisting of aliphatic chains and/or aromatic rings. The smaller structures often contain cellulose and/or lignin derivatives, which include oxygen-containing functional groups e.g. carboxyl, hydroxyl, carbonyl, ester and ketone. These oxygen- containing functional groups and other oxygen-linked moieties create potential
8 sites for activation or cleavage reactions that eventually result in the fragmentation of the coal macromolecule. Whilst the above studies linked this degradation to anaerobic processes (e.g. fermentation) to account for the in situ microbial methane formation, it is possible to initiate the reaction with aerobic fragmentation through the use of fungi and bacteria capable of degrading the geomacromolecule by a number of mechanisms including biosolubilisation and depolymerisation (Strąpoć et al. 2011b). In fact, aerobic biodegradation is likely to accelerate the coal-to-methane bioconversion process due to the presence of oxygen as the terminal electron acceptor, thereby stimulating the process more rapidly than anaerobic degradation alone.

The next step is degradation of the smaller fragmented coal molecules. This can be achieved through anaerobic oxidation or fermentation of the aliphatic and aromatic moieties, although exact mechanisms are not well understood (Jones et al. 2010; Strąpoć et al. 2011b). However, fragmented polymers and monomers are converted to short-chain molecules such as fatty acids, organic acids, hydrogen and carbon dioxide, which act as substrates for the last coal bioconversion process, i.e. methanogenesis.

Methanogenesis is a form of anaerobic respiration observed exclusively within the Archaea. Depending on the substrates, it can be categorised into hydrogenotrophic, acetoclastic or methylotrophic methanogenesis. Hydrogenotrophic (carbon dioxide and hydrogen-utilising) and acetoclastic (acetate-utilising) methanogens dominate coal methanogenesis due to the abundance of the substrates on coal beds (Li et al. 2008; Penner et al. 2010). However, generation of methanol, methylamines and methylsulfides from coal, which feed into methylotrophic methanogenesis is also possible, although less frequently found in coal beds (Strąpoć et al. 2011b; Strąpoć et al. 2011a). The pathways in bioconversion of coal to methane depend greatly on the location, type of coal beds (i.e. coal basins) and physicochemical properties unique to the coal microenvironment (Strąpoć et al. 2010). Different studies have found a major production of one particular substrate over the alternatives (e.g. acetate instead of H2 and CO2 (Beckmann et al. 2011; Jones et al. 2010) or methanol instead of acetate and H2 and CO2 (Guo et al. 2012; Penger et al. 2012)).
9

The rate-limiting step in bioconversion of coal into methane is the initial fragmentation of the complex organic matrix into smaller bioavailable compounds. To enhance this process, it is possible to use bacteria and fungi capable of degrading coal through a variety of mechanisms under aerobic conditions. A considerable number of previous studies have been conducted on the aerobic biodegradation of coal.

1.2.5 Aerobic coal biodegradation

For a very long time, coal was thought to be an unlikely substrate for microorganisms. Traditionally, microbiologists have been using simple sugars such as glucose and organic acids to study microorganisms, hence the use of coal as substrate that is recalcitrant and complex was not immediately considered. Furthermore, direct utilisation of coal has been associated with extreme conditions in the past (e.g. high temperature and pressure), in which microorganisms were not thought likely to participate. Since the discovery of coal-degrading microbes, a considerable number of studies have been conducted on the isolation and/or application of coal-degrading microbes under aerobic conditions (Cohen and Gabriele 1982; Fakoussa 1981).

Various research areas specifically on the microbial interactions with coal are currently actively studied, which include coal desulphurisation, coal fly ash leaching, coal-tar degradation and removal of polyaromatic hydrocarbons from coal-polluted environments (Seidel et al. 2001; Schneider et al. 1996; Sarti et al. 2010; Ghoshal et al. 1996). Although there are different underlying purposes to achieve coal biodegradation, a major motivation in utilising coal-degrading microbes is to satisfy the increasing global energy demand in a more sustainable manner (Fakoussa and Hofrichter 1999; Gao et al. 2012; Haider et al. 2013; Hofrichter et al. 1997b; Hofrichter and Fakoussa 2001). To achieve this aim, several approaches have been taken, including isolating new or applying previously isolated microorganisms; observing and measuring coal degradation through direct observation and analytical techniques; and investigating the possible mechanisms by which coal is (aerobically) degraded.
10 Such studies have been fundamental in understanding the relationship between coal and microbes, essential for any applied coal microbiology study.

1.2.5.1 Aerobic coal-degrading microorganisms

Many microorganisms have been shown to aerobically degrade coal (Table 1.1). Overall, both bacteria and fungi have been reported to degrade coal, although the majority of the coal-degrading microbes reported have been fungi. A noticeably larger number of isolates showed degradation of brown coal (e.g. lignite) as opposed to black coal (bituminous coal), reflecting the difficulty in degrading black coal by most microorganisms. Degradation of both coal types was achieved by different mechanisms (section 1.2.5.2).

1.2.5.1.1 Bacteria

A large portion of coal-degrading bacteria belong to the Bacillus (Gao et al. 2012; Maka et al. 1989; Quigley et al. 1989) and Pseudomonas (Fakoussa 1981; Fakoussa 1988; Fakoussa 1990; Gupta et al. 1990; Machnikowska et al. 2002; Osipowicz et al. 1994) genera. A smaller number of actinomycetes were also shown to degrade coal including Nocardia rubra (Osipowicz et al. 1994) and Streptomyces spp. (Gupta et al. 1988; Strandberg and Lewis 1987b). These bacteria have been shown to either directly solubilise coal in aqueous conditions (Maka et al. 1989; Strandberg and Lewis 1987b), and/or chemically alter the coal structure particularly at the functional groups (e.g. carboxylic and alcoholic molecules) found in coal (Fakoussa 1981; Fakoussa 1990; Gupta et al. 1988; Machnikowska et al. 2002).

Of particular interest is Pseudomonas fluorescens (Fakoussa 1981; Fakoussa 1988), as this is the only species that showed degradation of unmodified bituminous coal. Although there were other bacteria demonstrating hard coal degradation (Osipowicz et al. 1994), the coal had been incubated at 120 oC for 1 h, which likely caused structural changes making it more amenable to degradation. The coal samples from the P. fluorescens studies, however, were

11 ground to increase the coal surface area, but not chemically modified. The P. fluorescens strain was isolated from a forest fire region through approximately 3,100 cultivation experiments using the untreated hard coal. It was shown that the bacteria caused several modifications to the physical coal structure, including a change its colour, wettability and extractability (Fakoussa 1981; Fakoussa 1988; Hofrichter and Fakoussa 2001). This partly resulted from a significant amount of surfactants produced by P. fluorescens, which significantly lowered the coal surface tension (Fakoussa 1988). Furthermore, infrared (IR) and esterification analyses showed an increase in the carboxylic and hydroxyl content of the coal incubated with P. fluorescens, which strongly indicated an oxidative mechanism of coal degradation by the strain. P. fluorescens carries related catabolic activities, including the degradation of polyaromatic hydrocarbons (PAH) such as naphthalene and phenanthrene (Foght and Westlake 1996; Foght and Westlake 1991; Leblond et al. 2001). These aromatic structures are likely to be found abundant in coal and crude oil (Foght and Westlake 1996; Hearn et al. 2003). The ability shown by P. fluorescens to degrade hard coal and other complex hydrocarbons warrants a more fundamental investigation on the bacteria’s relationship with coal, including its attachment behaviour on the coal surface. Such attempts have been made in regard to P. fluorescens interactions with crude oil (Abbasnezhad et al. 2008), but so far not with coal.

1.2.5.1.2 Fungi

Coal-degrading fungi include Basidiomycota (e.g. Phanerochaete chrysosporium (Ralph and Catcheside 1994), Trametes versicolor (Cohen and Gabriele 1982)), Ascomycota (e.g. Fusarium oxysporum (Holker et al. 1995), Neosartoya fischeri (Igbinigie et al. 2008)) and Zygomycota (e.g. Cunninghamella sp. (Stewart et al. 1990; Ward 1990)) phyla. Fewer yeast-like fungi, mainly Candida spp. (Breckenridge and Polman 1994; Kucher et al. 1976; Ward 1985), have also shown coal degradation. Most coal-degrading fungi are filamentous in morphology, a factor that is advantageous for coal biodegradation. Actinobacteria like Streptomyces spp. that have been found to degrade coal (Gupta et al. 1988; Strandberg and Lewis 1987b) are also known
12 for their filamentous characteristics. Due to the porosity of coal, filamentous microorganisms can colonise both the outer and inner parts of the coal for effective degradation.

One of the well-known coal-degrading fungi is Phanerochaete chrysosporium. Phanerochaete chrysosporium is a wood decaying white-rot basidiomycete known for its ligninolytic enzyme system (Farrell et al. 1989; Tien and Kirk 1984; Tien and Tu 1987; Tien and Kirk 1988; Tien and Kirk 1983). The fungus produces a range of extracellular enzymes, including lignin peroxidase (LiP), manganese-dependent peroxidases (MnP) and laccase (Srinivasan et al. 1995; Tien and Kirk 1988; Wariishi et al. 1992), which have been found to depolymerise a wide range of aromatic compounds including lignin (Tien and Kirk 1983), polyaromatic hydrocarbons (Bumpus 1989), biphenyls (Eaton 1985), phenolic compounds (Alleman et al. 1995; Garcia et al. 2000), azo dyes (Spadaro et al. 1992), xenobiotics and other recalcitrant compounds (Cameron et al. 2000). The diversity of enzymes possessed by P. chrysosporium in degrading a wide range of aromatics is mainly due to the ability of LiP and Mn III from MnP to produce diffusible, small free radicals that are able to non- specifically oxidise polymer matrices such as coal (Ralph and Catcheside 1994; Gold et al. 1989). P. chrysosporium has become a key reference in coal degradation tests (Fakoussa and Frost 1999; Igbinigie et al. 2008; Monkemann et al. 1996; Stewart et al. 1990; Torzilli and Isbister 1994). Interestingly, however, no studies have yet shown P. chrysosporium being able to effectively degrade untreated bituminous coal.

As with bacteria, the majority of coal biodegradation studies have focused on low-rank coal (i.e. lignite). Whilst discovering microorganisms with lignite- degrading ability is a major development in its own right, finding microbes that degrade raw bituminous coal is a more challenging undertaking. The first known discovery was made by Monistrol and Laborda (1994), who demonstrated the conversion of untreated hard coal to softer, tar-like appearance by an isolated fungus (identity unknown). Unfortunately, the fungus lost its ability to solubilise hard coal over time, most likely due to spontaneous degeneration of the fungal cells (Hofrichter and Fakoussa 2001). Nevertheless, the authors succeeded in
13 isolating other hard coal degrading fungi (Trichoderma sp. and Penicillium sp.), but the tar-like coal conversion was no longer observed (Laborda et al. 1999; Laborda et al. 1997).

More recently, Igbinigie et al. (2008) discovered a new hard coal degrading fungus, Neosartoya fischeri, which was isolated from the roots of a grass species (Cynodon dactylon) grown on coal dumps. The fungus was able to show heavy engulfment and solubilisation of ground nitric-acid treated bituminous coal just after 3 days of incubation, which was not observed with reference strains P. chrysosporium or Trichoderma versicolor. Interestingly, however, whilst N. fischeri showed effective hard coal degradation, its performance on lower rank coal was lower than those of P. chrysosporium and T. versicolor. This shows a selective mechanism by N. fischeri that favoured hard coal degradation. This also shows that low rank coal degradation is not necessarily achievable by all coal degrading microbes, but rather depends on the availability of the necessary degrading mechanism.

Based on Fourier Transform Infrared (FTIR) and pyrolysis gas chromatography mass spectrometry (GC/MS) analyses of the coal biodegradation products by N. fischeri, an increase of C=O and N=O related compounds were observed, which indicated oxidation and nitration as possible mechanisms of coal alterations by the fungus. Depolymerisation of the coal molecule by the fungus was also speculated, as phenolics and fluorene in the degraded products were observed from the pyrolysis-GC/MS results. This discovery provided valuable information on the possible mechanisms of hard coal degradation, a process less understood than for low rank coal.

14 Table 1.1 Examples of brown (low-rank) and black (high-rank) coal-degrading microorganisms and associated mechanisms of degradation. Adapted from Hofrichter and Fakoussa (2001). Actino.= Actinobacteria, Zygo.=Zygomycota, Depoly.= Depolymerisation, Util.=Utilisation, Solub.=Solubilisation.

Organisms Brown/black coal Mechanism Reference Nocardia rubra Black Depolymerisation (Osipowicz et al. 1994) Actino. Streptomyces spp. Brown Solubilisation (Gupta et al. 1988) Streptomyces setonii Brown Solubilisation (Strandberg and Lewis 1987b) Bacillus sp. Brown Solubilisation (Quigley et al. 1989) Bacillus cereus Brown Solubilisation (Maka et al. 1989) Bacillus licheniformis Brown Solubilisation (Gao et al. 2012; Polman et al. 1994a) Bacillus pumilus Brown Solubilisation (Maka et al. 1989) Bacillus subtilis Brown Solubilisation (Maka et al. 1989)

BACTERIA Pseudomonas aureofaciens Black Depolymerisation (Osipowicz et al. 1994)

Eubacteria Pseudomonas cepacia Brown Depolymerisation (Gupta et al. 1990) Pseudomonas fluorescens Black Oxidation (Fakoussa 1981; Fakoussa 1988; Fakoussa 1990) Pseudomonas putida Brown Solubilisation (Machnikowska et al. 2002) Clitocybula dusenni Brown Depolymerisation (Ziegenhagen and Hofrichter 1998) Coprinus sclerotiger Black Physical disruption (Bublitz et al. 1994) Coriolus hirsutus Black Physical disruption (Bublitz et al. 1994) Nematoloma frowardii Brown Solubilisation, depoly. (Hofrichter and Fritsche 1996) Phanerochaete chrysosporium Brown Solubilisation, depoly. (Ralph and Catcheside 1994; Torzilli and Isbister 1994) Piptoporus betulinus Black Depolymerisation (Osipowicz et al. 1994) Trametes (Coriolus) versicolor Brown Solubilisation, depoly., util. (Cohen and Gabriele 1982; Fakoussa and Frost 1999) Agrocybe praecox Brown Depolymerisation (Hofrichter and Fritsche 1996) Stropharia rugosoannulata Brown Depolymerisation (Steffen et al. 2000) Basidiomycetous Isolates RBS 1k1, RBS 1b1 Brown Depolymerisation (Wilmann and Fakoussa 1997a; Wilmann and Fakoussa 1997b) Poria monticola Brown Solubilisation, utilisation (Cohen and Gabriele 1982) Alternaria sp. Brown Solubilisation (Hofrichter and Fritsche 1997) Aspergillus terreus Brown Utilisation (Ward 1985) Fusarium oxysporum Brown Solubilisation (Holker et al. 1995) Neurospora crassa Brown Solubilisation (Patel et al. 1996) Neosartoya fischeri Black Depolymerisation, oxidat. (Igbinigie et al. 2008)

FUNGI Paecilomyces spp. Brown Solubilisation (Scott et al. 1986; Ward 1985) Penicillium citrinum Brown Solubilisation (Polman et al. 1994a) Penicillium sp. Brown Solubilisation (Kitamura et al. 1993; Laborda et al. 1999) Black Solubilisation (Laborda et al. 1999; Monistrol and Laborda 1994; Stewart et al. 1990) Ascomycetous Trichoderma atroviride Brown Solubilisation (Holker et al. 1997; Holker et al. 1999; Laborda et al. 1999; Silva-

Deuteromycetous and Stenico et al. 2007) Black Oxidation, depoly., solub. (Laborda et al. 1999; Monistrol and Laborda 1994; Stewart et al. 1990)

Candida bombicola Brown Solubilisation (Breckenridge and Polman 1994) Yeast Candida sp. Brown Utilisation (Ward 1985) Candida tropicalis Brown Utilisation (Kucher et al. 1976) Cunninghamella sp. Brown Solubilisation (Ward 1990; Ward 1993) Zygo. Black (Stewart et al. 1990) Mucor lausannesis Brown Utilisation (Ward 1985)
15 1.2.5.2 Mechanisms of aerobic coal degradation

Hofrichter and Fakoussa (2001) described three possible mechanisms of coal degradation: depolymerisation, solubilisation and utilisation. Although originally meant to describe low rank coal (i.e. lignite and coal-derived humic acids) degradation, the mechanisms can also be applied to bituminous coal (Igbinigie et al. 2008; Laborda et al. 1999; Osipowicz et al. 1994; Stewart et al. 1990). Depending on the microorganism and culture conditions, these mechanisms may overlap with each other.

Depolymerisation occurs through the molecular fragmentation of coal that generates compounds with lower molecular masses. In some cases, decolourisation of degraded coal could also be observed as a result of depolymerisation (Hofrichter and Fritsche 1997). The mechanism is often associated with enzymatic reactions at low pH values (pH 3-6), most notably by the actions of the oxidative enzymes MnP, LiP and laccase (Hofrichter and Fritsche 1996; Hofrichter and Fritsche 1997; Scheel et al. 2000; Wilmann and Fakoussa 1997b; Wondrack et al. 1989). These enzymes work by cleaving certain coal structures (including aromatic rings) susceptible to their non- specific oxidative attack, leading to the generation of smaller coal fragments.

Solubilisation is a process where coal is converted from a solid state to liquid (Figure 1.4), usually driven by non-enzymatic processes and at a higher pH (7- 10). Microbial production of alkaline substances and chelating agents are usually associated with this mechanism (Gao et al. 2012; Quigley et al. 1989; Strandberg and Lewis 1987a; Strandberg and Lewis 1987b). This could occur + through the production of alkaline metabolites, e.g. ammonium ions (NH4 ) or amines (R-CH2-NH2) in response to a high amount of nitrogen in the medium, or the utilisation of organic acids (e.g. citric, oxalic) that results in an increased amount of free bases, thereby leading to alkalinisation (Hofrichter and Fakoussa 2001).

Despite usually being non-enzymatic, coal solubilisation has been associated with hydrolytic enzymes such as esterase (Gao et al. 2012; Holker et al. 1999;
16 Laborda et al. 1999). Further, studies have shown that the presence of MnP, which is predominately linked with depolymerisation, also plays a role in solubilisation (Gao et al. 2012; Laborda et al. 1997).

Figure 1.4 Example of coal (lignite) solubilisation by Alternaria sp. pre-grown on an agar plate. Image from Hofrichter and Fakoussa (2001).

It is important to note that almost all coal depolymerisation and solubilisation studies involved the use of another carbon source (e.g. glutamate, glucose), particularly to sustain the fungal growth (Holker et al. 1999). Direct observations on coal solubilisation were often made on fungi that were grown on a rich medium (e.g. Sabouraud agar) with depolymerisation or solubilisation, driven by co-metabolism (Cohen et al. 1987; Igbinigie et al. 2008; Monistrol and Laborda 1994). Utilisation of coal, in contrast, refers to microbial growth on coal as sole carbon source (Gupta et al. 1990; Kucher et al. 1976; Ward 1985). Coal-utilising microbes are able to obtain low-molecular mass aromatics (e.g. phenols) and aliphatics (e.g. n-alkanes) from coal and derive energy for growth (Gupta et al. 1990; Hofrichter and Fakoussa 2001; Kucher et al. 1976; Ward 1985). Furthermore, coal utilisation is a slow process because of the chemical complexity of coal. However this can be enhanced through the addition of mineral solutions (e.g. N, P, S, and mineral salts) (Hofrichter and Fakoussa 2001) and use of pre-treated coal (Ward 1985).

Apart from the three main mechanisms described, several other mechanisms exist for bituminous coal degradation. These include oxidation and biosurfactant production. Coal oxidation was observed by Fakoussa (1981), Fakoussa
17 (1988), Fakoussa (1990) and Igbinigie et al. (2008) as an increase in oxygen functionalities in coal (e.g. C=O) after microbial treatment, possibly through enzymatic systems (Dong et al. 2005; Ensley et al. 1982; Jeffrey et al. 1975; Leahy et al. 1996). Oxidation increases the bioavailability of certain compounds in coal, thereby making it less challenging for the microbes to degrade and/or utilise coal as a substrate (Hofrichter and Fakoussa 2001; Rosenberg and Rosenberg 1981). Surfactant production, such as those seen in P. fluorescens, considerably decreases the coal surface tension (Fakoussa 1981; Fakoussa 1988; Polman et al. 1994b), thereby increasing the bioavailability and consequently degradation of coal. Positive correlation between biosurfactant production and degradation of hydrocarbons (e.g. phenanthrene) has also been previously demonstrated (García
Junco et al. 2001), although this cannot be generalised for all hydrocarbon-degrading species (Johnsen and Karlson 2004; Willumsen and Karlson 1996).

All the above mechanisms described the chemical or biochemical factors produced by the microbes that influence coal biodegradation. Whilst these factors are important, attention should also be given to physical mechanisms by microorganisms that lead to coal degradation. A number of studies have mentioned the role of fungal hyphae in colonising both the inner and outer parts of coal (Achi and Emeruwa 1993; Cohen and Gabriele 1982; Igbinigie et al. 2008; Monistrol and Laborda 1994; Mukasa-Mugerwa et al. 2011). Tight attachment between hyphae and coal was observed, which eventually lead to coal being physically fragmented into smaller pieces (Hofrichter et al. 1997b). In some cases, ‘erosion’ of bituminous coal pieces was seen, e.g. by the basidiomycete Coprinus sclerotiger, which caused liberation of powder-sized particles and separation of the asphaltene layer (shiny material) of coal (Bublitz et al. 1994). Mass spectrometry analyses showed that the liberated coal material contained volatile and aliphatic (i.e. n-hexane) fractions of coal (Hofrichter et al. 1997b). Hence, this suggested a link between chemical alterations of coal with physical fragmentation caused by the fungus, only after fungal attachment and colonisation on coal. This showed the significance of cell adhesion on coal towards achieving effective coal degradation.

18 Many of the above studies suggest microbial colonisation of coal leads to coal degradation, however, there is still currently little information on this aspect. No studies have investigated the role of cell adhesion and surface colonisation of coal in detail. To understand cell adhesion and colonisation of coal, a general background in cell attachment, colonisation and biofilm formation is required. The next section of this review covers fundamental aspects of microbial interaction with surfaces.

1.3 Biofilms – Cell adhesion and microbial surface colonisation

For a long time, microorganisms were viewed as simple organisms that lived unicellularly without any complex multicellular interactions. However, this perception changed with the discovery that microorganisms, apart from being able to live as single cells, can attach to surfaces and form complex, multicellular units. The formation of aggregated cells embedded in polymer matrices and highly structured communities are known as ‘biofilms’ (Costerton et al. 1995; Costerton 1999a; Hall-Stoodley et al. 2004).

Ubiquitous in nature, biofilms can form on almost all types of surfaces provided sufficient moisture and accessible nutrients are available (Costerton et al. 1995). Cells are thought to adhere to surfaces and form biofilms as a form of protection against physical, biological or chemical attacks in the environment (Hall-Stoodley et al. 2004). Further, cells living as biofilms have the advantage of performing tasks that would otherwise be ineffective if conducted unicellularly. Many persistent and chronic bacterial infections, for example, depend on the effectiveness of the pathogens to grow as biofilms than single cells (Costerton 1999b).

Biofilms are widely recognised in various fields including disease and infections (Geipel 2009; Hawser 1994), biofouling (Shikuma and Hadfield 2010), bioprocessing (Wu et al. 2009), microbial fuel cells (Rabaey et al. 2007; von Canstein et al. 2008), steel corrosion (Mehanna et al. 2009; Zhang et al. 2007), wastewater treatment (Gabr et al. 2009) and in natural environments such as

19 streams and rivers (Besemer et al. 2009a; Luef et al. 2009) and hydrothermal vents (Westall et al. 2001). This shows the significance of biofilms in a diverse range of environments, which are either beneficial (e.g. in maintaining the ecosystem and through biotechnological applications) or destructive (e.g. infections and diseases) to humankind.

1.3.1 Biofilm formation

Biofilms form through a number of sequential stages (Figure 1.5), which are: 1) reversible cell attachment, 2) irreversible cell adhesion, 3) microcolony formation, 4) biofilm maturation and 5) dispersal. Both the beginning and end of a biofilm cycle consist of individual, free-living (planktonic) cells that result in continuous colonisation of new surfaces. The biofilm formation model is useful as a general guide to understanding the main mechanisms of cell attachment and surface colonisation. However, variations in terms of length of each stage and colonisation behaviour are to be expected across different microbial species, communities and/or environments. The biofilm model generally focuses on bacterial biofilm development, which differs from that of fungi. Thus a separate section on fungal biofilm formation is discussed in section 1.3.2.

Figure 1.5 Stages in biofilm formation: 1) Initial (reversible) cell attachment by free-living cells, 2) Adhesin production by attached cells (‘irreversible’ adhesion), 3) Microcolony formation, 4) Biofilm maturation and 5) Biofilm dispersal, where individual cells are released and restart a new biofilm cycle. Micrographs (1-5) consist of Pseudomonas aeruginosa biofilm development that correspond to the biofilm stages. Images adapted from Stoodley et al. (2002).
20

The first form of contact between cells and a surface, termed ‘reversible’ adhesion, begins by single cells ‘loosely’ attaching to the surface. This occurs mainly as a result of physico-chemical factors (section 1.3.4.2), which include van der Waals forces, hydrophobicity, surface free energy, electrostatic forces and surface roughness (Busscher and Weerkamp 1987; Busscher et al. 1990; Quirynen and Bollen 1995; Van Loosdrecht et al. 1990; Van Loosdrecht et al. 1987a). These factors are only effective once the cell is close enough (within 50 nm of distance) to the surface. Thus, cells are first transported towards the surface through a number of different modes, including diffusion by Brownian motion, convection through liquid or air flow, and/or chemotactic activity by the microorganism (e.g. flagellar-mediated motility) (Busscher and Weerkamp 1987; Quirynen and Bollen 1995). Only favourable physico-chemical interactions between the cell and substratum would result in its initial attachment to the surface before further adhesion takes place.

Once a particular surface is found suitable for further attachment, cells use adhesive substances (i.e. ‘adhesins’) that allow them to permanently attach to the surface. This attachment stage is called ‘irreversible’ adhesion as the cells are now committed to colonising the surface for biofilm development (Costerton et al. 1995). Different types of adhesins are produced by the adhering microbes, including exopolymeric substances (EPS), pili and fimbriae, which constitute a part of the biological factors promoting cell adhesion. The adhesins ensure a firm anchorage of the cells onto the surface, hence preventing any potential desorption by physico-chemical forces (i.e. shear force). At this stage, cells that are attached to the surface form a monolayer of biofilm, which serves as the basis for further attachment and early biofilm growth.

Subsequent microcolony formation consists of small clusters of cells adjoining to each other forming a multilayer biofilm. The microcolonies are developed via several ways, including multiplication (growth) of the originally adhered cells, attachment of new cells to the established monolayer and/or co-adhesion of cells in mixed-species population (Hall-Stoodley and Stoodley 2002; Quirynen and Bollen 1995; Stoodley et al. 2002; Tolker-Nielsen et al. 2000). In an
21 aqueous environment, certain cells are able to mobilise themselves on the surface to co-aggregate with other cells. Different motility behaviours possessed by bacteria (e.g. type IV pili in Pseudomonas aeruginosa, type I pili in Escherichia coli) allow the expansion of the microcolonies into different shapes and sizes depending on the species involved (Jackson et al. 2002; Klausen et al. 2003; O'Toole and Kolter 1998). Surface properties, including surface roughness, are also influential in microcolony development (Hall-Stoodley and Stoodley 2002; Quirynen and Bollen 1995).

Microcolony expansion progresses into a mature biofilm, which consists of cells encased in a highly structured and complex microbial ‘edifice’. A mature biofilm contains structures that resemble channels and pillars, which are thought to help transport nutrient and oxygen to the tightly packed cells, and efficiently remove waste material (Costerton et al. 1995; Davey and O'toole 2000; Tolker- Nielsen et al. 2000). Mature biofilms not only consists of cells, but also carbohydrates, exopolysaccharides, proteins, extracellular DNA (eDNA), RNA, cell waste, ions and other matrix-soluble substances, all of which are encased in a polymer matrix called exopolymeric substance (EPS) (Sutherland 2001a; Sutherland 2001b). Many of the biofilm components (e.g. exopolysaccharides and eDNA) play a vital role in ensuring the biofilm structural integrity and stability (Danese et al. 2000; Karatan and Watnick 2009; Whitchurch et al. 2002), thus protecting it, to a certain extent, against various chemical, biological and physical threats (Liu and Tay 2002; Mah and O'Toole 2001; Matz et al. 2005). Further, cell-to-cell signalling (i.e. quorum sensing) occurs actively at this stage, in which cells produce signalling molecules (e.g. N-acyl homoserine lactone) that regulate a diverse range of physiological needs (e.g. motility, metabolite production, stress response) to ensure a fully functional biofilm.

Biofilm formation ends at a stage called ‘dispersal’, where cells inside the biofilm are released into the environment as planktonic cells. Various factors can cause biofilm dispersal, which includes toxicity, limited oxygen and nutrient availability and increased shear stress (Barraud et al. 2006; Purevdorj et al. 2002; Schleheck et al. 2009; Thormann et al. 2005). The dispersal is induced by different mechanisms by the biofilm such as cell signal-mediated inhibition of
22 further biofilm growth or enzyme production that degrades biofilm components (Boles and Horswill 2008; Hammer and Bassler 2003; refer to McDougald et al. 2011 for a comprehensive review on biofilm dispersal). The dispersed cells would then search for other surfaces to colonise and begin a new biofilm formation cycle.

Amongst all stages in biofilm formation, it is clear that a key-determining factor to surface colonisation is the initial attachment of cells, which are primarily governed by physico-chemical factors. Cells need to overcome various physico- chemical obstacles (e.g. surface free energy and hydrophobicity) in order to reach a surface, where only then it is able to fully implement its own mechanisms for further adhesion. Thus, it is important to consider the non- biological influences apart from the biological roles in determining the initial cell attachment, from which biofilms can be formed.

1.3.2 Fungal biofilms

Although similarities exist between bacteria and fungi in biofilm formation, the bacterial biofilm model can only be used to a certain extent to study fungal surface colonisation due to several differences in morphology, physiology and growth. There are different opinions on whether fungal surface colonisation can be described as fungal biofilms, particularly since fungi are largely found in air interfaces instead of submerged in liquid, and that filamentous fungi typically have invasive hyphae that penetrate through the substratum, which most bacteria lack (Harding et al. 2009). However, for the purpose of this review, the term ‘biofilm’ is used to describe microbial surface colonisation in both air and liquid environments, and for both bacteria and fungi, which is widely accepted (Harding et al. 2009).

Fungi possess absorptive nutrition, extracellular enzymatic secretion and apical hyphal growth (Jones 1994). Many fungi are dimorphic and are capable of switching their growth form quickly in response to different environmental stresses (Chen and Fink 2006). These properties make fungi very adaptive in the environment, particularly in colonising a wide range of surfaces including
23 plants and building materials (Blanchette et al. 2010; Gaylarde and Morton 1999; Harding et al. 2009; Macia-Vicente et al. 2008).

Jones (1994) has written an extensive review on fungal adhesion, which focuses on the biological mechanisms of initial cell attachment. Following this, and general (bacterial) biofilm development (section 1.3.1), Harding et al. (2009) described a model for filamentous fungal biofilm formation (Figure 1.6), which involves the following stages: i) Propagule adsorption: Spores or hyphal fragments (equivalent to single cells for bacterial planktonic growth) reversibly attach to a surface through physico-chemical attractions; ii) Active attachment: The secretion of adhesin for irreversible attachment; iii) Microcolony formation I: This stage consists of apical growth and branching of the hyphae to produce a monolayer network of hyphal cells. EPS is also produced here and serves as an envelope for the cells; iv) Microcolony formation II: This is an initial maturation stage during which hyphal cells are dense and interjoined with each other in a complex hyphal network; v) Maturation: This stage is noted by the production of fruiting bodies and other reproductive structures; vi) Dispersal: This phase is analogous to bacterial biofilm dispersal, during which cells detach themselves from the biofilm. It involves the dispersal of spores or biofilm fragments into the air/liquid bulk phase, consequently restarting the biofilm cycle.

Figure 1.6 Biofilm formation stages for filamentous fungi. Ramage et al. (2009) extend the general biofilm model to describe the fungal biofilm formation as follows: i) reversible propagule (spore) attachment and ii) irreversible (active) attachment, iii) germling formation to form monolayer biofilm, iv) extensive mycelial and hyphal colonisation, v) maturation, i.e. development of fruiting bodies, vi) dispersal stage during which spores are disseminated from the biofilm. Image adapted from Harding et al. (2009).

24
Based on the bacterial and fungal biofilm formation models, it is evident that a major difference between the two biofilms is the ability for filamentous fungi to form extensive hyphae for colonisation, whilst limited cell extension can be observed for bacteria. Filamentous fungi are able to use hyphae to colonise the outer substratum surface and penetrate the surface of the colonised area. This is advantageous for fungi in its various roles in the environment including as pathogens, saprophytes or endophytes, where deep penetration of hosts is necessary for efficient infection, nutrient uptake and/or biofilm stability (Jones 1994; Macia-Vicente et al. 2008; Perera et al. 1997). This can also be extended to other environments including deteriorated buildings, where penetrative fungal colonisation ensures its persistence even in harsh environments (Warscheid and Braams 2000).

1.3.3 Environmental biofilms

In natural environments, biofilms often exist as complex, multispecies communities, rather than single species microorganisms (for a comprehensive review on the ecological role of biofilms, see Davey and O'toole 2000). The majority of biofilm investigations have focused on using pure microbial species to study its biofilm development, particularly due to its relative simplicity compared to mixed-community biofilms. Although the former is essential for a fundamental understanding of biofilms, it does not reflect the true nature of biofilms and their associated complexity in the natural environment.

Studies have shown the importance of biofilms in a wide range of natural settings including in soil, marine and benthic environments, and higher-order organisms (see biofilm introduction, section 1.3), where they exist as multispecies and multi-organism communities. It is now becoming increasingly apparent that communities in natural biofilms do not attach and form aggregation due to random occurrences, but rather to serve a specific role that may benefit itself and the community as a whole. For example, in activated sludge or sediment flocs (aggregated self-attached cells), it is believed that the presence of H2-producing acetogens and H2-consuming methanogens work
25 together to play a vital role in the anaerobic digestion of organic matter (Conrad et al. 1985; Macleod et al. 1990). Such mutualistic cooperation between biofilm members is not always present, however, as antagonistic behaviours e.g. competition and predation are also evident (Burmolle et al. 2006; Burmolle et al. 2007; Rao et al. 2005; Rypien et al. 2010). Environmental factors including temperature, seasonal weather, water flow and pH (Aguilera et al. 2007; Besemer et al. 2007; Besemer et al. 2009a; Besemer et al. 2009b; Horn et al. 2003) influence the biofilm community and structure and further add to the complexity in studying natural biofilms.

1.3.4 Factors that influence cell attachment and biofilm formation

1.3.4.1 Biological factors

Biological influences play an essential part in the biofilm formation, beginning from the irreversible surface attachment to dispersal of biofilm. Microbes utilise various cell structures and substances, including extracellular polysaccharide (EPS), pili and flagella (for bacteria) and hyphae (for filamentous fungi) that significantly affect their attachment and colonisation on surfaces (Costerton et al. 1995; Jones et al. 2007; O'Toole et al. 2000; Stoodley et al. 2002). Molecular genetic analyses showed that the production and use of these appendages involve numerous regulatory pathways, which greatly depend on environmental signals such as oxygen, nutrients, temperature, hydrodynamics and osmolarity (for reviews on molecular genetics of biofilms, see Davey and O'toole 2000; Molin and Tolker-Nielsen 2003; O'Toole et al. 2000 and Stanley and Lazazzera 2004). Further, it was shown that cell-to-cell signaling, i.e. ‘quorum sensing’ is essential in regulating biofilm formation through various regulatory pathways, which involve small-molecule autoinducer signalling mechanisms (Camilli and Bassler 2006; Davies et al. 1998). These genotypic and phenotypic regulations play an integral part in both the structure and function of biofilms.

26 1.3.4.2 Physico-chemical factors

Established physico-chemical theories have been used to describe cell adhesion, which hold prominence in determining the initial cell attachment to surfaces (Bos et al. 1997; Bos et al. 1999; Busscher and Weerkamp 1987; Busscher et al. 1990; Van Loosdrecht et al. 1990; Van Loosdrecht and Zehnder 1990). It is important to note that, as with all theories and models that describe biofilm formation, the degree of influence of these factors on cell attachment are varied across different microorganisms and environments. However, the following models are widely accepted and validated as a powerful tool in understanding the physico-chemical mechanisms of cell adhesion.

To reach a particular surface, cells must overcome several energy barriers in the water or air medium that are governed by various physico-chemical elements, depending on the distance between the cell and substratum. Movement of cells at a distance of > 50 nm away from the surface are mainly governed by Brownian motion, air or water convection, or flagellar activity (Busscher and Weerkamp 1987). Within 50 nm cells must overcome two more barriers, termed ‘secondary minimum’ and ‘primary minimum’ that are determined by long-range (< 50 nm) and short-range (< 2 nm) interactions.

1.3.4.2.1 Long-range interactions (< 50 nm)

In accordance to the ‘DLVO’ theory (i.e. ‘Derjaguin, Landau, Verwey and Overbeek’), long-range interactions mainly comprise of Lifshitz-Van der Waals,

GA, and electrostatic forces, GE. These forces make up the total Gibbs energy of interaction, GTOT, where a negative value would result in favourable long- range interactions. Three Van der Waals forces have been identified, i.e. induced dipole-induced dipole (London dispersion), induced dipole-dipole (Debye interaction) and dipole-dipole (Keesom interaction) that would occur between two surfaces. These forces are usually favourable between surfaces.

The electrostatic interaction, however, is largely determined by the surface charge and ionic strength of the medium. Opposite charge of two surfaces
27 would cause favourable electrostatic interaction, whilst surfaces with the same charge would repel each other. In nature, microbial cells and surfaces are predominantly negatively charged (Quirynen and Bollen 1995), therefore on this basis repulsion is expected. However, the distance between the cells and surface is greatly minimised at high ionic strength of the medium, hence weakening the repulsion. Through favourable van der Waals interactions, and at a high ionic strength, the total Gibbs energy of (long-range) interaction, GTOT is reduced, thus enabling the cell to reach the primary minimum.

1.3.4.2.2 Short-range interactions (< 2 nm)

Once in primary minimum range, short-range forces such as hydrophobicity, hydrogen bonding, steric interaction and bridging interaction are present to determine the adhesion strength between the cell and the substratum surfaces (Quirynen and Bollen 1995). In an aqueous medium, a water barrier between two interacting surfaces exists and must be removed in order for direct contact to occur. This water barrier can be easily removed by strong hydrophobic interactions, which is determined by the hydrophobicity of cell and substratum surfaces (Boks et al. 2008). Higher cell and/or substratum surface hydrophobicity would generally result in stronger hydrophobic interactions. The barrier can also be overcome by cell appendages (e.g. pili, fimbriae) that act as ‘bridges’ for cell attachment to occur (Boks et al. 2008).

When a cell finally reaches direct contact with the substratum, the free energy of adhesion



, determines whether or not cell adhesion is favourable. The


is the thermodynamic energy balance between the interfacial surface energies of all the interfaces involved (Absolom et al. 1983). This is given by:














(1)

where

 ,

 and

are the interfacial energies for the bacterium-solid (substratum), bacterium-liquid, and solid-liquid interfaces, respectively (the same equation can be derived at air interface:















, where

28

,

 and

 are cell-solid, cell-vapor and solid-vapor interfacial energies, respectively).

A favourable adhesion between the cell and substratum is given when



is negative, whereas positive



would result in unlikely adhesion based on thermodynamic principles. Under several different models, the interfacial energies for the above equation can be derived through experimental work on contact angle measurements based on the Young equation (Young 1805), which is further described in the methods section in Chapter 2 (section 2.2.8). One of the most frequently used models is the van Oss approach (Van Oss et al. 1986), which asserts that the major components of interfacial energies are Lifshitz-van der Waals and polar acid-base interactions, which are most important for thermodynamic balance (Busscher et al. 1990). This model further divides the acid-base component into electron donor and electron acceptor capacities to further describe the adhesion thermodynamics (Bellon-Fontaine et al. 1996). The surface thermodynamics model has been empirically tested and validated and known to be a powerful tool to predict microbial adhesion to solid substrata (Bos et al. 1999; Busscher et al. 1984).

1.3.5 Methods in studying cell attachment and biofilm formation

Given the diverse nature of biofilm studies, various methods have been developed to address numerous aspects of biofilm development (for reviews on common biofilm methods see Bos et al. 1999; Christensen et al. 1999 and Costerton et al. 1995). Physico-chemical approaches, microscopy and molecular techniques are most useful in this study and are discussed further in this section.

1.3.5.1 Physico-chemical approaches

Three different models are used to describe the strength of microbial adhesion to surfaces,



. These are termed the ‘thermodynamic’ approach (i.e. ‘equation of state’), ‘classical DLVO’ and ‘extended DLVO’ models (for a full

29 review on physico-chemical methods see Bos et al. 1999). All three models depend on the Young equation (see section 2.2.8) to generate



, however differences exist in the derivations used, which are briefly explained in this section.

The thermodynamic approach is the simplest model; it is based on short-range interactions to derive



and therefore does not include electrostatic interactions. In this model, the interacting surfaces are already assumed to be in contact (i.e. surpassing the secondary and primary minima) under conditions of thermodynamic equilibrium. The classical DLVO theory is the sum of Lifshitz- van der Waals and electrostatic interactions, and are distance-dependent due to the ionic strength and surface charge (section 1.3.4.2.1). Both models are valid for predicting microbial adhesion, however, they do not cover all possible aspects that influence adhesion for every type of strain, and therefore are limited only to certain microbes that fit under their model.

To accommodate this limitation, Van Oss et al. (1986) developed a third model, i.e. the extended DLVO, which combines the first two approaches. Under this model, the inclusion of the short-range Lewis acid-base interaction is essential in predicting



. The acid-base interactions are based on the electron- donating and electron-accepting capacities between polar moieties of the two interacting surfaces. It has been shown, theoretically (using the Hamaker constant), that the influence of the acid-base component on cell adhesion is far greater than that of Van der Waals and electrostatic interactions (Bos et al. 1999). Based on this notion, the Van Oss model can be used to calculate the surface energy components under the thermodynamic approach, where the distance-dependent interaction (i.e. electrostatic) is neglected, but includes the additional parameters van der Waals, acid-base and hydrophobic interactions. This improved model has been shown to be the most consistent across various bacterial and fungal species (Sharma and Rao 2002), and thus is most useful for the purpose of this study. Under this model, contact angle of three different liquids (i.e. water, formamide and diiodomethane) are measured on microbial lawns and substratum (i.e. coal) to derive the necessary surface energy components. Conveniently, the contact angle of water measured on surfaces is
30 widely used as a measurement of hydrophobicity (Abbasnezhad et al. 2008; Chau et al. 2009; Dorobantu et al. 2004; Ista et al. 2004; Webb et al. 1999) and thus was employed in this study.

1.3.5.2 Microscopy

Several microscopy techniques can be utilised to visualize the structural development of biofilms. These include Confocal Laser Scanning Microscopy (CLSM), Scanning Electron Microscopy (SEM) and Transmission Electron Microscopy (TEM).

CLSM is currently one of the most common visualisation aids in biofilm studies, where in conjunction with an image analysis software, real-time in-depth 3D analysis of the biofilm can be made, making it superior to conventional optical microscopy. Quantitative data on biofilm structure, e.g. density, average height and surface coverage can be obtained by using CLSM, which are useful particularly in studying the overall development of single-species biofilms (Lawrence et al. 1991; Moller et al. 1998; Tolker-Nielsen et al. 2000). However, a major drawback of CLSM is the limited resolution (up to 0.2 µm) in the images obtained, which makes detailed observation on individual cell attachment and morphology difficult.

Electron microscopy (SEM and TEM) has been used in various cell attachment and biofilm studies to produce images of high resolution (up to 0.05 nm) and magnification (up to 200,000x for SEM and 1,000,000x for TEM), with 3D imaging in a 2D projection (Adam et al. 2002; Aguilera et al. 2007; Baum et al. 2009; Dheilly et al. 2008; Grishin and Tuovinen 1989; Kawarai et al. 2007). Clear images can be produced through SEM, particularly on the morphology and attachment of individual cells on surfaces and each other (Hazrin-Chong and Manefield 2012). Further, detailed attachment and/or colonisation of cells on isolated area of uneven surfaces (e.g. crevices, small fissures found on coal) can be observed, which are not possible with any other microscopy techniques. This makes SEM a powerful tool for the purpose of this study, where a simple

31 technique on specimen preparation has been developed (Hazrin-Chong and Manefield 2012).

1.3.5.3 Molecular techniques in analysing biofilm communities

Recent advances in molecular based approaches have made it possible to investigate microbial diversity in complex and heterogeneous environments. This is seen as a major leap forward in microbial ecology, as it was previously impossible to properly estimate the diversity of microorganisms due to the large majority of microorganisms being non-cultivable (Amann et al. 1995; Giovannoni and Stingl 2005; Ward et al. 1990). Since the advent of the polymerase chain reaction (PCR), a number of molecular techniques have been developed that utilise the 16S and 18S rRNA genes. These methods have significantly increased our understanding in the diversity and role of microorganisms in different environments, including biofilms (Besemer et al. 2009b; Burmolle et al. 2007; Lee et al. 2008; Moss et al. 2006).

Among the commonly used PCR-based microbial community approaches are the Terminal Restriction Fragment Length Polymorphism (T-RFLP) and 454 pyrosequencing. T-RFLP is a microbial fingerprinting technique used to obtain community diversity profiles of environmental samples. Other similarly aimed methods include Denaturing Gradient Gel Electrophoresis (DGGE), Automated Ribosomal Intergenic Spacer Analysis (ARISA) and Single Strand Conformation Polymorphism (SSCP) (for a critical review of all methods, refer to Anderson and Cairney 2004 and Nocker et al. 2007). T-RFLP uses an automated DNA sequencing to separate fluorescently labelled PCR products digested with a restriction enzyme (Liu et al. 1997). A restriction fragment profile for each sample is thus obtained based on the size and fluorescence intensity of the fragments. This allows distance-based calculations of the fragments to be made across all samples for powerful data visualisation and statistical analysis (Bates et al. 2011). In comparison to gel-based methods like DGGE and SSCP, the use of automated sequencing in T-RFLP makes the latter more favourable for high-throughput analyses. Further, T-RFLP utilises only the terminal fragments and not the whole amplicons (e.g. in ARISA), thus making it less complex whilst
32 retaining high sensitivity and accuracy. On this account, the biofilm community analysis conducted in this study used T-RFLP as a fingerprinting method of choice.

Nevertheless, a major limitation of T-RFLP is its inability to generate taxonomic identification from its fragments. Although it is possible to perform in silico digestion analysis and assignment of peaks through database comparison, the approach is error-prone due to several assumptions (Anderson and Cairney 2004). Thus, to overcome this limitation, a high-throughput sequencing technique, i.e. 454 pyrosequencing was employed. 454 pyrosequencing, along with other sequencing methods such as Illumina and SOLiD, are referred as ‘next-generation’ (NGS) sequencing (for a recent review, refer to Soon et al. 2013). In contrast to the Sanger method, which involves chain-termination sequencing, NGS techniques involve large-scale parallel sequencing, capable of generating thousands to millions of sequences simultaneously (Hall 2007). In particular, 454 pyrosequencing utilises an emulsion method for DNA amplification, with each droplet containing a bead that carries ten million copies of a unique DNA template (Margulies et al. 2005). DNA fragments carried by the beads are denatured into single-stranded DNA after the emulsion is broken, and are deposited into pico-litre wells where sequencing takes place. 454 pyrosequencing is able to sequence up to 25 million bases in a relatively short time (i.e. one 4-hour run). Its high accuracy, large throughput and relative affordability make it a popular choice across various studies in life sciences.

1.4 Microbial adhesion and colonisation on coal

Little is known about microbial cell attachment and colonisation on coal. Whilst mechanisms of coal degradation have been intensively studied (section 1.2.5.1), a more fundamental knowledge on the microbial colonisation of coal is currently missing.

Coal biodegradation studies have previously mentioned evidence of biofilm formation on coal in a number of ways. The most often cited observation is the

33 intense colonisation of coal by coal-degrading fungi through their extensive hyphae formation (Achi and Emeruwa 1993; Cohen and Gabriele 1982; Hofrichter et al. 1997b; Monistrol and Laborda 1994; Mukasa-Mugerwa et al. 2011). Colonisation was not observed on other materials that acted as controls (i.e. stones), demonstrating that the coal colonising activity observed was active and preferential (Monistrol and Laborda 1994). In all studies, the colonisation was followed by an intense degradation of coal through solubilisation or depolymerisation. Further, gelatinous material characteristic of exopolymeric substance (EPS) production in biofilm formation was also observed on coal through fungal colonisation in submerged conditions (Igbinigie et al. 2008; Laborda et al. 1999), which shows the role of coal as substratum for attachment and biofilm formation by these fungi. The examples above show the important role of cell attachment and biofilm formation in coal degradation; however, no investigations were conducted to further explore this aspect.

More elaborate studies pertaining to cell attachment to coal have been conducted, particularly in the area of coal processing technology. Coal flocculation, desulphurisation and mineral leaching have explored the use of certain microorganisms that specifically adsorb to non-carbon components in coal (e.g. sulphur, fly ash, titanium) (Chen and Skidmore 1988; Ding 2009; Pawlik et al. 2004; Shabtai and Fleminger 1994; Vijayalakshmi and Raichur 2002; Vijayalakshmi and Raichur 2003). Sulphur-oxidising bacteria Sulfolobus acidocaldarius and Thiobacillus ferrooxidans, for example, have been shown to adsorb to coal by selectively adhering to the pyritic sulphur component (Grishin and Tuovinen 1989; Ohmura et al. 1993b; Ohmura et al. 1993a; Vitaya and Toda 1991). This enhances coal recovery since sulphur, a contaminant in coal combustion, can be effectively removed by these microbes. Useful findings on the kinetics of adsorption (i.e. via Langmuir isotherm) and surface thermodynamics have been found (Chen and Skidmore 1988; Ding 2009; Vijayalakshmi and Raichur 2002). However, the above studies specifically focused on mineral obtainment from coal, which differs from studies on coal biodegradation where carbon is of interest. Further, very low pH (< pH 2) and/or high temperature (> 70oC) were necessary for bacterial growth in these studies, which do not reflect the optimum conditions for activity by aerobic coal-
34 degrading microorganisms. Thus, accurate representations of cell adhesion on coal as a carbon substrate are not available.

Interestingly, whilst there is a clear gap of knowledge on cell attachment to coal, extensive research on similar grounds have been conducted for other hydrocarbons, particularly crude oil (for a comprehensive review see Abbasnezhad et al. 2011). Crude oil and coal share a number of similarities as hydrocarbons. They are both low in bioavailability (i.e. degree of interaction with microorganisms) due to their low solubility in aqueous solution, and both contain a large amount of polyaromatic and aliphatic hydrocarbon molecules, which are target substrates for hydrocarbon-utilising microbes.

Cell attachment studies on common substances in crude oil, e.g. hexadecane and phenanthrene, have shown important findings on the role of cell adhesion in liquid hydrocarbon degradation. A common result was that cell attachment and biofilm formation played a significant role in the uptake of the hydrocarbon molecules by certain microbes (García
Junco et al. 2001; Johnsen and Karlson 2004; Rosenberg and Rosenberg 1981; Rosenberg et al. 1982; Wick et al. 2002). There were cases, however, where cell attachment did not play a major role in degradation, particularly when high concentration of oil-emulsifying substance e.g. biosurfactants were produced by the cells, or the solubility of the substrate was high. This shows a greater dependency for microbial attachment to degrade hydrocarbons when they lack extracellular hydrocarbon-degrading mechanisms and/or when the solubility of the substrate is low. Extensive studies have also been conducted on mechanisms for cell attachment on crude oil, which include cell surface hydrophobicity, surface charge and EPS production (Abbasnezhad et al. 2008; Obuekwe et al. 2009). These studies showed the importance of cell attachment in understanding microbial hydrocarbon degradation. Thus, similar investigations on cell attachment on coal should be performed to gain a deeper appreciation of the mechanisms of coal degradation as a solid hydrocarbon. This is an exciting area of research and would broaden the current knowledge of microbial interactions with hydrocarbons.

35 1.5 Aims of this study

The aim of this study was to describe in detail how bacteria and fungi colonise and degrade coal, thus bringing together the fields of biofilm biology and coal biodegradation. The motivation for this research is the immense potential for the application of coal bioconversion to methane.

In an attempt to bridge the knowledge gaps on cell attachment and colonisation on coal, this study investigated the role of coal as a substratum for cell attachment and substrate for degradation. More focus was given on the former aspect. The general biofilm formation on coal was investigated using Scanning Electron Microscopy (SEM). Focus was particularly given on the initial cell attachment, which determines subsequent microbial colonisation on coal. Thus, physico-chemical analyses on the hydrophobicity of cell and coal surfaces were conducted, and surface thermodynamics of the two surfaces were investigated. It is hypothesised that, similar to adhesion studies of other hydrocarbons, the physico-chemical interactions play a significant role in determining the overall cell adhesion on coal.

Although the focus of this study was on cell attachment to and degradation of untreated bituminous coal, other coal types were included for comparison. Nitric-acid treated bituminous coals, which are more oxidised than untreated coal, were used. Nitric-acid treated coal has been used in previous studies as an amenable substrate for degradation (Laborda et al. 1999; Machnikowska et al. 2002; Maka et al. 1989; Strandberg and Lewis 1987a; Wondrack et al. 1989). This study aimed to investigate how this related to cell attachment and colonisation. A low-rank coal, i.e. lignite, the most commonly used in coal biodegradation studies, was also included for comparison in both attachment and degradation analyses. It is postulated that cell attachment and colonisation is a prerequisite for coal degradation to occur, and a positive correlation between the two is evident across all coal types.

Coal is degraded by both bacteria and fungi, and there are recognisable differences in their mechanism of cell attachment to surface. Similarly, these
36 differences are expected in regard to their adhesion and colonisation on coal. Chapter 2 is dedicated solely to bacterial cell attachment and biofilm formation on coal, using Pseudomonas fluorescens Pf-5 as the model bacterium. P. fluorescens Pf-5 was obtained from a culture collection (Centre for Marine Bioinnovation, Sydney, NSW) and its genome has been completely sequenced. Although it is known that P. fluorescens is able to degrade coal, the Pf-5 strain has not been previously tested for coal degradation. A section in this chapter aimed to assess its coal-degrading ability using Fourier Transform Infrared Spectroscopy (FTIR).

Another objective of this study was to obtain a fungal isolate native to Australia capable of degrading coal for potential field applications. Hence, Chapter 3 is dedicated to the isolation of a coal-degrading fungus. The isolated fungus was used as the model organism to investigate fungal cell attachment to and colonisation of coal, as described in Chapter 4. Characterisation of this fungus in regard to its taxonomic identity and coal-degrading ability was also included in this chapter.

Chapter 5 explores the application of coal in a natural environment, where microorganisms exist as communities instead of single species. Through the use of terminal restriction fragment length polymorphism (T-RFLP) and 454 pyrosequencing technology, unique microbial communities adhering to coal were analysed and identified, giving a new perspective on the types of microorganisms, largely uncultivable, that would attach to and colonise coal in a natural environment.

37 2 Pseudomonas fluorescens attachment and oxidation of bituminous coal

2.1 Introduction

As studies pertaining to the attachment and colonisation of coal by microbes are currently under-established, a basic level of investigation is necessary before conducting more complex work. In this study, a known coal degrading bacterium, Pseudomonas fluorescens was chosen as model organism for attachment and colonisation of bituminous coal. P. fluorescens is a Gram- negative rod-shaped bacterium that is commonly found in the environment, particularly in soil and water. Fakoussa (1988) showed that a P. fluorescens strain, isolated from forest fire regions, was able to modify the physical and chemical structure of bituminous coal. In his study, P. fluorescens was shown to secrete surfactants, lowering the surface tension and resulting in the change of color and solubility of the coal particles. Based on infrared spectroscopy and esterification experiments the supernatant revealed a high content of carboxylic and hydroxyl groups, signifying coal oxidation by the strain. Furthermore, a P. fluorescens strain cLP6a was shown to degrade a number of polyaromatic hydrocarbons (PAHs) also found in bituminous coal (Hearn et al. 2003; Foght and Westlake 1996).

P. fluorescens has been studied in diverse areas of cell attachment research, i.e. fundamental work on biofilm formation, novel gene discoveries, effects of antibiotics, and role of lipopolysaccharides on biofilm formation (Williams and Fletcher 1996; Korber et al. 1994; Hinsa and O'Toole 2006). Studies have described P. fluorescens attachment to nano-microstructured surfaces of metals, liquid hydrocarbons and soil, including cell and substratum topography, morphology and surfactant production (Dorobantu et al. 2008; Diaz et al. 2010; Baum et al. 2009; Abbasnezhad et al. 2008). Thus, based on the literature coverage of P. fluorescens in both coal degradation and cell attachment

38 studies, this is a suitable model organism for studying bacterial attachment to coal.

The first aim of this chapter was to demonstrate coal oxidation by the P. fluorescens strain under investigation. This would ensure that the strain used was capable of degrading coal (as the sole carbon source or not), and further investigation on its attachment behaviour would be warranted. Secondly, direct observation using Scanning Electron Microscopy (SEM) was used to describe the bacterium’s attachment behaviour on coal as substratum under different conditions. As coal is an attractive hydrocarbon source for coal-degrading bacteria such as P. fluorescens, it is hypothesised that the cell attachment behaviour would change under carbon-limiting conditions. Thirdly, contact angle measurements were conducted to analyse the hydrophobicity of P. fluorescens and coal of different oxidation ranks to predict thermodynamics of adhesion between cells and coal surfaces. This was done to analyse the influence of physico-chemical factors governing the first form of cell interaction with the coal surface. These findings provide the foundation for further work on this subject, e.g. on other coal degrading bacteria and the biological factors that may contribute to cell adhesion to coal.

2.2 Material and Methods

2.2.1 Medium preparation
A minimal salts medium (M9) pH 7.5 was used in all experiments with P. fluorescens and bituminous coal. The medium contained the following (per L of dH20): 64 g Na2HPO4-7H2O, 15 g KH2PO4, 2.5 g NaCl, 5.0 g NH4Cl, 2 ml of 1 M

MgSO4, 100 μl of 1 M CaCl2 and 20 ml of 20% (w/v) D-glucose solution. The

MgSO4, CaCl2 and glucose solutions were sterilised separately prior to mixing with the other salts solution. In addition to the M9 medium, 0.25% (w/v) of casamino acids (CAA) were added to a number of experiments.

39 2.2.2 Culture preparation
A 3-day old culture of P. fluorescens Pf-5 (Paulsen et al. 2005) was grown in minimal M9 medium at 30 oC shaking at 90 rpm. This served as the inoculum for experiments involving P. fluorescens attachment to and oxidation of coal. Unless otherwise stated, the inoculum was washed by centrifugation (4000 x g for 10 min each wash) with M9 medium without glucose and casamino acids (CAA) at least three times before it was added into fresh medium to remove excess carbon and nitrogen sources. The washed cells were inoculated into a 6 or 12-well plate (Iwaki Microplate, Japan) that contained ethanol-sterilised coal pieces (refer Section 2.2.3 for coal preparation) in sterile M9 medium with and without glucose and/or CAA. The final cell concentration was approximately 5 x 106 cells/mL. The cultures were incubated for 24 h at 30 oC shaking at 20 rpm.

To prepare killed cells, the above culture was pelleted by centrifugation and re- suspended in 50 mg/L mercuric chloride (HgCl2), incubated at room temperature for 24 h and agitated at 20 rpm. After treatment, the cells were washed with sterile M9 media by centrifugation. Cells were checked for viability and integrity using LIVE/DEAD cell stain (L7012, Life Technologies Australia Pty. Ltd., Australia) and viewed using an Olympus BX51 fluorescence microscope. Approximately 90% of the cells were dead when compared against the live cells. Cells were also shown to be intact.

2.2.3 Coal preparation
In this study, two main sources of coal were used: 1) High-ranked bituminous coal and 2) Low-ranked lignite coal. In addition, to create a variety of coal surfaces for the experiments and to investigate the effects of various treatments on coal surface properties, the bituminous coal underwent several chemical treatments involving peroxidase, hydrogen peroxide and nitric acid that were mainly used for Attenuated Total Reflectance- Fourier Transform Infra Red (ATR-FTIR) spectroscopy and contact angle analyses. Coal incubated with P. fluorescens was also included in this analysis. Note that for all coal samples analysed for FTIR, contact angle and SEM imaging, a preparation procedure of
40 fixation, dehydration and drying (section 2.2.6) was conducted.

2.2.3.1 Bituminous coal
Bituminous coal was obtained from within solid, bacteria-free cores taken on August 2010 from a coal seam 80 m below ground level at Lithgow (New South Wales, Australia). For the cell adhesion experiment involving SEM imaging only, the coal was crushed into small pieces of approximately 2 x 1 x 1 mm in size for each piece. For experiments involving ATR-FTIR and contact angle analyses, coal pieces of approximately 30 x 20 x 15 mm were cut and polished to attain a flat and smooth surface. The polishing procedures used the Kent 3 Automatic Lapping and Polishing Unit (Kermet International Ltd, Maidstone, UK) on a 1 μm grit polishing paper (Leco Australia Pty Ltd, New South Wales, Australia). All coal pieces were washed and sonicated for 1 min to remove excess debris. The coal pieces were then sterilised by immersion in 70% ethanol for 30 min, washed with sterile MilliQ water (EMD Millipore Corp., USA) and dried at 22 oC in a biosafety cabinet for at least 1 h before being used as substrata.

2.2.3.2 Lignite coal

Naturally weathered lignite coal was obtained from an open mine in Loy Yang, La Trobe, Victoria, Australia. The coal had been stored loosely in sealed bags in the dark at room temperature for a number of years prior to its transfer and use at the University of New South Wales. The coal was used for ATR-FTIR and contact angle analyses and underwent the same polishing and sterilisation procedures to that of bituminous coal.

2.2.3.3 Peroxidase treated bituminous coal
Polished bituminous coal pieces were treated with versatile peroxidase in 100 mM tartaric acid buffer pH 3.5 with 1% v/v hydrogen peroxide with the following concentrations (mg/L): 0, 0.0024, 0.024 and 0.24. The coal samples were

41 incubated at 37°C for 12 h, rinsed three times with sterilised deionised water, air dried for 48 h and analysed for ATR-FTIR spectroscopy and contact angle.

2.2.3.4 Hydrogen peroxide treated bituminous coal
Polished bituminous coal pieces were treated with hydrogen peroxide solution (Sigma- Aldrich, USA) at different concentrations (% v/v): 0, 1, 10 and 30. The treated coal samples were then washed with sterilised deionised water, air dried for 48 h and analysed for ATR-FTIR spectroscopy and contact angle.

2.2.3.5 Nitric acid treated bituminous coal
Polished bituminous coal pieces were immersed in 4.69 N of nitric acid for 48 h, washed thoroughly with deionised water (pH> 5), air-dried and analysed for ATR-FTIR spectroscopy and contact angle.

2.2.3.6 P. fluorescens treated bituminous coal

P. fluorescens treatment of coal involved following the same procedure to culture preparation and incubation (section 2.2.2) in M9 medium (without glucose and CAA) and including polished coal pieces of approximately 30 x 20 x 15 mm each in a sterile 12-well plate. The incubation was conducted at 30 oC for 24 h and shaking at 20 rpm. At the end of incubation, the polished coal pieces were gently rinsed in M9 medium and underwent cell fixation of 2.5% glutaraldeyhde in potassium phosphate buffer pH 7.2 for 2.5 h and drying before being measured for contact angle.

2.2.3.7 P. fluorescens treated bituminous coal – scraped

In order to measure the contact angle of P. fluorescens treated coal without P. fluorescens attachment, following the procedure in section 2.2.3.6, the coal surface was scraped using sterile cell scrapers (Blade 25, Sarstedt AG & Co., Germany) several times using both blade sides (once from each side). The scraped coal was then measured for contact angle. SEM (refer to section 2.2.6
42 for standard preparation) of the scraped coal was performed to ensure that the scraped coal surface was free of cells. Based on the micrographs approximately 90% of the cells were successfully removed after scraping (Appendix III).

2.2.4 Control surfaces preparation

For the P. fluorescens adhesion to coal experiments involving SEM imaging, control surfaces, i.e. glass (inert surface) and pebbles (similar topography to coal surface) were used. A microscope glass slide was crushed into small pieces of approximately 15 x 1 x 2 mm using pestle and mortar to create a rougher surface.

Small pebbles of approximately 2 x 1 x 1 mm were collected from the seashore of Coogee Beach, New South Wales, Australia. To remove potential carbon sources (e.g. calcium carbonate), the pebbles were acid-washed using 50% (v/v) nitric acid for 1 h at room temperature and agitated at 50 rpm. The pebbles were then washed thoroughly with deionised water until the pH of the water stabilised close to neutral.

The fragmented glass pieces and pebbles were sterilised by immersing in 70% (v/v) ethanol solution for at least 30 min, rinsed in sterile MilliQ water and dried at room temperature before further use.

For ATR-FTIR and contact angle measurements, an unbroken microscope glass slide was used as a flat surface. The glass slide underwent similar sterilisation, rinsing and drying procedures as above prior to use.

2.2.5 Attenuated Total Reflectance- Fourier Transform Infra Red (ATR- FTIR) Spectroscopy of coal

Infrared spectroscopy of coal surfaces was conducted on a Perkin Elmer Spotlight 400 FTIR microspectrometer fitted with a liquid nitrogen-cooled 16 element linear array mercury cadmium telluride (MCT) detector (PerkinElmer,

43 Beaconsfield, UK). A 400 x 400 μm region was analysed using a germanium Attenuated Total Reflectance (ATR) imaging crystal and processed using the PerkinElmer Spectrum IMAGE software. The data was corrected for the effect of atmosphere, then an image generated using the baseline corrected amide I C=O peak of P. fluorescens. For the cell attachment experiment, controls of sterile coal and glass were also measured.

Stackplots of the average of 64 spectra were prepared using the PerkinElmer Spectrum 6 software. Peak fitting of individual spectra was performed using the Thermo GRAMS software.

2.2.6 Scanning Electron Microscopy of P. fluorescens attachment on bituminous coal
Using pre-sterilised forceps, coal pieces that had been incubated with P. fluorescens were taken out of the culture and gently washed with pre-warmed minimal medium to remove unattached cells and minimise the potential dislodging of attached cells from using cold medium. Fixation, dehydration and drying methods were followed from the works of Fratesi et al. (2004) and Hazrin-Chong and Manefield (2012).

Cells adhering to washed coal pieces were chemically fixed using 2.5% (v/w) glutaraldehyde (Sigma-Aldrich, USA) for 2.5 h in 0.1 M potassium phosphate buffer, pH 7.2. The coal particles were gently rinsed with M9 minimal medium three times and underwent a graded dehydration series of ethanol (35, 50, 70, 80, 90, 95, 100, 100, 100% for 10 min each) and hexamethyldisilizane (HMDS) (Fluka, Castle Hill, Australia) drying (50 and 100% for 10 min each).
The treated coal pieces were left to dry for 2 days in a fume hood at 22 °C.
After drying, the samples were mounted on aluminum stubs, sputter coated with a layer of gold using an Emitech K550 sputter coater (Quorum Technologies, UK) and viewed using an ESEM Quanta 500 microscope (FEI, Oregon, USA) at 10 kV.

For experiments involving all three SEM, FTIR and contact angle

44 measurements, the coal samples firstly underwent the SEM preparation up until the HMDS drying, and were then analysed for FTIR and CA prior to gold sputtering for SEM. This is to avoid interference of gold coating to the FTIR and CA analyses.

2.2.7 Imaging of P. fluorescens cells in supernatant in the presence of coal and without glucose and CAA
Cells in the supernatant that were incubated with coal and without glucose and CAA were visualised through fluorescence microscopy to compare the free- living cell morphology against those that were attached on the coal surface. After 24 h of incubation, the cells in the supernatant were washed three times with M9 medium without glucose and CAA by centrifugation (2000 x G, 10 min each). The cells were then stained with SYTO 9® Green Fluorescent Nucleic Acid Stain (Life Technologies Australia Pty. Ltd., Australia) before imaging using Olympus BX51 Fixed Stage Fluorescence Microscope. At least 20 images were captured at random locations within the sample.

2.2.8 Contact angle measurements and surface thermodynamics of adhesion between P. fluorescens and coal
Contact angle measurements were made on several different types of coal and P. fluorescens surfaces to measure hydrophobicity, surface energies and subsequently free energy of adhesion between two interacting surfaces. The contact angle of glass as a control surface was also measured. The contact angles were measured using a contact angle goniometer (KSV Cam 200, KSV Instruments Ltd., Finland) at room temperature (approximately 22 oC).

The bacterial surface was prepared by depositing cells on a 0.45 μm pore filter paper (Merck Millipore, USA) using negative pressure to create a smooth and flat lawn of compactly organised cells. Sterile polished coal surfaces and glass were used directly (refer to sections 2.2.3 and 2.2.4 for polishing and sterilisation procedures). The static sessile drop method was used to measure

45 the contact angles using three liquid types: deionised water, formamide and diiodomethane. These three test liquids were used as probes for surface free energy calculations and surface free energy of adhesion. The test liquids and their surface tension components, which comprised the apolar or Lifshitz-van

 der Waals component

, the polar or acid-base component

, electron

acceptor

and the electron donor

sub-components are summarised in Table 2.1 The surface energies of P. fluorescens and coal were calculated using the van Oss acid-base approach (see section 1.3.5.1) (Van Oss et al. 1988).

Table 2.1 Test liquids and their surface energy components.




Surface tension









(mN m-1)

Water, H2O 72.8 21.8 51.0 25.5 25.5 Formamide, 58.0 39.0 19.0 39.6 2.3

(CH3NO) Diiodomethane, 50.8 50.8 0.0 0.0 0.0

(CH2I2)

The Young equation describes the theory of the contact angle of pure liquids on a solid (Young 1805):










(2)

where

is the experimentally derived surface tension of the liquid,
is the contact angle,

is the solid surface energy and

is the solid/liquid interfacial energy. In order to obtain

, an estimate of

needs to be acquired.

An acid-base theory for calculating surface energy was developed by van Oss (1988) which comprises the sum of a Lifshitz-van der Waals apolar component



and a Lewis acid-base polar component

:













(3)

46

 by which

can be further divided to an electron donor and an electron acceptor subcomponent:













(4)

Thus, the solid/liquid interfacial energy can be obtained by:









































(5)

A relation between the measured contact angle and the solid and liquid surface free energy can be obtained by combining equation (5) with the Young equation (2):








































(6)

The contact angle of at least three liquids with known surface tension components (like given in Table 2.1) on the tested surface must be known in


order to obtain the surface energy components (

,

,

).

It is possible to predict whether or not adhesion between two surfaces, in this case between P. fluorescens and coal, is thermodynamically favourable. This can be achieved by using the interfacial surface energies of all the interfaces involved. When a bacterium attaches to a solid in a liquid environment, three types of interfaces are involved: bacterium-solid (substratum), bacterium-liquid, and solid-liquid, each with its own interfacial energy, i.e.

 ,

 and

respectively. Therefore, the strength of bacterial adhesion to a particular surface, i.e. the surface free energy of adhesion,



, can be given by:














(1)

All three interfacial energies

,

 and

can be calculated from measured
47 contact angle data and the van Oss acid-base approach. Thermodynamically (assuming zero electrostatic interactions), adhesion of bacteria is favored if


is negative whereas a positive



implies an unfavorable adhesion.

2.2.9 Enumeration of P. fluorescens cells on different types of coal
In parallel to the contact angle measurement and surface thermodynamics of adhesion calculation, P. fluorescens cells attaching to different types of coal (i.e. bituminous, nitric acid treated, peroxidase treated, peroxide treated and lignite) were enumerated using SEM. The same coal pieces used for contact angle measurements were incubated with 3-day grown P. fluorescens of the same concentration (9.4 x 106 cells/ml) across all coal types in M9 medium without glucose and CAA for 24 h. The coal pieces were then rinsed gently three times in the same medium before being processed for SEM (section 2.2.6). On average 50 images were taken for each sample and cell numbers were calculated to achieve number of cells/mm2.
2.2.10 Live and dead P. fluorescens attachment on coal

Live and HgCl2-killed P. fluorescens were compared to observe if there was any biological influence on cell attachment to coal. Both live and dead cultures (refer Section 2.2.1 for killed culture preparation) were incubated with polished bituminous coal pieces in M9 media without glucose and CAA. The incubation was conducted for 24 h at 30 oC with agitation at 20 rpm. The coal pieces then underwent an SEM preparation (see Section 2.2.6) and were viewed using SEM for differences in cell counts.

2.2.11 Statistical analyses

The student t-test (one-tailed distribution, paired) was employed using Microsoft Excel 2011 to determine statistical significance in the difference between two samples of interest.

48 2.3 Results
2.3.1 ATR-FTIR of coal oxidation by P. fluorescens
To demonstrate that the P. fluorescens Pf-5 strain used in this study was capable of degrading coal, FTIR of the coal surface exposed to bacteria was conducted. Polished coal pieces were incubated with P. fluorescens for 24 h, processed for SEM (up until the drying step), left dried and sampled for ATR- FTIR. Figure 2.1A shows the ATR-FTIR spectrum of P. fluorescens on coal along with two control samples: P. fluorescens on a glass slide and a bacteria- free coal. The key chemical bond vibrations are labeled in the figure, of which the most notable are the amide I C=O stretch, the amide II NH deformation mode, and the C=C stretching peak arising from aromatic ring deformation modes. The P. fluorescens and coal spectrum also shows a peak at 1463 cm-1 that can be assigned to the CH2 deformation mode.

The bacteria-free coal spectrum shows a peak for the C-H stretch around 2900 -1 cm that is observed in the biological samples. A CH2 deformation mode is also observed on this sample, however the peak is at a lower frequency of 1438 cm- 1. Two peaks in the region of interest for this study were the C=C stretch at 1595 cm-1 and a C=O stretch present as a high frequency shoulder at around 1715 cm-1. This C=C stretch can also be seen in the bacteria and coal spectrum. Furthermore, the region between 1784 and 1560 cm-1 in the spectrum shows an excellent fit to four Gaussian peaks (Figure 2.1B). These are assigned as follows: coal C=C (1592 cm-1); amide I C=O in protein (1659 cm-1); coal C=O (1715 cm-1); and a biogenic C=O (1736 cm-1).
The fourth peak (C=O stretch at 1736 cm-1) is not observed in P. fluorescens on glass or the bacteria-free coal spectra. Table 2.2 summarises the parameters obtained from the peak fitting analysis, along with proposed assignments of each peak. A four- Gaussian peak fit produced calculated spectra in excellent agreement with the experimental data.

49
Figure 2.1 (A) Representative FTIR stackplot of: (i) P. fluorescens on a glass slide; (ii) P. fluorescens on coal; and (iii) coal sample with no biological material. (B) Expanded region of spectrum (ii) showing experimental points (circular markers), four Gaussian peaks (broken lines), the linear baseline and calculated spectrum (solid lines).

50 Table 2.2 Peak fitting statistics of a region rich in P. fluorescens on coal. The spectrum in the region 1784-1560 cm-1 was fit using a linear baseline (slope -0.0035, offset 107.6) and four Gaussian functions to give a correlation of 0.9988817 to the experimental data. ‘Biogenic C=O stretch’ (first row) refers to C=O stretch that was only found in P. fluorescens and coal spectrum.

Peak Assignment Peak Peak Height Peak Width Peak Area

Position (%T) (cm-1) (%)

(cm-1)

Biogenic C=O 1736 2.53 30.9 14.8 stretch

Coal C=O stretch 1715 0.59 16.1 1.8

Amide I C=O 1659 5.71 71.9 77.6 stretch

Coal C=C stretch 1592 0.93 33.4 5.9

A positive control for coal oxidation by FTIR using peroxidase as a biochemical oxidising agent was conducted. The peroxidase-treated polished coal samples were prepared in a similar fashion to the bituminous coal sample treated with P. fluorescens and sampled for ATR-FTIR.

Table 2.3 shows peak fitting results for different concentrations of peroxidase- treated coal approximately at the 1500-1700 cm-1 of ATR-FTIR spectral range (for peak fitting curves, see Appendix IV). The C=O stretch (peak 1) found in the range 1732-1714 cm-1 is observed at high frequency only when peroxidase is present (0.0024-0.24 mg/mL). This peak corresponds approximately to the 1736 cm-1 C=O peak found in P. fluorescens-treated coal sample (Figure 2.1A). Furthermore the ratio of peak 1 to native coal C=O peak (peak 2) stabilises at 0.024 mg/mL peroxidase and the highest concentration of peroxidase (0.24 mg/mL) does not increase the ratio of biogenic C=O to native C=O on the coal.

51 Peak 3 (coal C=C) did not change with increasing peroxidase concentration, which indicates its relative stability in the presence of peroxidase.

Table 2.3 Peak fitting results for influence of peroxidase on coal. Three main peaks are assigned for each spectrum of coal treated with different peroxidase concentrations (0, 0.0024, 0.024 and 0.24 mg/mL): Peak 1, C=O resulting from peroxidase treatment, Peak 2, native coal C=O and Peak 3, C=C from coal. Peak 1 is observed at 1727-1732 cm-1 only on peroxidase- treated coal spectra, where it increases in intensity (shown by peak area, %) but stabilises at 0.024 mg/mL. Peaks 2 (observed at 1695-1705 cm-1) and 3 (1593-1600 cm-1) do not show increase in peak area (%) at increasing peroxidase concentrations.

Peak Peroxidase concentration (mg/mL) assignment 0 0.0024 0.024 0.24

Peak 1 Not observed 1732 1714 1727 -1 C=O (cm ) 0% 0.9% 4.7% 4.4% (area) Peak 2 1703 1706 1695 1705 Native coal 15% 4.6% 1.4% 1.4% C=O (cm-1) (area) Peak 3 1600 1594 1593 1596 Coal C=C 85% 94.5% 93.9% 94.2% (cm-1) (area)

The ATR-FTIR analyses showed that P. fluorescens Pf-5 degrades bituminous coal via oxidation, possibly through peroxidase enzyme mechanism. This strain was thus used in subsequent experiments assessing its mechanisms of adhesion and colonisation.

2.3.2 Scanning Electron Microscopy (SEM) of P. fluorescens attachment on bituminous coal in different media
To investigate P. fluorescens attachment behavior on coal under different media conditions, SEM was performed on coal pieces incubated with P.

52 fluorescens in minimal salts (M9) medium with a) 0.4% (v/v) glucose and 0.25% (v/v) casamino acids (CAA) (Figure 2.2), b) glucose only (Figure 2.3) and c) no external carbon source other than coal (Figures 2.3-2.5) at 30 oC for 24 h. At least 50 images for each sample were taken.

Figure 2.2 shows representative micrographs of P. fluorescens attachment on coal in the presence of glucose and CAA at different magnifications. In this medium the cells formed extensive biofilms with complex architecture consisting of small channels, voids and inter-linking material between the cells (Figure 2.2C). Furthermore there were areas on the coal surface where individual cells or small clusters of three to five cells were present (Figure 2.2D).

Figure 2.2 Representative micrographs of P. fluorescens attachment on coal in M9 medium with glucose and CAA after 24 h of incubation. Cells were observed to form extensive biofilms (A-B) with characteristic biofilm entities (C) such as water channels (block arrow) and material interlinking between the cells (narrowed arrow). Individual (full arrow) or clusters of three to five cells (dashed arrow) were also observed in some areas (D).
53
Incubations were made where glucose was omitted from the growth medium while CAA was kept. The visual observations by SEM were similar to those with both glucose and CAA in Figure 2.2 (images not shown). Therefore, to limit external biofilm promoting agents, CAA was omitted in subsequent experiments.

Where glucose was present but CAA absent, P. fluorescens cells again attached to the coal surface (Figure 2.3). However, there were no extensive biofilms formed like those observed in glucose and CAA medium. Cells in glucose-only medium were mostly individually attached (Figure 2.3A), and in cases where cell clusters were observed the cells were often seen trapped in between dents and crevices native to the coal surface (Figure 2.3 B).

A B

Figure 2.3 Representative micrographs of P. fluorescens attachment on coal in glucose-only M9 medium after 24 h of incubation. No extensive biofilm was observed, however cells were attaching individually (A) and forming clusters within crevices of the coal surface (B). Elongated cells could also be found across the coal surface (arrow).
The P. fluorescens attachment to coal study was further conducted without glucose and/or CAA but only coal as the sole carbon source. Figures 2.4 to 2.6 represent P. fluorescens SEM observations under these conditions. The cells exhibited two forms of cell attachment, i.e. individual cell attachment (Figure 2.4A) and microcolony formation (Figure 2.4B). Interestingly, the latter was not observed in medium with external carbon source/s present.

54

Figure 2.4 Representative micrographs of P. fluorescens attachment on coal with coal as the sole carbon source after 24 h of incubation. The cells exhibited A) individual cell attachment and B) microcolony formation (arrows).

The microcolonies were further magnified to illustrate its structure in more detail. These microcolonies each of approximately 40 x 50 x 30 μm in size consisted of aggregated microbial ‘balls’ in which small channels or voids characteristic of biofilm architecture could be observed (Figures 2.5-2.6). Small grainy material could also be observed across all microcolonies (Figure 2.5) that were not present among individual cells or the biofilm formed in glucose and CAA M9 medium.

The average length of a short cell was approximately 1.7 μm, however it was difficult to measure long cell length due to its intertwining nature. Nevertheless, the width for both short and long cells was approximately 0.6 μm. The microcolony mix of short and long cells could also be observed colonising coal microparticles (Figure 2.6). Other representative micrographs of microcolonies can be found in Appendix III.

55
Figure 2.5 Microcolony of P. fluorescens in M9 medium without external carbon source present. Each microcolony consisted of short and extended cells and small airways (full arrow) characteristic of biofilm formation. Small grainy material in between the cells (dashed arrow) could also be observed in each microcolony that was not present among individual cells or in biofilms formed when glucose and CAA was present.

Figure 2.6 A microcolony of P. fluorescens consisting of short and extended cells (square) colonising a coal microparticle in the absence of an additional carbon source.

56 Prior to SEM observations, optical density (OD600) measurements of the supernatant in the coal attachment by P. fluorescens study were taken as an indication of biomass growth within 24 h in media with and without glucose. Figure 2.7 shows that cell density in medium with glucose almost doubled in 24 h whereas there was no significant change in cell density in the supernatant without glucose.

10 Without glucose With glucose 600 1 OD

0.1 0 h 24 h

Figure 2.7 Optical density (OD600) of supernatants of P. fluorescens used in the coal adhesion experiment in the presence and absence of glucose. Significant increase in cell density was observed in medium with glucose, whereas there was no change observed in the absence of glucose. Error bars indicate standard errors of three pseudo-replicates.

Based on the SEM micrograph, in all tested conditions P. fluorescens exhibited rod-shaped morphology typical of Pseudomonas species. However, the cell length and width varied from one medium type to another. Table 2.4 summarises the average length and width of P. fluorescens cells in all three medium types.

57 Table 2.4 Cell length and width (μm) of P. fluorescens on coal in different M9 medium.

M9 Medium Cell length (μm) Cell width (μm) Glucose + CAA 1.4 ± 0.2 0.7 ± 0.1 Glucose 2.1 – 8.4 0.5 ± 0.1 No additional carbon > 1.7 0.6 ± 0.1

The supernatant of the culture incubated with coal in M9 medium with no additional carbon was analysed using fluorescence microscopy to assess whether the cells were elongated similar to those observed on the coal surface. Figure 2.8 shows a representative micrograph of stained P. fluorescens cells in the supernatant. No elongation of cells was observed; the average cell length is approximately 2.3 ± 0.4 μm, similar to that of the shorter cells observed attaching on the coal surface.

Figure 2.8 Representative image of free-living P. fluorescens in M9 medium without glucose and CAA. No cell elongation was observed compared to cells that were attached on the coal surface.

To test whether cell attachment, elongation and aggregation on coal without the presence of additional carbon sources were exclusive to coal surfaces, two

58 controls using a glass and a rock surface were conducted. Figure 2.9 illustrates P. fluorescens attachment to glass at 24 h in M9 medium without any carbon source. Cell attachment still occurred; however, there was neither signs of cell elongation nor microcolonies formed on glass or rock as observed on the coal surface in the absence of glucose and CAA.

Figure 2.9 Representative micrographs of P. fluorescens on broken glass (A) and acid-washed pebble (B) in M9 medium without external carbon source after 24 h of incubation. Cells were attached to both surfaces, however there was no elongation or aggregation of cells.

2.3.3 Contact angle (CA) measurements

In order to investigate the effect of physico-chemical factors on P. fluorescens adhesion to coal surfaces in different conditions, hydrophobicity measurements determined by contact angle were conducted. The hydrophobicity of each surface was compared using water contact angles; the Gibbs surface free energy of adhesion was calculated using water, formamide and diiodomethane contact angles of each surface.

2.3.3.1 Water contact angle measurements of P. fluorescens
Figure 2.10 shows the water contact angle measurements for P. fluorescens surface in which cells were subjected to three different pre-treatments: glucose- fed, glucose-starved and mercuric chloride incubated in M9 salts medium. All cell surface types showed contact angles between 10-14o, which were considered low compared to other surfaces studied previously (Abbasnezhad et

59 al., 2008; Boks et al., 2008; Das et al., 2010; Sharma & Rao, 2002). A low contact angle of a cell surface would imply that the cell is hydrophilic. Whilst there was hardly any difference between the contact angles of glucose-starved and killed P. fluorescens, the contact angle of glucose-fed cells was significantly lower than the former two surfaces (P<0.05).

20

18

) 16 W

θ 14

12

10

8

6

Water contact angle ( Water 4

2

0 Glucose-fed Glucose-starved Killed

Figure 2.10 Average water contact angles of three types of P. fluorescens: Glucose-fed, starved and killed. Error bars represent standard error of three independent replicates, each with at least two measurements at different sites.

2.3.3.2 Contact angle measurements and surface energy components of various surfaces
The water contact angle of P. fluorescens was then further compared against different types of coal and glass as controls. Overall, all types of coal (except for

HNO3-treated coal) have significantly (P<0.01) higher water contact angles than that of P. fluorescens cells (Figure 2.11). The highest contact angle for coal was the bituminous type, which on average was approximately 118o, whilst the contact angle of lignite coal was approximately 85o, significantly lower than that of bituminous coal, nevertheless still relatively high among other coal surfaces.

Chemical treatments of bituminous coal using HNO3, peroxidase and peroxide produced substantially lower contact angles than the original bituminous coal.
60 This suggests the oxidation of coal lowers its hydrophobicity. The lowest contact angle produced on the chemically treated coal surface was the HNO3 treated (approximately 25o), followed by peroxidase (48o) and peroxide (65o).

The contact angle of P. fluorescens treated bituminous coal was found to be of approximately 47o, i.e. more than half the contact angle of untreated bituminous coal. However, when the cells were scraped from the coal surface, the contact angle of the coal increased significantly (approx. 97o), slightly higher than that of lignite coal but lower than the untreated bituminous coal. This shows that the presence of P. fluorescens on bituminous coal lowered the coal surface hydrophobicity markedly, however when cells were removed the coal hydrophobicity returned closely to its original hydrophobic state.

A control from a glass slide for contact angle measurement was also conducted. The contact angle for glass was substantially lower (approx. 4o) compared to those of coal surfaces and P. fluorescens.


140

) 120 W

θ 100 80 60 40 20 0 Water contact angle ( Water

Figure 2.11 Water contact angles of surfaces of P. fluorescens (glucose-fed), various coal types and glass slides. HNO3, peroxidase, peroxide, and P. fluorescens treated coal consist of bituminous coal. Error bars represent standard deviations of at least three independent replicates.

61
Table 2.5 summarises water, formamide and diiodomethane contact angles along with the calculated surface energy components (γ) for each tested surface. The contact angles of these three liquids on the surfaces were used as probes in calculating the surface energies, which were subsequently used to calculate total free energy of adhesion ∆GAdh between two surfaces (Table 2.6).

62 W F Di Table 2.5 A summary of all contact angle work, i.e. water (θ ), formamide (θ ) and diiodomethane (θ ) on various P. fluorescens and coal surfaces LW + - and glass as control. Based on these values, the Lifshitz-van der Waals apolar component (γ ), electron acceptor (γ ) and electron donator (γ ) sub- AB TOT components of Lewis acid-base polar component (γ ) were calculated to give the total surface energy (γ ) for each surface.
-1 Contact angle θ (o) Surface energy components (mN m ) Surface θ W θ F θ Di γ LW γ + γ - γ TOT

P. fl. normal 9.7 ± 2.7 14.6 ± 5.1 47 ± 3.0 42 0.7 55.6 54.6 P. fl. starved 14.3 ± 2.6 18.0 ± 3.6 69.6 ± 1.8 23.2 6.1 52.9 59.1 P. fl. killed 14.6 ± 3.7 17.5 ± 7.2 57.3 ± 5.9 30.1 3.3 53.4 56.5 Bituminous 118.2 ± 3.1 101.7 ± 6.8 57.9 ± 9.6 29.8 0 0.4 29.8 Lignite 85.4 ± 3.6 16.2 ± 3.1 16 ± 3.5 48.9 5.5 0 48.9

HNO3 24.7 ± 4.8 18.6 ± 5.5 20.3 ± 6.8 48.2 0.1 48.6 53.3 treated Peroxidase 48.1 ± 7.2 20.5 ± 7.3 28.2 ± 2.0 47.6 0.4 25.2 53.9 treated Peroxide 64.6 ± 4.7 58.3 ± 11.0 28.7 ± 2.3 29.6 6.9 7 43.4 treated P. fl. treated 46.5 ± 4.5 18.5 ± 2.6 12.8 ± 2.3 49.5 0.7 22.9 57.5 P. fl. treated 96.5 ± 5.7 62.8 ± 5.3 42.9 ± 4.4 38.1 0.3 0 38.1 - scraped Glass 3.8 ± 6.7 3.9 ± 4.6 37.8 ± 5 41 1.2 54.8 57

63 2.3.4 Thermodynamics of adhesion between P. fluorescens and coal
Based on water, formamide and diiodomethane contact angle measurements and using the Lifshitz-Van der Waals/acid-base surface free energies concept, the thermodynamics (i.e. free energy) of adhesion between two surfaces can be calculated. This gives valuable predictions on probable adhesions between two surfaces based on physico-chemical influences. A negative value of the surface free energy would indicate that the likelihood of attachment between two surfaces is high. In contrast, positive surface free energies would indicate attachment is unlikely based on physico-chemical factors.

Table 2.6 summarises the total Gibbs free energy of adhesion ∆GAdh between various types of coal surfaces and P. fluorescens. Certain surfaces, in particular coal surfaces without modification, i.e. bituminous and lignite, resulted in negative ∆GAdh with all P. fluorescens surface types. Peroxide and scraped P. fluorescens treated coal also produced negative ∆GAdh with P. fluorescens. The most negative ∆GAdh among all tests was between glucose-fed P. fluorecens and lignite. This was followed by the same glucose-fed cells but with scraped P. fluorescens treated bituminous coal and the non-treated bituminous coal. Killed and starved P. fluorescens followed a similar order of negative ∆GAdh to the glucose-fed cells with the coal surfaces (lignite, P. fluorescens treated-scraped and bituminous coal) but with a higher ∆GAdh (albeit still negative).

In contrast, surfaces that had been treated with nitric acid, peroxidase and P. fluorescens produced positive free energy of adhesion with any of the bacterial surfaces. Positive ∆GAdh were also produced between all P. fluorescens and control surfaces: glass and their own cell surface (e.g. glucose-fed cells against glucose-fed cells). The highest ∆GAdh values were between all P. fluorescens cell surfaces with nitric acid treated coal, the cells’ own surface and the glass slide (with the exception of glucose-fed P. fluorescens). This was followed by peroxidase and P. fluorescens treated coal with all P. fluorescens.

64 Despite the differences in free energy of adhesion between different P. fluorescens surfaces for a particular coal/control surface, the negativity or positivity remained the same across all bacterial surfaces with a particular coal/control surface. For example, all types of P. fluorescens cell surface had negative ∆GAdh with bituminous coal and positive ∆GAdh with peroxidase treated coal, despite there may be a large difference in the value between each other.

65 Table 2.6 Total Gibbs free energy of adhesion (∆GAdh) between differently treated P. fluorescens cell and coal surfaces with glass as reference. The total Gibbs free energy accounted for the sum of adhesion of Lifshitz-van der Waals and acid-base components. ‘Normal’ refers to glucose-fed cells whereas ‘starved’ is glucose-starved cells.

2 Coal surface Total free energy of adhesion ∆GAdh (mJ/m )

P. fluorescens cell surface

Normal Starved Killed

Lignite -38 -15 -24

Bituminous -15 -1 -7

Nitric acid-treated 31 30 30

Peroxidase- 13 19 16

treated

Peroxide-treated -11 -2 -6

P. fluorescens- 10 17 13

treated

P. fluorescens -26 -7 -15

treated-scraped

Control surface

Glass slide 10 29 30

Own surface 34 23 28

66 Figure 2.12 shows the number of pre-washed P. fluorescens cells attached to different types of coal (except for P. fluorescens treated and scraped coal) after 24 h of incubation in M9 medium without glucose (referred to as ‘starved’ cells). The initial cell density in the supernatant incubated was constant across all coal samples (9.4 x 106 cells/ml).

The highest number of cells attached were observed to be on lignite coal (8.96 x 104 cells/mm2), which corresponded well to the total free energy of adhesion data in Table 2.6, where it gives the lowest ∆GAdh between P. fluorescens and lignite surface. This was followed by peroxide-treated bituminous (5.8 x 104 cells/mm2) and untreated bituminous coal (4.67 x 104 mm2). There was a significant difference between the number of cells on lignite coal and both peroxide-treated and untreated bituminous coals (P=0.0001 and P<0.0001 respectively), however no significant differences were found between cells attached on the latter two coals (P=0.11). The ∆GAdh for these two coals (against starved P. fluorescens) were also highly similar (Table 2.6).

The lowest numbers of cells attached were on nitric acid-treated and peroxidase-treated bituminous coals, which were 7.4 x 103 cells/mm2 and 1.17 x 103 cells/mm2 respectively. The difference of cells attached on these two coal surfaces is significant (P=0.02). In contrast to the other coal surfaces, the ∆GAdh results for nitric acid-treated and peroxidase-treated coal do not reflect the number of cells attached. Nitric acid-treated coal had the greatest ∆GAdh value followed by that of the peroxidase-treated coal, nevertheless, it was peroxidase- treated coal that has the least number of cells attached based on Figure 2.12.

67 cells/mm2

1.20E+05

1.00E+05

8.00E+04

6.00E+04

4.00E+04

2.00E+04

0.00E+00 Lignite Bituminous Nitric acid- Peroxidase- Peroxide-treated treated treated

Figure 2.12 The number of starved P. fluorescens cells (cells/mm2) attached to various types of coal surfaces. Lignite coal surface held the highest number of cells attached, followed by peroxide-treated bituminous, (untreated) bituminous, nitric acid-treated bituminous and peroxide-treated bituminous coal. The general trend in this graph corresponded well with the

∆GAdh values in Table 2.6.

2.3.5 Live and dead P. fluorescens cell attachment to bituminous coal
To observe whether there was any difference in the numbers of cells attached between alive and killed P. fluorescens on bituminous coal, an experiment was conducted in which both alive and killed P. fluorescens cells of the same density were incubated with bituminous coal for 24 h at 30oC, observed and quantified using SEM.

Figure 2.13 represents SEM images of killed and alive P. fluorescens on bituminous coal. Based on the images alone, it was apparent that the number of live cells attached on the coal surface outnumbered those of killed cells significantly. Quantitation of at least 30 images of the cells on the coal surface revealed that the live cells that were attached to coal was approximately 2.5 x 102 cells/mm2, whereas there were approximately 30 dead cells/mm2 attaching to the coal surface.

68
Figure 2.13 Representative SEM micrographs of A) killed and B) live P. fluorescens on bituminous coal. Significant differences (P<0.05) in cell attachment were observed between the cell types. Arrows are pointing to the dead cells attaching to the coal surface.

2.4 Discussion

This chapter investigates the interaction between a coal-degrading P. fluorescens strain and bituminous coal, where coal was used as a substratum for cells to attach to and use as a carbon and energy source. Based on FTIR analyses, the P. fluorescens strain used has the ability to oxidise coal, which was shown by unique oxidation peaks found only when the bacteria and coal were present. Through microscopy observations and surface thermodynamics analyses, the attachment behaviour of P. fluorescens was different across a number of conditions and coal types. This implies that not only was P. fluorescens altering the coal surface, but that different conditions and coal types influenced the cells’ behaviour on coal.

ATR-FTIR spectra of P. fluorescens on coal and glass and uninoculated coal showed peaks that correspond uniquely to P. fluorescens and coal. The presence of the amide I (C=O) and amide II (N-H) peaks is definitive evidence of cellular protein and, as expected, is only observed in P. fluorescens on glass and coal (Figure 2.1). A peak at 1464 cm-1 from the coal and P. fluorescens spectrum can be assigned to the CH2 deformation mode expected from aliphatic chains in cell membrane lipid molecules with a generic structure such as that shown in Figure 2.14A (below). Both P. fluorecens on coal and glass
69 spectra clearly show that the ATR-FTIR measurement could detect P. fluorescens on coal. This is the first time that ATR-FTIR was used to detect chemical alterations in coal.

The bacteria-free coal spectrum showed a peak for the C-H stretch around -1 2900 cm that is also observed in the P. fluorescens with coal sample. A CH2 deformation mode was also observed on the coal only sample, however the peak was at a lower frequency of 1438 cm-1, as expected from cyclic chemical residues such as in Figure 2.14B in bituminous coal (Friedel and Queiser 1956). The most notable peak in the coal is the C=C stretch at 1595 cm-1 from the aromatic residues in the structure shown in Figure 2.14B that is not expected in large quantities in biological material. This C=C stretch can be seen in the P. fluorescens on coal spectrum and is clear evidence that ATR-FTIR is detecting the underlying coal substrate as well as the cells.

As a naturally derived sample, coal comprises a mixture of organic compounds and polymers, along with a variety of minerals such as quartz and silica, all of which are active in the “fingerprint” region of the infrared spectrum (below ~1400 cm-1). As a result of natural variation in the mineral content in coals it was difficult to link changes in this part of the infrared spectrum to the presence of a biological agent. However the region between 1560 and 1784 cm-1 in the coal with P. fluorescens spectrum is particularly useful in distinguishing the influence of the bacteria on the coal surface. Gaussian peak fitting (Figure 2.1B), shows a peak from C=C in coal (1595 cm-1), protein amide I C=O (1659 cm-1), a peak from non-biogenic C=O groups in coal (1715 cm-1), and a C=O stretch at 1736 cm-1 that is not observed in P. fluorescens on glass or the bacteria-free coal. This C=O functional group must form as a result of bacterial oxidative processes on the coal surface. Therefore, it can be described as a biogenic C=O peak that is a key indicator for the chemical changes occurring at the coal-bacteria interface.

Figure 2.14 below indicates the oxidation positions in coal hydrocarbon groups that are consistent with several notable FTIR peaks. These are aliphatic chains corresponding to cell membrane lipids at 1463 cm-1, cyclic chemical residues in
70 coal at 1438 cm-1 and a C=O resulting from biogenic oxidation of coal at 1736 cm-1. Chemical groups that could give rise to the C=O peak include aldehydes and ketones as well as esters and carboxylic acids (labelled (a), (b) and (c) respectively in Figure 2.14C)
(Socrates 2001).

It is likely that other functional groups resulting from oxidation of the coal surface were present but not observed. A study on oxidative weathering of coals (Liotta et al. 1983) noted extensive formation of aliphatic C-O-C ether groups. Unfortunately the FTIR in this region is not informative for the coal samples in this study since silicate mineral matter was evident in strong Si-O stretching peaks that overlap peaks expected from C-O stretching in ethers.

Figure 2.14 Proposed chemical groups based of FTIR spectra in this study: A) Aliphatic chains found in cell membrane lipid that represents peak 1463 cm-1 on P. fluorescens and coal spectrum, B) Generic cyclic chemical residue representing peak 1438 cm-1 on bacteria-free coal spectrum and C) Oxidised form of B at 1736 cm-1 found on P. fluorescens and coal spectrum, indicative of oxidation of coal by P. fluorescens. Possible oxidation (C=O) outcomes are a) aldehyde or ketone, b) ester and/or c) carboxylic acids. R1-R6 denote potentially different hydrocarbon side chains.

71
A likely agent that may have caused oxidation of the coal analysed is the peroxidase enzyme from P. fluorescens (Lenhoff and Kaplan 1956). This hypothesis was investigated by incubating a polished bituminous coal in various concentrations of aqueous peroxidase and applying ATR-FTIR to the coal surface to look for changes in the C=O bond vibrations indicative of oxidation of organic material. Unlike the P. fluorescens measurements, there was no protein from cellular aggregates on the coal surface, so the amide I C=O peak was absent. As a result, a small C=O peak observed between 1706-1695 cm-1 was evident and appeared to be characteristic of oxidised organic material native to coal, but distinct from the higher frequency biogenic C=O observed when microorganisms were present. A three-Gaussian peak fit with peak positions constrained between 1732-1714 cm-1 (peak 1, C=O), 1706-1695 cm-1 (peak 2, C=O), and 1600-1593 cm-1 (peak 3, C=C) produced calculated spectra in excellent agreement with the experimental data (Table 2.3; see Appendix IV for the entire concentration range).

The C=O stretch found in the range 1732-1714 cm-1 and assigned as peak 1 was only observed when peroxidase was present (Table 2.3). The peak found at 1736 cm-1 in the P. fluorescens and coal spectrum was slightly outside of the range of peak 1 assigned in the peroxidase and coal measurement, but the latter high frequency strongly suggested that P. fluorescens was oxidising the coal surface via a peroxidase pathway. Furthermore, the ratio of peak 1 (biogenic C=O) to peak 2 (native coal C=O) stabilized at 0.024 mg/mL peroxidase, suggesting that this concentration was sufficient to complete the oxidation of the coal surface under the incubation conditions of this experiment. The high concentration of peroxidase (0.24 mg/mL) did not increase the ratio of biogenic C=O to native C=O on the coal. A possible explanation is that diffusion of peroxidase into the surface was limited in terms of depth and time. An increase in peroxidase concentration would correspond to a faster rate of diffusion until the surface region was saturated with peroxidase and oxidation could not proceed faster. This suggests that the production of oxidative enzymes by P. fluorescens did not fully determine coal oxidation, rather other

72 factors such as the bioavailability of the coal to the bacteria may have lead to coal degradation.

With regards to cell attachment on coal surfaces, the SEM micrographs (Figures 2.2-2.6) showed that there were differences in the adhesion and colonisation behaviour of P. fluorescens on bituminous coal in the presence and absence of an external carbon source.

Extensive biofilm formation on coal by P. fluorescens was observed when glucose and casamino acids (CAA) were present (Figure 2.2). Formation of complex architecture consisting of compacted cells with channels and voids, as well as exopolymeric substances (EPS) interlinking the cells was observed, as normally expected in the formation of biofilms by these bacteria (Allison et al. 1998). This was also similar to previous biofilm analyses of wild type P. fluorescens strains conducted on glass or polystyrene in the presence of glucose and CAA where extensive biofilm formation was observed (Hinsa and O'Toole 2006; Klausen et al. 2003; Read and Costerton 1987). CAA in particular are known biofilm promoting agents and their presence even without glucose in the medium was sufficient for biofilm formation by P. fluorescens on coal in this study.

The extensive biofilm could only be observed with P. fluorescens in glucose and CAA medium, and not in medium with only glucose or without glucose and CAA. There were clear differences in attachment and biofilm formation of P. fluorescens on coal between nutrient-rich and nutrient-poor medium, which implies that where external available nutrients (i.e. glucose and CAA) were present, P. fluorescens may have used the coal surface as an inert substratum similar to glass and polystyrene. However, since coal is rich in polyaromatic and straight-chained hydrocarbons, known substrates for P. fluorescens (Barathi and Vasudevan 2001; Bugg et al. 2000; Leblond et al. 2001), it is possible that the cells could have utilised coal as a carbon-yielding substratum. The FTIR results showed that P. fluorescens could oxidise parts of the coal surface in 24 h in the absence of glucose and CAA. Thus, in the glucose and CAA-rich medium, cells may have been simultaneously using the additional nutrients and
73 oxidising coal to utilise it as a carbon source once the more bioavailable glucose and CAA were depleted. This may have been advantageous at this stage, as the cells would have been fully matured as biofilms, increasing their ability to utilise the low-bioavailable coal by further increasing coal oxidation.

Apart from the extensive biofilm formation on coal in the glucose and CAA-rich medium, there were also random distributions of individual cells or small clusters of cells (Figure 2.2D). The individual cell attachment could be observed on coal in all three media types (i.e. glucose and CAA, glucose-only, without glucose and CAA. See Figures 2.2D, 2.3A and 2.4A for respective images). The individually attached cells in glucose and CAA medium were most likely in the initial biofilm stage of attachment and would form extensive and mature biofilms when incubated for longer than 24 h. Based on SEM alone, it is unknown if the coal surface on which cells formed extensive biofilms were any different chemically from those where the cells were individually attached. However, it can be speculated that the topography on which biofilms were formed was more advantageous than that where the individual cells were attached. As observed in previous studies, surface topography is influential in the level of attachment of the cells on a particular surface (Diaz et al. 2010; Mitik-Dineva et al. 2008; Mitik-Dineva et al. 2009a; Kroupitski et al. 2011; Mitik-Dineva et al. 2009b).

Apart from the random attachment of individual cells, cell clusters could also be observed albeit more sporadically and particularly in more hollowed surfaces. These dents could be seen as ‘pockets’ for the cells to aggregate into, which may be an advantage for the cells to adhere to coal. A study conducted on Salmonella typhimurium distribution on lettuce leaves showed that the bacteria were more frequently attached to abaxial parts of the lettuce leaves, which had rougher topography than its adaxial counterparts with relatively smoother surface (Kroupitski et al. 2011). Thus, a similar effect may have occurred in these samples, where coal surfaces are naturally topographically complex.

Furthermore, random occurrences of elongated cells could be seen across the coal surface in the glucose-only medium (Figure 2.3B). These cells were approximately three to four times longer than the usual length of the rod-shaped
74 P. fluorescens cells while maintaining a similar width. Cell elongation by P. fluorescens has been observed in previous studies mainly as a result of stress conditions such as antibiotic presence, temperature, agitation, surface external polarization and nutrient limitation (Busalmen and de Sanchez 2003; Diaz et al. 2007; Korber et al. 1994; Sillankorva et al. 2011; Steinberger et al. 2002). Since CAA, which comprises of a mixture of amino acids and peptides, was omitted from the medium, this may have created a nitrogen limitation that induced some of the cells to elongate.

Steinberger et al. (2002) found that when P. aeruginosa was subjected to low carbon and nitrogen in the media, the cell length extended further, which differed from a key finding whereby P. fluorescens became shortened during starvation (Van Overbeek et al. 1995). The former study had included low bioavailable carbon (i.e. hexadecane), whereas any carbon source was strictly limited in the latter. Interestingly, Steinberger et al. observed that cells extended the longest when grown in a minimal medium with hexadecane as the sole carbon source than in other nutrient-limiting media. This finding was similar to P. fluorescens behaviour on coal, in which cells became longer when nutrients became more limited (Table 2.3). Cells extended to greater lengths in medium without glucose and CAA than when they were in glucose-only and glucose and CAA media. Since coal is a low bioavailable carbon source, the extreme cell elongation observed in P. fluorescens on coal without external carbon and nitrogen could represent an adaptation to enhance acquisition of carbon and nitrogen. Based on the supernatant imaging, no cells were shown to elongate in their planktonic state in medium without glucose and CAA (Figure 2.8). Thus, this implies that the elongation was coal surface-specific and could have resulted from a general stress response to low amounts of nutrients in the medium.

In addition to cell extension, mixtures of the long and short P. fluorescens aggregates were formed, complete with water channels and EPS formation that resembled microcolonies of biofilms (Figures 2.4B, 2.5, 2.6). Some microcolonies were observed to ‘engulf’ sub-particles of coal (Figure 2.6). These observations were exclusive to cells grown without glucose and CAA and
75 on coal (glass and pebble controls did not show similar results). The cause behind microcolony formation is unclear; the sophisticated channel and EPS formation ensured that the formation was not simply driven by physical influence such as agitation. Since the control surface, in particular the topologically similar pebble did not produce such results, topography cannot be considered as a main factor. This suggests that both cell elongation and microcolony formation could be seen as a mechanism for stress response and utilisation of carbon from coal by P. fluorescens. The ‘engulfing’ behavior by P. fluorescens on coal only without external carbon and nitrogen source also strengthens these speculations.

To further investigate the attachment behaviour of P. fluorescens on coal, physico-chemical factors, in particular, surface hydrophobicity and thermodynamics of adhesion, were tested on P. fluorescens and coal surfaces. Contact angle measurements of water, formamide and diiodomethane, and the Gibbs free energy of adhesion calculations were conducted in accordance to the Lifshitz-van Der Waals and acid-base interactions. Direct cell counting using SEM was also performed to sync with the hypothetical thermodynamic calculations.

In this study, water contact angles were used to compare hydrophobicity of one surface to another. The higher the contact angle of a surface, the more hydrophobic it is and vice versa. In general, P. fluorescens exhibited highly hydrophilic surfaces across normal (glucose-fed), starved and killed cells based on the low water contact angles (10-14o) for all surfaces (Figure 2.10). However, glucose-fed P. fluorescens were significantly (P<0.05) more hydrophilic than the starved and killed cells. P. fluorescens is known to be hydrophilic (Abbasnezhad et al. 2008; Dorobantu et al. 2004), however both starvation and killing treatments may have compromised the cell membranes, making it more permeable for release or uptake of hydrophobic components that were present inside the cell or from the environment into the membrane. Further, P. fluorescens is known to produce hydrophilic lipopolysaccharide on its outer membrane (Williams & Fletcher, 1996). This may be lost from the cell surface during starvation or death, which caused an increase in hydrophobicity.
76 P. fluorescens is also known to produce biosurfactants, of which the rhamnolipids are highly hydrophilic (Nguyen and Sabatini 2011; Vasileva- Tonkova et al. 2011). This could have been one of the factors towards a hydrophilic P. fluorescens cell surface. Compromised P. fluorescens cells may have difficulty in upholding and/or producing hydrophilic surfactants, contributing to its increased hydrophobicity.

When compared against most coal surfaces, P. fluorescens was significantly (P<0.01) more hydrophilic (Figure 2.11 and Table 2.4) particularly when compared to the untreated bituminous coal, which was the most hydrophobic surface tested. The high hydrophobicity in bituminous coal was not surprising since it consists of a relatively higher proportion of packed aromatic rings and lower oxygen content compared to its sub-bituminous and lignite counterparts (Hofrichter and Fakoussa 2001). This also makes bituminous coal more difficult than lower ranked coals to breakdown by microorganisms as aromatic rings are less bioavailable.

However, the high hydrophobicity of bituminous coal decreased when it was chemically or biologically oxidised. Nitric acid, hydrogen peroxide and peroxidase treatments significantly lowered its surface contact angle and therefore its hydrophobicity. The increase in hydrophilicity was also shown in other studies involving oxidation of bituminous coal through high temperature

(artificial weathering) and KMnO4 (Ding 2009; Pawlik et al. 2004). KMnO4 coal oxidation creates –C—O— and –COO— hydrophilic functional groups ((Ding 2009) resulting in a chemical structure similar to a lower ranked coal, e.g. lignite (Hofrichter and Fakoussa 2001). Interestingly however, lignite was more hydrophobic (due to its higher contact angle) than the chemically treated bituminous coal in this study. This implies that the chemical treatments had strongly oxidised the bituminous coal surface, rendering it more hydrophilic than the untreated lignite. Presumably, chemical oxidation of lignite would result in a much more hydrophilic surface than the oxidised bituminous coals.

P. fluorescens treatment of bituminous coal also substantially increased its surface hydrophilicity (Figure 2.11). At first, the contact angle of coal was
77 measured with cells attached (as in Figure 2.4), resulting in an inaccurate value of coal hydrophobicity after biogenic oxidation. The reduced hydrophobicity measured in this instance may have resulted from the interfering reduced hydrophobicity of the cells. For this reason, further contact angle measurements were carried out after the cells were scraped from the coal. The hydrophobicity of coal doubled after removing P. fluorescens from its surface, but it was still well below that of the non-treated bituminous coal. This result implies that either P. fluorescens indeed oxidised the coal surface or that it left behind a strongly adhered layer of biosurfactant. A similar observation was made by Bunster et al. (1989) on P. fluorescens increasing the hydrophilicity of a leaf surface. The reduced hydrophobicity caused by P. fluorescens on the leaf was prolonged despite a decrease in cell density on the surface, which was likely caused by the surfactant production of the strain that continued to adhere to the surface.
However given the previous ATR-FTIR observations for this sample, it is clear that an oxidation event took place on the surface of this bituminous coal, which may also lowered the coal surface hydrophobicity.

The hydrophobicity of coal and P. fluorescens surfaces obtained by contact angle measurements were used to determine the surface energy components for each surface and to predict the surface energy of adhesion between two surfaces (Tables 2.5 and 2.6). Based on this study, all P. fluorescens surface types (normal, starved and killed), together with P. fluorescens-treated coal had the highest surface free energy. This was followed by nitric acid-treated coal, peroxidase-treated coal and glass surfaces. The similarities between all these surfaces are that they exhibit a high hydrophilicity (based on contact angle) and had relatively higher electron donating capacity γ - than other surfaces. Thus, the positive surface free energy of adhesion, ∆GAdh (Table 2.6), between P. fluorescens and the above-mentioned surfaces means that their attachment to one another was highly unfavourable.

In contrast, surface attachment of P. fluorescens to the untreated, peroxide- treated and P. fluorescens-scraped bituminous coal and lignite was favourable due to the negative surface free energy of adhesion. All of these surfaces were more hydrophobic and had a significantly lower electron donating capacity (but
78 not necessarily overall surface energy) than the more hydrophilic surfaces. In general, microbial adhesion to a particular surface increases when the surface is higher in hydrophobicity and has a lower surface energy. This generalisation, however, is not always the case, as the balance of the interfacial free energy components of the two surfaces needs to be taken into account.

Although the untreated bituminous coal exhibited the greatest hydrophobicity and lowest surface energy, the greatest adhesion by P. fluorescens was not to this surface but to lignite. Lignite was lower in hydrophobicity and had a higher surface energy than bituminous coal, however, P. fluorescens was more attracted to lignite than bituminous coal or other hydrophobic surfaces. This is perhaps due to a higher electron accepting capacity γ + and Lifshitz-van der Waals component γ LW on lignite, which accounts for its greater interactions with P. fluorescens that has a higher γ - and γ LW. Therefore, the interaction between P. fluorescens and the coal substratum was stronger when both surfaces had a high γ LW and an opposing electron donor/acceptor capacity. Bituminous coal, although highly hydrophobic, had zero γ + and lower γ LW, which resulted in a similar surface energy of adhesion (with P. fluorescens) to peroxide treated coal that has a lower hydrophobicity but high γ + and lower γ LW. Thus, when determining thermodynamic interactions between P. fluorescens and coal, the hydrophobicity and surface energy components (γ LW, γ + and γ -) rather than just the overall surface energy of the two surfaces needs to be considered.

In terms of the types of P. fluorescens cells influencing its adhesion on coal, it seems that normal (glucose-fed) cells would adhere the strongest to a surface (of favourable adhesion) than starved and killed cells, despite the former being more hydrophilic. A relatively higher apolar or Lifshitz-van der Waals component γ LW was found on normal P. fluorescens surfaces that supports stronger adhesion to other substrata of similar or higher γ LW value. The value of γ LW for normal P. fluorescens was close to what is expected for many bacterial cells (Sharma and Rao 2002). Hence the lower γ LW obtained for starved and killed P. fluorescens shows that both cells have had their membrane compromised that affected not only its hydrophobicity but also surface energy.
79 Interestingly, this observation was more prominent in starved than killed cells, which contributed to a thermodynamically lower adhesion of the former to hydrophobic coal surfaces. This implies that in a carbon and nitrogen-limiting environment, starved P. fluorescens cells have a lesser ability to adhere to coal than normal cells, although cell adhesion would still take place (due to the negative adhesion energy). These results support the SEM observations.

Furthermore, the attachment of P. fluorescens to P. fluorescens- treated/attached coal and also to its own surface was found to be thermodynamically unfavourable. This may explain the limited aggregation or biofilm formation by P. fluorescens observed on coal in media with limited bioavailable carbon and nitrogen. Therefore, extensive biofilm formation on coal would have resulted from expression of biological factors used to overcome the potential energy barrier of cell aggregation. Exopolysaccharide, lipopolysaccharide and microbial appendages production, e.g. pilli, are common in P. fluorescens and have been shown as a part of its adhesion mechanisms (Abbasnezhad et al. 2011; Williams and Fletcher 1996). As demonstrated further in this study, the number of killed P. fluorescens cells attached to coal was considerably smaller than live P. fluorescens (Figure 2.13). This showed that although the thermodynamics of adhesion is an essential factor to consider particularly during the first cell-coal surface contact, biological determinants are equally important to determine cell adhesion on coal for a longer period of time. The fact that P. fluorescens could still attach to glass (Figure 2.9) despite the positive surface free energy of adhesion also implies that P. fluorescens is able to attach to any surface through biological means which are strong enough to overcome the physico-chemical barriers. This phenomenon was also mentioned by Boks et al. (2008), in that microbial appendages play an influential role in cell attachment despite the high-energy barrier the cells need to overcome. Some appendages are capable of extending hundreds of nanometers away from the cell surface to reach the substratum and achieve adhesion. However the surface properties of these adhesive factors are difficult to detect by physico- chemical studies alone due their nanoscale and hence might not be reflected in surface energy calculations.

80 To account for the theoretical aspect of ∆GAdh values in Table 2.6, cells attaching to the various coal types were directly counted after 24 h of incubation in M9 without glucose and CAA (Figure 2.12). Reassuringly, lignite had the highest number of starved cells attached, followed by peroxide-treated and untreated bituminous coals. This corresponded well to the ∆GAdh calculation where the most negative, hence the most favourable adhesion for starved P. fluorescens would be that with lignite, followed by untreated bituminous and peroxide-treated bituminous coals. The differences in the ∆GAdh and direct cell attachment between the latter two coals are not significant, thereby the order of which of the two surfaces is more adherent to cells is not critical. Nitric-acid treated and peroxidase-treated bituminous coal had the lowest number of cells attached, which reflects the positive ∆GAdh values indicating unfavourable adhesion. Nevertheless, the order to which coal surface had the least attachment between the two surfaces was reversed in ∆GAdh and cell attachment measurements. This shows that although ∆GAdh calculation gives a generally reliable prediction of cell attachment, it may be irrelevant to make comparisons between surfaces that have high positive ∆GAdh values (i.e. unfavourable attachment).

Previous studies have often cited and/or demonstrated the use of chemically treated coal, particularly from nitric-acid treatment, as an enhancement to coal degradation (Achi and Emeruwa 1993; Igbinigie et al. 2008; Laborda et al. 1999; Machnikowska et al. 2002; Maka et al. 1989; Strandberg and Lewis 1987a; Strandberg and Lewis 1987b). However, the favourable results obtained from the studies were evident for fungi and not bacteria. In cases where pretreated coal showed favourable degradation by bacteria, the coal used was lignite and not bituminous coal (Gupta et al. 1990; Maka et al. 1989; Polman et al. 1994b; Quigley et al. 1989). Interestingly, almost all bacterial degradation reported from the literature were of pretreated lignite, untreated lignite and untreated bituminous coal (Fakoussa 1981; Fakoussa 1988; Gao et al. 2012; Laborda et al. 1997; Monistrol and Laborda 1994; Strandberg and Lewis 1987a; Strandberg and Lewis 1987b). Only one study, however, conducted by Laborda et al. (1999), showed successful bacterial degradation of nitric acid-treated

81 bituminous coal. Another study performed by Kitamura et al. (1993) showed favourable bacterial degradation of peroxide-treated bituminous coal.

The lack of previous findings on bacterial degradation of nitric acid-treated bituminous coal correlates with the results in this study, where adhesion between P. fluorescens and nitric acid-treated bituminous coal was thermodynamically unfavourable compared to the favourable adhesion on lignite, untreated bituminous coal and peroxide-treated bituminous coal (Table 2.6. and Figure 2.12). This was due to the high surface energy of the nitric acid- treated coal, which was similar to that of peroxidase-treated coal, on which cells were least attached. Thus, the unfavourable adhesion shown between P. fluorescens and nitric acid-treated bituminous coal may imply a limitation on degrading the treated coal by bacteria. However, nitric acid treatment of coal results in a more oxidised form of coal. This means that chemically speaking, nitric acid-treated coal is more favourable for degradation due to the higher number of oxygen-linked moieties introduced. Therefore, it is likely that certain bacteria would utilise biological mechanisms to overcome the energy barrier in order to degrade the oxidised coal. Further investigation is necessary to confirm this speculation.

As mentioned earlier, many coal degradation studies showed favourable degradation of nitric acid-treated coal by fungi, which conflicted with the attachment results in this study. This may imply differences in surface properties and mechanisms between fungi and bacteria that enabled the former to effectively degrade nitric acid-treated coal. This will be examined further in Chapter 4.

Results from the thermodynamic analysis of cell aggregation (cell-to-cell attachment) implied that the microcolony formation of P. fluorescens observed exclusively on coal in media without glucose and CAA (Figure 2.4-2.6) could not have been driven by physico-chemical factors, but was rather influenced by cell biology. What drove P. fluorescens to form ‘mini’-biofilms on coal without the support of excess carbon and nitrogen is unclear. However, the need to derive carbon from coal may have led to this response by means of surfactants and
82 other cell adhesion factors. Since more bioavailable carbon like glucose was absent, the cells were forced to form much smaller aggregates than in glucose- rich medium. However, together with cell elongation as observed in this study, these strategies could constitute a part of the mechanisms of adhesion of P. fluorescens on coal to utilise it as both substratum and carbon source. Further investigation particularly on microbial adhesin production by this bacterium on coal would benefit this research area.

In this study, the surface attachment and oxidation of coal by P. fluorescens were described. P. fluorescens Pf-5 was shown to oxidise bituminous coal through the formation of C=O bond, which is in line with previous studies involving P. fluorescens and hydrocarbon degradation. The oxidation corresponded closely to coal chemically oxidised by peroxidase, which suggested the production of similar oxidising agents by P. fluorescens. The attachment of P. fluorescens to bituminous coal was described and evaluated through direct observation (SEM) and physico-chemical analyses. Under limiting bioavailable carbon conditions, P. fluorescens exhibited unique morphological behaviour on coal, which suggests the unique role of coal as a substratum for attachment and substrate for growth. To the best of our knowledge, this study is the first that investigates and describes bacterial attachment on coal in detail, which opens up more research on coal-microbe interactions and provides some background for similar studies on other hydrocarbons. Further investigation on the biological factors concerning microbial adhesion on coal is necessary to better understand the mechanism of adhesion and degradation of coal.

83 3 Isolation of coal-degrading fungi from an abandoned shale mine in The Wolgan Valley and Newnes forest, New South Wales

3.1 Introduction

Coal is one of Australia’s main and important natural resources. The country contains approximately 76.4 Giga tonnes (Gt) of black and brown coal reserves, the fourth largest coal reserve in the world (Bureau of Resources and Energy Economics 2010). Further, there is still a vast area that is unexplored for coal reserves, potentially amounting to one trillion tonnes of coal (Bureau of Resources and Energy Economics 2010). These resources have secured Australia as one of the current (and future) major coal producers and exporters in the international arena.

Coal is also the main energy resource for the country, providing approximately 75% of its electricity production through coal-fired power stations (Bureau of Resources and Energy Economics 2010). More recently however, there has been increased interest in capturing methane generated within coal seams, which would serve as a major alternative to coal burning for electricity both locally and overseas (New South Wales Department of Trade and Investment 2013). Coal bed methane (CBM) in Australia is a largely untapped resource, having an Economic Demonstrated Resource (EDR) of approximately 35,055 Petra Joules (PJ) per year, which is worth about 175 years of gas reserve (Bureau of Resources and Energy Economics 2012). This makes CBM in Australia a promising avenue for the country’s economic growth.

A number of studies have shown interest in the biogenic origin of CBM in Australia (Ahmed and Smith 2001; Faiz and Hendry 2006; Li et al. 2008). As evidenced by isotopic data, CBM originated through microbiological activity across a large region of Australia (Faiz and Hendry 2006). This shows the
84 methanogenic conversion potential that could derive from coal through the action of microorganisms. The microbial conversion of coal, as a biotechnological tool, would serve as a sustainable approach in utilising coal for methane in comparison to CBM capturing or coal mining.

Strapoć et al. (2008) and Jones et al. (2010) have outlined the potential mechanisms by which microbial conversion to methane is derived. Three main stages are involved in this process, namely, the initial fragmentation of coal macromolecules, anaerobic oxidation and/or fermentation of the fragmented molecules and methanogenesis. Coal fragmentation is seen as the rate-limiting step due to the recalcitrant molecular structure, which is more difficult to break down as the coal rank increases. Both Strapoć et al. and Jones et al. have focused on microbial fermentation as a main process to defragment the coal. However, it is known that aerobic coal degraders can alter the coal macromolecule to produce similar smaller entities, which would then feed into the next step of the coal conversion process. Thus, the application of coal degrading isolates is seen as a useful bioaugmentation tool for coal to methane generation (more so for bituminous coal than lignite). No such attempt has been made to isolate bituminous coal degrading microorganism native to Australia for this purpose, though there has been a successful application of a known peroxidase-producing fungus, Phanerochaete chrysosporium, to degrade Australian lignite coal (Ralph and Catcheside 1994).

The isolation and screening of aerobic coal degrading microorganisms have been conducted for a number of decades outside Australia. Fakoussa (1981) has discovered that certain bacteria are able to utilise parts of coal as a sole carbon source, whilst Cohen and Gabriele (1982) demonstrated the solubilisation of lignite by wood-decaying basidiomycetes Trametes versicolor and Poria monticola. Whilst screenings have isolated both bacteria and fungi (Kitamura et al. 1993; Monistrol and Laborda 1994), filamentous fungi including basidiomycetes and deuteromycetes, have attracted most interest (Bublitz et al. 1994; Granit et al. 2007; Hofrichter and Fritsche 1997; Igbinigie et al. 2010; Igbinigie et al. 2008). Both groups exhibit similar properties of filamentous cell morphology and are associated with the production of extracellular enzymes
85 linking to coal degradation (Hofrichter and Fakoussa 2001; Laborda et al. 1997; Monistrol and Laborda 1994; Wilmann and Fakoussa 1997b). These properties are more common in fungi than bacteria thus could explain the preferential isolation of the former group.

More studies have focused on lignite and humic acid degradation (Granit et al. 2007; Grinhut et al. 2011; Hofrichter and Fritsche 1997; Kitamura et al. 1993) than bituminous coal (Bublitz et al. 1994; Hofrichter et al. 1997b; Igbinigie et al. 2008), which reflects the difficulty in the microbial transformation of the latter. Lignite is more amenable to biodegradation due to the presence of more oxygen-linked moieties found in various functional groups (e.g. carboxylic acids, ethers), which can be more easily fragmented compared to the more aromatically condensed structure found in bituminous coal (Strapoć et al., 2008). Pre-treatments of bituminous coal by chemical agents such as nitric acid and hydrogen peroxide in order to oxidise the coal were commonly used in many isolation studies (Igbinigie et al. 2008; Kitamura et al. 1993; Monistrol and Laborda 1994). This shows that isolating microorganisms that can degrade untreated bituminous coal over the more chemically or naturally oxidised coal is challenging. However, it has been achieved in the past, as demonstrated in the above papers, and has thus provided the basis for this study.

This chapter describes the isolation and identification of bituminous coal- degrading microorganisms, and in particular fungi from a native Australian environment. Several steps were employed in the screening: 1) Growth on coal agar and nutrients (with no additional carbon source), 2) Growth on coal silica and nutrients (with no additional carbon source or impurities), and 3) Coal degradation capacity of the isolates. Untreated and nitric acid treated bituminous coals were used alongside lignite coal to compare degradation efficiency. Active isolates were then identified using rRNA gene amplification and sequencing. Oxidative enzyme tests were also conducted for selected isolates. The most outstanding isolate based on all screenings was then used in the following chapter for further characterisation. This included an investigation into its colonisation and degradation behaviour on coal that is in line with the work described in Chapter 2.
86
3.2 Material and Methods

3.2.1 Coal preparation

In preparation for coal agar and coal silica plates, bituminous coal (for coal origin see Section 2.3.1) was pulverised into powder with the average diameter size of approximately 60 μm using an industrial milling machine (Standard Ring Mill Batch Pulveriser, Rocklabs, New Zealand).

For coal colonisation and degradation assays, the bituminous coal sample was fragmented into small pieces of approximately 1 x 0.5 x 0.5 cm. Four types of coal were used for this assay: 1) Untreated (raw) bituminous coal, 2) Nitric acid treated bituminous coal (refer to Section 2.2.3 for preparation), 3) Lignite coal from the Loy Yang open cut mine in Victoria, Australia, and 4) Lignite coal sourced from Kalimantan, Indonesia. For lignite coal, 2-10 mm pieces were used without any pre-treatment except for sterilisation. All coal pieces were dried at room temperature for 2 days before being sterilised under ultra violet light for 30 min on each side of the coal piece.

3.2.2 Elemental analysis of coal

Elemental (carbon, hydrogen, nitrogen, oxygen, sulphur, phosphorus) analyses were conducted for the four types of coal used in this study. For C, H, N, O and S analyses, a TruSpec Micro CHNSO determinator (Leco Corporation, Michigan, USA) was employed. Coal ash was used to analyse P content through a Genesys 6 UV-Visible spectrophotometer (Thermo Electron Corporation, Madison, USA). Samples were analysed in duplicate and content calculated on a dry, ash-free basis percentage (% daf). The analyses were conducted according to Australian Standard Methods AS1038.3, AS1038.6.4, AS1038.6.3.3 and AS1038.9.1. Differences between the replicates of samples were less than 1% of standard deviation.

87 3.2.3 Sample collection

Samples for the screening of coal-degrading isolates were collected in May 2010 from the Old Oil Shale Mine in The Wolgan Valley and in the vicinity of Newnes State Forest, New South Wales, Australia. The mine was in operation 100 years ago with the environment experiencing widespread hydrocarbon contamination. It is hypothesised that this would lead to enrichment of hydrocarbon-degrading fungi and bacteria, therefore justifying the choice of sampling environment. Table 3.1 shows a list of samples that were collected and numbered accordingly. The samples consisted of coal, wood, and soil samples with evidence of microbial growth (mostly fungi). These samples were transferred into sterile falcon tubes or airtight plastic bags before being further processed for inocula preparation.

Table 3.1 A list of samples obtained from Wolgan Valley, NSW, as inocula for the isolation of coal-degrading microorganisms. The specific description and location of the samples are included.

Inoculum No. Description Location

5 lumps of coal and fungi test coal mine

6 lumps of coal and fungi ash pile

7 soil slag heap

8 wood coal washing tank

9 coal coal washing tank 10 flakes of rotten wood soil within the Newnes forest

11 coal and soil coke oven 12 rotten wood soil within the Newnes forest

13 coal and soil test coal mine pond

14 coal and soil ash pile

15 ash ash pile/wall 16 coal and soil test coal mine

88 3.2.4 Inocula preparation

To separate cells from the original carbon source collected from the environment, all samples were initially suspended in an M9 minimal salts medium with reduced phosphate concentration (10 mM) at pH 7.5 (see Appendix I). The samples were vortexed for 30 sec and then sonicated at 100 W for 2 min. These samples were then centrifuged at 50 x g for 2 min and the supernatant was transferred into sterile falcon tubes. This separation method had been tested against several other protocols and was found to be the least damaging to cell viability, based on LIVE/DEAD BacLight Bacterial Viability staining (Invitrogen, Molecular Probes Inc., USA).

3.2.5 Initial screening for coal-degrading microbes on coal agar

Pre-prepared inocula from environmental samples were inoculated at different dilutions (from 10-1 to 10-8) onto five different types of agar containing the following: 1) Coal and agar only, prepared in water (isolates obtained from these plates were labelled with an ‘R’ as the first letter of identity), 2) Coal in M9 agar and 1 x trace element solution (TES) (see Appendix I) (isolates labelled as ‘B’), 3) Coal in M9 agar and 6 x TES (isolates labelled as ‘Y’), 4) Coal in BIII fungal agar (see Appendix I) and 6 x TES (isolates labelled as ‘G’), and 5) Agar only. Each medium type, except for the agar only control, contained 10 mg/mL of pulverised untreated bituminous coal. All medium types were adjusted to pH 5.5 to promote fungal growth. For each plate, 100 μL of each inoculum at a given dilution was spread aseptically on to the agar plate and sealed with parafilm. All plates were incubated at 22 oC in the dark in a contained but aerated space for at least two weeks. The plates were observed for growth every 3 days.

3.2.6 Further screening of coal-degrading isolates on coal silica
A second screening using coal silica was conducted in order to remove isolates that were potentially utilising carbon from impurities in the agar or the agar itself,

89 and thereby selecting those that could utilise coal as their sole carbon source. Coal silica plates were prepared according to a protocol outlined by Thatcher and Weaver (1974), with some minor changes. For 100 mL of dH20, 13.4 g of

Na2SiO2.9H20 was dissolved and the pH adjusted to 10 by adding cation exchange resin (AG 50 WX-8, Bio-Rad Laboratories, USA) with constant stirring. After 30 min of mixing, the resin was added to the solution to re-adjust the pH to 10 and was then removed through filtration. In constant stirring, the pH was lowered to 5.5 by adding 5 N phosphoric acid. The ingredients, according to the four different agar types (refer to section 3.2.4) excluding agar, were then mixed into the silicate solution. Approximately 25 ml of the mixed solution was transferred into a sterile 90 mm petri dish and immediately placed in a 60oC oven for 45 min to 1 hour to solidify. The plates were then cooled at 22 oC overnight before being used. All ingredients except coal were pre- sterilised through autoclaving and the procedures were conducted aseptically.

Approximately 1,200 colonies from the coal agar plates were picked using a sterile 10 μL inoculating loop and were transferred by streaking onto the coal silica plates. Each silica plate was divided into eight small sections (on average) for eight individual isolates. The inoculated plates were then sealed with parafilm to prevent them from drying, and were incubated at 22 oC in the dark and observed each week for growth for at least 6 weeks.

3.2.7 Molecular identification of coal-degrading isolates

3.2.7.1 DNA extraction of coal-utilising fungal isolates

Coal-utilising isolates were grown on Sabouraud agar and after approximately 10 days of incubation a portion of the fungal mat was scraped and suspended into 500 μL of sterile phosphate buffer in a 2 ml cryo tube containing 200 μl of 0.5 mm Zirconia/Silica beads (BioSpec Products Inc., USA), 500 μl of phenol: chloroform: isoamyl alcohol solution (25:24:1) and 800 μl of Tris-EDTA (TE) buffer. All tubes were placed in a FastPrep® cell disrupter (FP120, Thermo Savant, Canada) for 30 sec at 5.5 min s-1 and then centrifuged at 16,000 x g for
90 4 min at room temperature. The aqueous phase was transferred into a clean 1.5 ml tube before adding 500 μL of chloroform: isoamyl alcohol (24:1). The tubes were centrifuged at 16,000 x g for 4 min. The aqueous phase was recovered and mixed with 0.5 x volume of 7.5 M NH4Ac and two volumes of isopropanol. The tubes were then incubated at room temperature for 10 min and centrifuged at 16,000 x g for 20 min at 4 oC. The supernatant was removed and the pellet washed with 80% ethanol solution before being reconstituted in 30 μL TE buffer. DNA yields were measured using a NanoDrop spectrophotometer (ND-1000, Thermo Fisher Scientific Inc., USA).

3.2.7.2 Polymerase chain reaction (PCR) amplification of fungal ITS regions

ITS1 and ITS2 regions were sequenced using primer pair ITS1/ITS4 (see Appendix II for primer sequences and references). Only a subset of the isolates could be identified using this primer pair alone. Therefore, amplification of ITS1 and ITS2 regions separately was carried out using ITS1/ITS2 and ITS3/ITS4 primer pairs, respectively.

The PCR mixture (25 μL total volume) for each isolate consisted of 12.5 μL of 2x Promega PCR Master Mix (Promega Corp., USA), which contained 50 units/ml of Taq DNA polymerase supplied in a proprietary reaction buffer (pH 8.5), 400 μM dATP, 400 μM dGTP, 400 μM dCTP, 400 μM dTTP and 3 mM

MgCl2, 0.5 μL of 10 μM of each forward and reverse primer, 10.5 μL of molecular biological grade water, and 1 μL of the extracted fungal isolate DNA.

All amplifications were conducted using an MJ Mini Personal Thermal Cycler (Bio-Rad Laboratories Inc., USA). The thermal cycling profile and reference for the amplification using each primer set is as follows:

1. ITS1/ITS4 (White et al. 1990): Initial denaturation at 94 oC for 5 min; 30 cycles of denaturation at 94 oC for 45 sec, annealing at 60 oC for 30 sec, extension at 72 oC for 45 sec, followed by a final extension at 72 oC for 10 min. 2. ITS1/ITS2 (Kumar et al. 2005): Initial denaturation at 96 oC for 10 min; 30

91 cycles of denaturation at 95 oC for 1 min, annealing at 60 oC for 1 min, extension at 72 oC for 1 min, followed by a final extension at 72 oC for 10 min. 3. ITS3/ITS4 (Kumar et al. 2005): Initial denaturation at 94 oC for 4 min; 30 cycles of denaturation at 94 oC for 30 sec, annealing at 60 oC for 55 sec, extension at 72 oC for 1 min, followed by a final extension at 72 oC for 4 min.

All PCR products were verified using electrophoresis of the amplicons on a 1.5% agarose gel stained with SYBR® Gold nucleic acid gel stain (Molecular Probes Inc., USA). Products were visualised by transillumination with a Molecular Imager® Gel Doc™ XR System (Bio-Rad Laboratories Inc., USA).

3.2.7.3 Sequencing of the ITS regions of the fungal isolates

PCRs were prepared as described above (section 3.2.7.2) except in reaction volumes of 100 μl (i.e., four times all reagents and DNA). Products were verified using electrophoresis before undergoing DNA purification using the DNA Clean & ConcentratorTM-5 kit (Zymo Research Corp., USA). The purified PCR products were checked by electrophoresis to ensure that the DNA was not lost during the cleaning procedure and that the purification was successful.

The DNA concentration of the purified products was quantified using a Qubit® fluorometer (Life Technologies Australia Pty Ltd., Australia). This DNA was further amplified for sequencing using 1 μL of Big Dye® Terminator v3.1, 3.5 μL of its 5x sequencing buffer (Life Technologies Australia Pty Ltd., Australia) and 1 μM of the forward primer (i.e. ITS1 or ITS3). DNA in the reaction amounted to 20 to 50 ng. Molecular biology grade water was added to a final volume 20 μL. The thermal profile was conducted as follows: 25 cycles of 96 oC for 10 sec, 50 oC for 5 sec, and 60 oC for 4 min.

The products were purified by adding 5 μL 125 mM EDTA and 60 μL of 100% ethanol solution, followed by incubation at 22 oC for 15 min and centrifugation at 14,000 x g for 20 min. The DNA pellet was washed twice with 70% ethanol and dried at 90 oC for 1 min. Sequencing reactions were carried out at the

92 Ramaciotti Centre (University of New South Wales, Sydney, Australia) using an Applied Biosystems 3730 DNA Analyzer (Life Technologies Corp., USA).

Sequences were trimmed for quality using the 4Peaks software version 1.7.1 (Griekspoor and Groothuis 2005). Species identification was based on the results of BLAST searches for homology comparison with known sequences in GenBank (National Centre for Biotechnology Information (NCBI; http://www.ncbi.nlm.nih.gov/)). Successful identification required at least 95% of sequence overlap to a known ITS sequence of a minimum of 400 base-pairs (bp) in the GenBank database and were named at the species or genus level. Only matched sequences with the Expect (E)-value of 0.0 were deemed identified. This denotes statistical significance of a matching sequence from BLASTn searches.

3.2.8 Coal colonisation and degradation assay

Isolates grown on coal silica were further tested for the ability to colonise and degrade coal under different conditions. Positive isolates from the coal silica assay were suspended and lightly vortexed for 10 seconds in 100 μl of phosphate buffer pH 5.5. This suspension was then plated onto Sabouraud agar pH 5.5 (Appendix I) and the plates sealed with parafilm and incubated at 22 oC in the dark for 10-14 days. After the agar plate was fully covered by the fungal isolate, untreated and nitric-acid treated coal pieces of approximately 1 x 1 x 0.5 cm were placed onto the fungal mat. Phanerochaete chrysosporium, a known coal solubilising fungus, was used as a positive control for this assay. Plates were observed weekly for progress on colonisation and degradation of coal by the isolates. Images of the assay were captured by a high performance digital camera (PowerShot G12, Canon Inc., Japan) to record coal colonisation and solubilisation by the isolates. Observations were conducted on average each week for a period of approximately 6 months.

To confirm initial observations, the assay was repeated two more times using a few isolates that showed complete colonisation of both untreated and treated coal, and either each of the untreated or treated coals. A sterile negative control
93 made from polytetrafluoroethylene (gas chromatography vial caps, THC11241050, Thermo Fischer Scientific Inc., Australia) with a jagged surface, or a sterile 100 µl micropipette tip made from polypropylene (Edwards Instruments Co., New South Wales, Australia), was included in the second and third assays to confirm that coal was actively preferred for colonisation by the isolates. This was verified by observing and comparing colonisation on the control surface against colonisation on the coal.

Apart from bituminous coal, lignite coal pieces were also included in the second and third assays to compare high- and low-ranked coal degradation. Lignite coal pieces of 2-10 mm in diameter were placed alongside the bituminous coal pieces on a full-grown fungal mat and observed for solubilisation, i.e. through the formation of brown liquid on the agar plate or diffused into the agar (in the latter case a brown halo surrounding the coal is an indication of solubilisation). Observations were conducted weekly for a period of approximately 3 months.

3.2.9 Oxidative enzyme assay

Three isolates, G9o, G5o and Y7e, which exhibited positive results in the solubilisation assay, were tested for ligninolytic or oxidative enzyme activity according to the works of Granit et al. (2007) with modifications. Each fungus in three independent replicates was tested for manganese peroxidase (MnP) and laccase production, as well as the ability to degrade phenolic complexes.

The isolates were subcultured onto agar plates containing (per L): 39 g potato dextrose agar, 5 g agar and 50 ml microelement stock solution (refer to

Appendix I for recipe). For the MnP assay, 100 mg MnCl2 was added to each plate. Observation of black pigments (MnO2) surrounding the fungal colony signified Mn2+ oxidation to Mn3+. To detect laccase production, 100 mg of 2,2’- azino-bis (3-ethylbenzothiazoline-sulfonic acid) (ABTS) was added to each agar plate. A formation of dark-green coloured pigments in the agar surrounding the fungal colony indicated production of ABTS cation radicals (ABTS
+), a direct product of laccase activity (Steffen et al. 2000). To detect degradation of phenolic complexes, 0.02% (w/v) of an anthraquinone dye Remazol Brilliant
94 Blue R and an azo dye Reactive Violet 5 (both from Sigma-Aldrich Co., Australia) were each added to a plate to observe decolourisation of the dye by the fungi (Keharia and Madamwar 2002; Palmieri et al. 2005). All plates were incubated at 28 oC in the dark and were observed for changes every 2-5 days for 4-6 weeks.
3.3 Results
3.3.1 Coal elemental analysis
Table 3.2 shows the elemental analysis data for the coal samples used in this study. The analysis was done to compare the coal biodegradation outcomes in this study. The proportions of the elements in each coal sample generally differed from one another, particularly on the carbon, oxygen and nitrogen content. Carbon and hydrogen were found to be the highest in untreated bituminous coal whilst the proportions were lower in nitric acid-treated bituminous and lignite coals. In reverse, both Loy Yang and Kalimantan lignite coals had a higher proportion of oxygen than the bituminous coals. The oxygen content in nitric acid-treated bituminous coal was also much higher than the untreated bituminous coal. The nitrogen content in the treated bituminous coal was found to be the highest among other coal types. Phosphorus and sulphur content in all coal types appear to be in smaller proportions than the other elements, although they are found to be higher in bituminous coals than lignite coals.

The ratios of carbon-to-oxygen (C:O) and carbon-to-nitrogen (C:N) were also calculated for each coal sample (Table 3.2). These ratios reflect the order of the coal based on the reduction state and type of organic matter respectively. The untreated bituminous coal appeared to have the highest C:O ratio, followed by the treated samples, Kalimantan lignite, and then Loy Yang lignite. The C:N ratio was the highest in the lignites, but lower in the bituminous coals. These ratios provide good insight into coal biodegradability and were used to compare with the results from the coal colonisation and solubilisation assay obtained in this study.
95
Table 3.2 Elemental analysis data of coal samples used: 1) Untreated bituminous, 2) Nitric acid- treated bituminous, 3) Loy Yang lignite, and 4) Kalimantan lignite. All data presented were calculated to a dry, ash-free basis (% daf).

Elements Coal sample Bituminous Nitric acid- Loy Yang Kalimantan treated lignite lignite bituminous Carbon 84.40 72.73 66.06 68.50 Hydrogen 5.54 4.61 4.67 4.75 Oxygen 7.40 17.94 28.40 25.49 Nitrogen 1.90 4.08 0.64 1.08 Phosphorus 0.023 0.033 <0.001 <0.001 Sulfur 0.76 0.64 0.23 0.19 C:O 11.41 4.05 2.33 2.69 C:N 44.42 17.83 103.22 63.43

3.3.2 Screening of isolates through coal agar and coal silica

Fungi from soil, wood or coal samples were initially screened for optimal growth on coal agar in five different media. After approximately two weeks of incubation at 22 oC in the dark, growth of colonies on the coal plates could be observed across all samples in different media (Figure 3.1). A mixed range of different colony morphology types could be found on the different coal plates. In general, small (2 mm diameter), white/beige and filamentous colonies were prevalent across many different samples and media types. Medium-sized colonies (50 mm diameter) of brown and beige colour, and both filamentous and non- filamentous morphologies were observed. There was also the presence of smooth and non-filamentous colonies resembling those of bacteria. More distinctive and wider (approx. 2 cm diameter) colonies of green and brown filamentous morphologies were found on plates containing an excess of trace elements (6x concentration) than plates with only 1x trace elements (in M9 medium) or without any nutrients (coal and agar only). Amongst all media used,
96 coal plates with M9 medium and 6x trace elements were observed to have the most growth of distinctive isolates against the coal-free agar controls.

Although a large number of isolates were obtained on coal agar plates, there were also colonies growing on agar-only plates (Figure 3.1). In fact, many colonies mainly of small and white morphology on pure agar closely resembled those growing on coal plates. To exclude isolates deriving carbon and energy from agar a second screening was conducted. Approximately 1,200 colonies were transferred to coal silica plates of the same media. The isolates were selected based on different morphology types: filamentous, non-filamentous, colour, and size irrespective of the medium used in the coal agar. Colonies that closely resembled those on agar-only controls were excluded from subculturing. The sub-cultured colonies on coal silica plates were maintained at 22 oC in the dark.

Figure 3.1 Examples of isolates from a non-diluted sample grown on coal agar in different media types: A) Agar-only control, B) Coal and agar, C) Coal in M9 agar and 1x TES, D) Coal in M9 agar with 6x TES, and E) Coal in BIII agar with 6x TES. The arrows show brown coloured filamentous colonies present in media with 6x TES (D and E) and absent from other media plates. Small white colonies were present on all media types including agar-only control plate.

97 From the 1,200 sub-cultured isolates, 150 were shown to grow on coal silica plates after three weeks of incubation. Figure 3.2 shows examples of isolates that grew on coal silica.

Figure 3.2 Examples of isolates selected from previous coal agar screening grown on coal silica of the same medium. The plates were divided into seven or eight sections and each isolate was streaked once. Out of the 1,200 colonies streaked, approximately 13% successfully grew on coal silica after approximately 3 weeks of incubation. For each plate, the average colony grown on coal silica was one out of eight colonies.

A large proportion (approximately 85%) of the isolates were filamentous, whilst the rest of the colonies were hard or smooth in appearance. Also, a greater number of colonies grew on plates with nutrients than without (i.e. coal and silica only). Despite the positive outcome, no observable changes were noted on the coal physical appearance (e.g. colour) in the silica medium.
3.3.3 Identification of fungal isolates

Fifty isolates were selected for taxonomic identification through ITS sequencing and comparison with GenBank (Table 3.3). Out of 50 isolates, 37 were successfully amplified by the ITS primers.

In addition to taxonomic identity, Table 3.3 presents a summary of the coal colonisation and solubilisation capacity of the isolates, the morphology, and source/location of the fungus. Whilst a range of fungi were identified, certain genera and phyla were dominant. Ascomycetous fungi made up 91% of the total phyla, whereas Basidiomycetous and Zygomycetous fungi were 6% and

98 3%. Penicillium was the most frequently identified genus (38%), followed by Paecilomyces (14%) and Aspergillus (6%).

Interestingly, most isolates identified originated from ash piles (38%) in an abandoned oil shale mine. Samples from the ash pile included lumps of coal or soil (refer to Table 3.1 in Section 3.2.2). The sources that were less frequently associated with the identification include, in decreasing order: coal mine (16%), coal washing tank (14%), coke oven (11%), coal mine water (8%), and slag heap (5%). Additionally, with respect to the medium used for screening the isolates, M9 minimal medium with 6x trace elements (TE) (labelled as ‘Y’) gave most results (59%), followed by M9 with 1x TE (30%) (‘B’), and BIII fungal medium with 6x TE (11%) (‘G’). No isolates were found to grow only on coal (i.e. without additional nutrients).

99 Table 3.3 Molecular identification of several fungal isolates obtained in this study. The source, phylum, morphology and coal colonisation and solubilisation ability of each isolate are indicated. Isolate Taxonomic Identity Source Phylum Morphology Coal colonisation, solubilisation B6o Penicillium sp. Ash pile Ascomycota Filamentous N Y6a Paecilomyces sp. Ash pile Ascomycota Filamentous UT Y6d Paecilomyces parvisporus Ash pile Ascomycota Filamentous UT Y6e Penicillium sp. Ash pile Ascomycota Filamentous UT Y6k Paecilomyces parvisporus Ash pile Ascomycota Filamentous UT B5b Penicillium sp. Coal mine Ascomycota Filamentous N B5e Penicillium sp. Coal mine Ascomycota Filamentous UT Y7d Penicillium sp. Slag heap Ascomycota Non-filamentous N Y13b Paecilomyces lilacinus Coal mine pond Ascomycota Filamentous UT Y13d Pupureocillium lilacinum Coal mine pond Ascomycota Filamentous U Y15c Paecilomyces lilacinus Ash pile Ascomycota Filamentous UT Y15b Penicillium sp. Ash pile Ascomycota Filamentous UT B11c Aspergillus fumigatus Coke oven Ascomycota Filamentous N B14d2 Paecilomyces variotii Ash pile Ascomycota Filamentous N B16ab Penicillium glabrum Coal mine Ascomycota Filamentous N B6a Penicillium sp. Ash pile Ascomycota Filamentous N B6j Penicillium sp. Ash pile Ascomycota Non-filamentous N B6q Penicillium sp. Ash pile Ascomycota Filamentous N BFun8 Fusarium oxysporum Air in laboratory Ascomycota Filamentous UT GFun4 Cladosporium sphaerospermum Air in laboratory Ascomycota Filamentous N G5o Penicillium sp. Coal mine Ascomycota Filamentous UT, L G8n Penicillium verruculosum Coal washing tank Ascomycota Filamentous UT Y11t Umbelopsis sp. Coke oven Zygomycota Non-filamentous N Y13m Penicillium sp. Coal mine pond Ascomycota Non-filamentous N Y15b Paecilomyces lilacinus Ash pile Ascomycota Filamentous UT Y15d Ophiocordyceps heteropoda Ash pile Ascomycota Filamentous UT Y5a Aspergillus viridinutans Coal mine Ascomycota Filamentous T Y9a Emericella nidulans Coal washing tank Ascomycota Non-filamentous N Y13d Ophiocordyceps heteropoda Coal mine pond Ascomycota Filamentous U Y5c Chytridiomycota sp. Coal mine Ascomycota Filamentous UT Y9e Hypocreales sp. Coal washing tank Ascomycota Non-filamentous N Y9j Ustilago tritici Coal washing tank Basidiomycota Non-filamentous N Y15h Torrubiella sp. Ash pile Ascomycota Non-filamentous U Y11f Acremonium sp. Coke oven Ascomycota Non-filamentous N B11b Malassezia restricta Coke oven Basidiomycota Non-filamentous N Y7e Penicillium sp. Slag heap Ascomycota Filamentous T, L G9o Fusarium sp. Coal washing tank Ascomycota Filamentous U, L, B The first letter of the isolate name corresponds to the type of coal medium, where B= M9 minimal medium with 1x trace elements (TE), Y= M9 minimal medium with 6x TE, and G= BIII fungal medium with 6x TE (refer to Section 3.2.4 for further information). The number in the isolate name corresponds to the inoculum/sample number (Table 3.1) and lower case letter refers to consecutive labelling of the isolates on a plate. N= no colonisation, U and T correspond to untreated and treated coal colonisation, respectively. L= Lignite solubilisation, B= bituminous coal degradation.
100 3.3.4 Coal colonisation and degradation assay

Approximately 100 isolates were tested for their capacity to colonise and solubilise coal. Isolates were grown on a rich medium (Sabouraud) agar until a full-grown mat was achieved. Untreated and nitric-acid treated bituminous coal pieces and lignite coal pieces were placed on the microbial mats and observed for colonisation and degradation of coal.

3.3.4.1 Coal colonisation

After approximately two weeks of incubation, cell attachment to the coal pieces was observed for several isolates that were filamentous in morphology. Each colonised coal piece had hyphae tightly adhered surrounding and underneath the coal surface. The colonisation of coal pieces by these filamentous isolates continued for a further 2-3 weeks of incubation until the coal pieces were completely covered by a filamentous mat. A minority of filamentous isolates showed partial attachment after months of incubation. Certain isolates infiltrated small fractures of the coal pieces. With one exception, no colonisation was observed on polytetrafluoroethylene used as an inert control surface. Interestingly, certain isolates preferentially colonised untreated coal whilst others preferentially colonised nitric acid treated coal. Other isolates showed no preference or did not colonise coal at all (Figure 3.3 and Figure 3.4). The assay was repeated twice with identical results obtained. A summary of the proportion of isolates that colonised bituminous coal is illustrated in Figure 3.6.

None of the non-filamentous isolates were found to attach to coal (Figure 3.5). These isolates either showed hard-crusted or smooth and shiny morphology. The coal pieces were easily moved by light agitation, which confirmed the lack of attachment between the isolate and coal. In several cases, after several weeks of incubation, the untreated coal pieces were overgrown by filamentous organisms that were not present at the start of the assay. Thus, the growth was considered a contamination and did not contribute to the isolate’s colonisation activity on coal.

101

Figure 3.3 Representative images of preferential and non-preferential colonisation on coal by isolates. A and B, C and D, E and F and G and H represent images of before and after incubation of four isolates with untreated bituminous coal (U) and nitric-acid treated bituminous coal (T). A diverse colonisation preference of coal (squared outline) could be observed: A-B shows both untreated and treated coal colonisation; C-D untreated coal only and E-F treated coal only. G-H did not show any colonisation of coal.

102

Figure 3.4 Second test of coal colonisation and solubilisation assay. Selected isolates labelled as G5o, G9o and Y7e (images A, C and E, respectively) showed non-preferential (B), untreated (D) and treated (F) coal colonisation, respectively, after several weeks of incubation. The polytetrafluoroethylene control (P) was not colonised by the isolate throughout the incubation. Small white particles in F (dotted arrows) corresponded to water droplets on the inner side of the agar plate lid as a result of condensation. L= Lignite coal that was freshly added for a follow- up experiment (see Figure 3.5). The squared outline indicates complete coal colonisation.

103

Figure 3.5 Representative images of non-filamentous isolates showing no colonisation on coal after several weeks of incubation. The main morphologies of these isolates consisted of either hard-encrusting (A) or smooth and shiny (B). There were cases, as shown by the white arrows in C, of a filamentous organism surrounding the untreated coal. This was deemed as contamination and hence not considered as positive colonisation by the isolate. U= Untreated bituminous coal, T= Nitric-acid treated bituminous coal.

10%

Colonisation 40% 50% 40%

6% 4%

No colonisation Contamination UT U T

Figure 3.6 A summary of the proportion of isolates colonising coal. Half of the total isolates tested showed some form of colonisation, of which 40% colonised both untreated and treated coal (UT). A smaller percentage of the isolates colonised either the untreated coal (6%) (U) or treated coal (4%) (T) only. Almost half of the total isolates (40%) comprising of filamentous and non-filamentous morphology did not colonise coal. A small proportion (10%) of the isolates were deemed contaminated.

104 3.3.4.2 Bituminous coal degradation

Whilst a considerable proportion of isolates showed positive coal colonisation, most showed no apparent coal degradation. Coal colonisation occurred actively on average for one month but remained stagnant thereafter. Only one isolate showed direct degradation of bituminous coal after six months of incubation at 22 oC. This isolate was labelled G9o and identified as a Fusarium sp. The isolate originated from coal pieces in the coal washing tank of the abandoned oil shale mine (Table 3.1) and was selected from coal agar and silica in BIII medium with 6x trace elements. Based on the colonisation assay, G9o preferentially colonised untreated coal, whilst there was little or no attachment of its hyphae on the treated coal even after several repetitions. Likewise, coal degradation by the isolate was solely observed on the untreated coal. Whilst coal solubilisation was not observed, parts of the coal piece were found to be soft when the biomass was scraped from the coal surface and light pressure was applied (Figure 3.7).

In comparison, the positive control P. chrysosporium showed a more intense degradation of the untreated bituminous coal after approximately six months of incubation. This was shown by the formation of a dark brown halo in the agar directly underneath and surrounding the untreated coal (Figure 3.8). The dark brown halo was not observed on the treated coal. Interestingly, whilst the untreated bituminous coal was degraded, it was not completely colonised by P. chrysosporium. This was not the case with G9o, in which the untreated coal was both colonised and degraded.

105

Figure 3.7 Fusarium sp. G9o colonisation (A) and degradation (B) of the untreated coal (U) in comparison to the treated coal (T) that was hardly colonised or degraded. Softening of some parts of the untreated coal (arrow) was observed. Biomass was scraped from the untreated coal surface (B). The degradation was observed after approximately six months of incubation.

106
Figure 3.8 P. chrysosporium degradation of untreated bituminous coal after 6 months of incubation at 22 oC. A) Top view of the plate where untreated (U) and treated (T, square) coal pieces were placed on the fungal mat. Brown halo surrounding U could be seen. B) Mirror view from the bottom of the plate, where the brown halo could be observed more clearly as a result of degradation by P. chrysosporium.

3.3.4.3 Lignite coal solubilisation
Three isolates, Fusarium sp. G9o, Penicillium sp. Y7e and Penicillium sp. G5o, which showed strong colonisation of bituminous coal, were selected to test for lignite coal colonisation and solubilisation. After approximately 3 months of incubation at 22 oC in the dark, all these isolates showed positive lignite solubilisation based on direct observation of liquid formation on agar or its diffusion into the agar (Figures 3.9, 3.10 and 3.12). P. chrysosporium also showed similar results in a similar time (Figure 3.11). Whilst lignite degradation was preceded by full colonisation by the isolates, P. chrysosporium showed only partial colonisation.

107

Figure 3.9 Lignite coal (L, circle) solubilisation by isolate Penicillium sp. Y7e before (A) and after (B) 3 months of incubation. Liquid coal formation (arrow) could be observed. Based on the colonisation assay, Y7e preferred colonising treated coal (T) over untreated coal (U). Polytetrafluoroethylene control (P) was not colonised.

Figure 3.10 Lignite coal (L, circle) solubilisation by isolate Penicillium sp. G5o before (A) and after (B) 3 months of incubation. Lignite coal was added several weeks after colonisation of the bituminous coal was observed. A brown halo (arrow) could be seen in the agar directly underneath the colonized lignite, indicating solubilisation by the isolate. G5o was observed to colonise both untreated (U) and treated coal (T), but not the control surface (P).

108
Figure 3.11 Lignite coal (L) solubilisation of P. chrysosporium after approximately 3 months of incubation. The solubilisation of untreated (U) bituminous coal followed 3 months after. T= Treated bituminous coal and P= Polypropylene pipette tip as the control. The arrows highlight a lignite piece in (A), where part of it has been solubilised and diffused into the agar, forming a dark brown halo underneath the lignite piece (B).

109 Isolates G9o and G5o were used to compare the extent of colonisation and degradation between bituminous coal and lignite coal. In addition to the untreated and treated bituminous coals, two types of lignite coal were used, i.e. from Loy Yang, Australia and Kalimantan, Indonesia. The Kalimantan lignite differs from the Loy Yang lignite based on the elemental analysis. Figure 3.12 shows colonisation and degradation of all the described coal types by G9o over seven weeks. G9o colonised both lignite coals more profusely than bituminous coals based on the formation of biomass on the coal. After nearly 3 months of incubation, a brown halo could be observed from the bottom of the agar plate directly underneath both lignite coals, indicating coal solubilisation. Similarly, isolate G5o, colonised lignite coals more profusely and consequently showed increased solubilisation than for the untreated and treated bituminous coals (see Appendix V).

Figure 3.12 G9o colonisation and solubilisation test of four different types of coal: Untreated (U) and nitric acid treated (T) bituminous coals, and Loy Yang (L) and Kalimantan (K) lignite coals. In comparison with the initial incubation (A), the colonisation and solubilisation of the Loy Yang and Kalimantan lignites was quicker than the bituminous coals (images B and C). The control surface (P) was not colonised throughout incubation. Arrows refer to solubilised lignite (brown halo) diffused into the agar.

110 3.3.5 Oxidative enzyme assay
Based on their positive results in the solubilisation assay, isolates Fusarium sp. G9o, Penicillium sp. Y7e and Penicillium sp. G5o were also screened for the production of different oxidative enzymes, which have been shown to be associated with coal degradation (Hofrichter & Fritsche, 1997; Silva-Stenico et al., 2007; Wilmann & Fakoussa, 1997b). The assays included testing for laccase, MnP and phenolic complex degrading abilities by growing the isolates on agar plates with specific indicators of enzymatic activity. Both G9o and G5o showed positive results for laccase (plates with ABTS), manganese peroxidase

(plates with MnCl2) and phenolics degradation enzymes (plates with Reactive Violet 5 dye), although the duration varied from each isolate.

Table 3.4 summarises the results for all three isolates for each test. For plates containing an MnCl2 indicator, both G9o and G5o produced black pigments in the agar surrounding the colonies after approximately one day of incubation. As for the ABTS-containing plates, formation of dark green pigments appeared approximately after 2 weeks of incubation for G9o, while it took 3.5 weeks to see the final colour change for G5o. Both G9o and G5o decolourised the Reactive Violet 5 dye plate, however the duration also varied with each isolate, as G5o decolourised the agar plate in 4-5 days whilst G9o took twice as long.

Isolate Y7e did not produce any observable changes to the MnCl2, ABTS and Reactive Violet 5 agar plates. However, Y7e could decolourise the more recalcitrant Remazol Brilliant Blue dye in approximately 4 weeks, quicker than G9o and G5o, which took approximately 6 weeks to decolourise. Figures 3.13 – 3.16 show examples of changes in the agar plates containing the four indicators with the isolates.

111 Table 3.4 Oxidative enzyme assay results with isolates G9o, G5o and Y7e. Varying results were observed with each isolate and test. The time (in brackets) denotes the duration it took to first observe any colour change/loss in the agar plate.

Oxidative enzyme test indicator

Isolate MnCl2 ABTS Reactive Remazol Violet 5 Brilliant Blue R G9o Black pigments Dark green Decolourised Decolourised (1 day) pigments (10 days) (6 weeks) (2 weeks) G5o Black pigments Dark green Decolourised Decolourised (1 day) pigments (4-5 days) (6 weeks) (3.5 weeks) Y7e No change No change No change Decolourised (4 weeks)

Figure 3.13 Bottom view of agar plates containing MnCl2 grown with isolates G9o, G5o and Y7e. Black pigments, indicative of Mn2+ oxidation, were produced in the agar grown with isolates G9o and G5o. Isolate Y7e did not produce any black pigments, and the control (C) plate is a sterile

MnCl2-containing agar plate.
112

Figure 3.14 Bottom view of agar plates containing ABTS grown with isolates G9o, G5o and Y7e. After approximately 2 - 3.5 weeks of incubation, both G9o and G5o produced dark green pigments indicating oxidation of the ABTS by laccase production. Y7e did not produce these pigments. Control is a sterile ABTS-containing agar plate (C).

Figure 3.15 Reactive Violet 5 plate grown with G5o on days 0 (A), 2 (B) and 10 (C). By the end of the incubation time the isolate could decolourise the bright purple dye to light pink.

113
Figure 3.16 Bottom view of Remazol Brilliant Blue R plate grown with Y7e at the start (A), middle (B) and end (C) of incubation (approximately 4 weeks in duration). Unlike G9o and G5o,

Y7e did not cause positive changes to other plates (ABTS, MnCl2 and Reactive Violet 5). However, it could decolourise the Brilliant Blue dye (arrows showing the edge of the plate where decolourisation is more visible) 2 weeks quicker than the other isolates.

Based on the performance of the isolates on tests conducted above, isolate Fusarium sp. G9o was chosen for further characterisation in the subsequent chapter. Although isolates G5o and Y7e also showed interesting results, G9o was the only isolate that could directly degrade the more recalcitrant bituminous coal, apart from being able to solubilise lignite and degrade phenolic complexes. The unique ability of G9o to degrade bituminous coal in this study made it an ideal candidate for further investigations.

3.4 Discussion

Previous research has found that microorganisms, especially filamentous fungi isolated from the environment, have the ability to degrade coal (Cohen et al. 1987; Cohen and Gabriele 1982; Cohen and Wilson 1990; Fakoussa and Hofrichter 1999; Hofrichter et al. 1997b; Hofrichter and Fritsche 1997; Hofrichter and Fakoussa 2001; Laborda et al. 1997; Machnikowska et al. 2002; Wilmann and Fakoussa 1997a). Whilst intense research has been conducted in this field over a few decades, predominantly in Germany and the United States, isolation of coal degrading microorganisms from a native Australian environment is lacking despite the vast natural coal resources. Furthermore, the coal bed methane (CBM) industry in Australia is growing rapidly. Therefore, finding potential microbial candidates native to the Australian continent, which could be

114 used in biogenic CBM production, would be a useful tool to utilise the country’s natural resource. In addition, strict Australian quarantine laws do not permit deployment of foreign organisms in the environment, which stresses further on the need to isolate native coal-degrading microorganisms. This chapter describes work carried out to find microbes that have the ability to degrade hard coal that would have potential future use in the biogenic CBM industry.

Several stages were implemented for the isolation process. The first stage was the initial screening of various environmental samples from The Wolgan Valley, NSW on untreated bituminous coal agar and subsequently coal silica, both in various salts media. Isolates that were able to utilise coal as a sole carbon source (i.e., those that grew successfully on coal silica) were identified using molecular techniques (DNA extraction and ITS sequencing). The isolates were then screened for bituminous and lignite coal colonisation and degradation through direct observation on a solid medium. The best performing isolates were tested for the production of oxidative enzymes. The screening process culminated with the selection of a Fusarium species that displayed a desirable bituminous coal degradation capacity.

To eliminate microbes that were not utilising coal as a carbon source, the isolation assay was conducted with a carbon-free solidifying substance (i.e. silica). After approximately three weeks of incubation, around 13% of the total isolates from the coal agar screening showed visible growth on coal silica. The longer incubation time and substantial reduction in number of isolates indicated that the second screening on coal silica was more rigorous than that on coal agar. Hence it is more likely that the isolates that grew on coal silica were utilising coal as a sole carbon source. Previous research agrees that microbial growth on bituminous coal, especially without any external carbon source present (e.g. glucose), takes longer (i.e. three to six weeks) than when a simpler carbon source is introduced (Hofrichter and Fakoussa 2001). The lack of physical changes seen on the coal (e.g. colour bleaching or solubilisation) could have also reflected the slow utilisation of coal by the isolates. A previous screening of thousands of environmental samples on untreated bituminous coal revealed only 0.2% of isolates could degrade coal (Fakoussa 1988). The
115 similarity in results obtained in previous research and those obtained in the coal silica screening is suggestive of effective isolation of coal utilising microorganisms.

The subcultured isolates only grew on coal silica that contained salts and trace elements. There was no growth found on coal silica without these, despite the fact that there was growth in the original coal agar (without the minerals). This shows the importance of providing adequate minerals in order for coal utilisation to occur, which is in agreement with previous studies (Hofrichter & Fakoussa, 2001). Growth on coal silica mostly came from isolates that grew on coal with M9 minimal salts medium with 6x trace elements over other media. The preference of M9 medium (89%) over BIII medium (11%) by many isolates is likely due to the former containing five times more ammonium than the latter. Therefore, this may indicate that a higher amount of nitrogen was preferred by many coal-utilising fungi, which contradicts studies that indicate a low amount of nitrogen is more preferable for coal degradation to occur (Hofrichter & Fritsche, 1997; Wilmann & Fakoussa, 1997a). Nevertheless, certain fungi (basidiomycetes) have been found to degrade coal more efficiently in a high nitrogen environment, although their ligninolytic ability was suppressed (Hofrichter et al. 1997a).

Of the total number of isolates that grew on coal silica, approximately half were randomly chosen for molecular taxonomic identification prior to the coal colonisation and solubilisation assay (Table 3.3). Based on the identification data, a large proportion (91%) of Ascomycetous fungi were acquired. Whilst some Basidiomycetous fungi (i.e. white-rot fungi) are known for their coal degrading ability (e.g. Phanerochaete chrysosporium and Trametes versicolor), many Ascomycetous fungi (e.g. Penicillium citrinum and Trichoderma atroviride) also have such properties (Hofrichter and Fakoussa 2001). The most frequently identified genera in this study were Penicillium sp. (38%), followed by Paecilomyces sp. (14%), and Aspergillus sp. (6%). These three genera have previously been found to degrade coal, have been associated with degrading components found in coal (e.g. PAHs) or have been shown to produce

116 enzymes known to alter the physical and chemical properties of coal (Faison et al. 1991; Gesell et al. 2004; Hofrichter and Fakoussa 2001; Yuan et al. 2006).

Other isolates obtained that play a role in coal degradation, but are less frequently identified include: Fusarium oxysporum, a known coal solubiliser (Holker et al. 1999), Cladosporium sphaerospermum that has been found to degrade PAHs (Potin et al. 2004) and Acremonium sp., which produce an alkaline compound that solubilises coal (Quigley et al. 1989). The identification results are in line with previous research and thus give confidence that coal- degrading fungi have been isolated based on the multiple screenings on coal agar and silica in this study. Nevertheless, other isolates identified as Umbelopsis sp., Ustilago tritici, Torrubiella sp., Hypocreales sp., Lecanicillium lecanii and Malassezia restricta, have not previously been found to be associated with coal degradation, but are known instead for their human, animal, insect and plant pathogenicity or saprophyticity (Batts 1955; Bischoff and White 2004; Ro and Dawson 2005; Sugiyama et al. 2003). It is possible that these fungi could also degrade coal, perhaps through similar mechanisms of attack, due to resemblances in molecular structures with natural substrates (e.g. between lignin monomers in plants and humic acids in coal). P. chrysosporium, for example, could degrade lignin molecules, textile dyes and phenols, which all contain aromatic ring structures that could be cleaved using oxidative or hydrolytic enzyme systems (Baldrian 2004; Garcia et al. 2000; Spadaro et al. 1992; Tien and Kirk 1984).

Isolates that successfully grew on coal silica were subject to another screening based on direct observation of coal colonisation and degradation. As the isolates were initially screened for utilisation of coal as a sole carbon source, this assay utilised a richer medium with external carbon and energy source (i.e. glucose and peptone) in order to promote greater biomass to achieve a more effective coal colonisation and biodegradation (if any) within a limited time. Bituminous coal, untreated and nitric acid-treated, were initially used for this assay.

117 Within approximately two weeks of incubation, around half of the total isolates were observed to colonise coal, albeit to different degrees and preferences. Whilst a number of isolates managed to colonise a whole coal piece, there were other isolates that partially colonised coal, with the remaining isolates showing no attachment and colonisation on coal (Figures 3.3-3.5). All coal colonisers (full and partial) were filamentous in appearance and the non-colonisers exhibited rough and hard or smooth and shiny colonies similar to yeast or bacterial colony morphology respectively. The filamentous fungal isolates consisted of a complex hyphal network, which gave them an advantage for colonisation and thus biofilm formation ability in an aerial environment (Harding et al. 2009). The fungi not only extended their hyphae on the outer parts of coal, but also infiltrated through coal fractures. This colonising property seemed to be lacking in non-filamentous isolates, which presumably would form biofilms more efficiently in submerged environments than in aerial settings. It is unknown whether the filamentous fungal isolates in this study could perform as well in liquid as they would in aerial environments, but such occurrences by other filamentous fungi have been cited previously (Gamarra et al. 2010; Villena and Gutierrez-Correa 2007).

The colonising isolates showed active preferential attachment to coal. Except for one isolate, the colonisers were not observed to attach and colonise the control surface made up of polytetrafluoroethylene or polypropylene (Figure 3.4). Hence, this was evidence that coal was a substratum for active colonisation compared to passive colonisation where all surfaces are colonised irrespective of the surface type. Importantly, whilst all colonisers were filamentous isolates, there were isolates of filamentous morphology that demonstrated no form of adherence to coal. This suggests that not all the isolates obtained here had the same physiological properties to colonise coal. Nevertheless, this observation further supported the evidence of active attachment of the colonising filamentous isolates. These results, however, did not conclude that non-colonising filamentous, or non-filamentous isolates, would not participate in coal degradation. The coal degrading fungus P. chrysosporium used as positive control, for example, showed little or no colonisation to either the untreated or treated bituminous coal. This shows
118 versatility in nature to degrade and utilise coal through different degradation strategies.

The preferential attachment and colonisation of coal was further observed in the colonising isolates’ preference on different coal types. Interestingly, some of the isolates were observed to colonise either untreated or treated bituminous coal, or both (Figures 3.3, 3.4 and 3.6) whilst the majority had no preference. The cause of the observed preference is unknown, however, this could be indicative of the different mechanisms of coal degradation of the various isolates. Based on elemental analysis of the untreated and treated bituminous coal (Table 3.2), both coals differ substantially in oxygen and nitrogen content, meaning that there were differences in the functional groups that serve as attachment sites for coal degrading fungi (Hofrichter and Fakoussa 2001). Therefore, whilst the colonisers of both coal types would have mechanisms for attacking both untreated and treated coal, the preferential colonising isolates may only have the mechanism to degrade its preferred colonised coal.

Despite the observation of active colonisation on coal, most isolates did not show obvious bituminous coal degradation. For many isolates, the degree of activity was only seen on full or partial colonisation of coal that lasted for about one month of incubation and remained stagnant thereafter. However, one isolate, identified as Fusarium sp. (sample ID G9o), showed a form of coal degradation (Figure 3.7) after approximately six months of incubation. The scarcity of isolates that could degrade bituminous coal is somewhat expected given the recalcitrant and less oxidised nature of this type of coal. However, it was unexpected not to find more isolates capable of degradation of artificially oxidised bituminous coal (i.e. through nitric acid, hydrogen peroxide or autoclaving) as has been observed previously (Achi and Emeruwa 1993; Igbinigie et al. 2008; Monistrol and Laborda 1994). Nevertheless, this limitation might not be due to the nature of bituminous coals per se. The conditions to which all isolates were subjected (e.g. medium, temperature) were the same across all isolates and thus may not have been optimal for many fungi to degrade coal.

119 The isolate Fusarium sp. G9o showed partial degradation of untreated bituminous coal. This was observed through the softening of the edge of the coal when light pressure was applied. Softening of the coal only occurred in the untreated coal. A similar observation was made by Monistrol and Laborda (1994), where some fungi could soften the bituminous coal pieces after completely engulfing the coal with mycelia. No solubilisation of bituminous coal occurred. In comparison with the positive control P. chrysosporium, untreated coal was also preferred for degradation over treated coal (Figure 3.8). However, in the latter no softening of coal was observed; instead, leaching of coal soluble substances was evident from the colouration of the agar directly underneath the coal piece.

The difference in the coal modification capacity between these two fungi indicate variations in their coal degradation mechanism despite a shared preference to degrade untreated coal. Based on the oxidative enzyme analysis, G9o could produce MnP and laccase, which are also found in P. chrysosporium from previous studies (Godfrey et al. 1996; Srinivasan et al. 1995). A Fusarium species called F. oxysporum isolated by Holker et al. (1995) was also found to contain a gene encoding a ligninase H8 homologous to that found in P. chrysosporium (Monkemann et al. 1996). The properties shared by both fungi might be useful for bituminous coal degradation, although it is known that the enzymes have also been associated in the breakdown of other coal types and humic acid substances.

A difference between G9o and P. chrysosporium worth noting is that whilst both fungi preferred degrading untreated bituminous coal, only G9o showed complete coal colonisation prior to degradation. P. chrysosporium did attach to coal, however the area covered was only that in contact with the medium on the agar plate. This shows that there was a varying degree of coal colonisation among coal-degrading fungi and full colonisation or engulfment of coal was not necessary for coal degradation to occur.

It is uncertain as to what drives colonisation (or the lack of it) by coal degrading fungi prior to degradation. In light of this, Abbasnezhad et al. (2011) outlined
120 that cell attachment and colonisation to hydrocarbons (specifically that of liquid) is not necessarily a prerequisite to their degradation. However, adherence becomes more prominent and advantageous when the hydrocarbon is low in bioavailability and the microorganism lacks certain metabolites (e.g. surfactants, enzymes) that would enhance the degradation further. It is known that P. chrysosporium, a white-rot fungus, contains extracellular enzymes that are able to degrade not only coal but also a range of pollutants including polychlorinated biphenyls (PCBs) and carbon tetrachloride (Cameron et al. 2000; Eaton 1985; Ralph and Catcheside 1994). Hence, whilst the isolates obtained in this study may possess similar properties, those of P. chrysosporium are different in such a way that they warrant the fungus to achieve effective degradation without full colonisation. Whereas for Fusarium sp. G9o, a larger cell-to-coal surface area of contact is probably necessary for more efficient coal biodegradation.

As the bituminous coal assay proved to be difficult for degradation, lignite coal was consequently tested to observe if the isolates could degrade a lower-rank coal. Isolates that showed different preferential colonisation of bituminous coal (i.e. both untreated and treated, untreated only, and treated only) were selected and incubated with lignite. All isolates, including the positive control, showed lignite solubilisation within three months of incubation (Figure 3.9-3.12). This showed the ability of the isolates to degrade coal despite their inability to attack the bituminous coal. It is possible that the colonising isolates would degrade bituminous coal if more time was given and/or incubation conditions were modified. However these aspects were not central to this study and therefore were not investigated further.

The quicker and more obvious results in lignite degradation is evidence that lignite coal is more easily degraded than bituminous coal, which is in line with many previous studies (Hofrichter and Fritsche 1997; Laborda et al. 1997; Monistrol and Laborda 1994; Silva-Stenico et al. 2007; Wondrack et al. 1989; Yuan et al. 2006). The preference for lignite over bituminous coal for degradation is most likely due to its chemical structure, consisting of more oxygen-linked moieties (e.g. carboxylic and carbonyl bonds) that act as sites for microbial attack (Hofrichter and Fakoussa 2001). Based on the elemental ratios
121 (Table 3.2), the lignites used had a much lower C:O ratio compared to bituminous coal, which makes it a more oxidised coal and more susceptible to microbial degradation. Furthermore, the lignites have a higher C:N ratio than the higher-ranked coal, which indicates that the isolates may have preferred the type of organic matter contained in lignite to bituminous coal for degradation. Nonetheless, this generalisation cannot be applied to all microbes, since it has been previously shown that certain fungi could more effectively degrade bituminous coal than lower-ranked coal (Igbinigie et al. 2008). Thus, coal degradability is not only dependent on the coal structure but also the presence of appropriate mechanisms to degrade different coal types.

Lignite solubilisation was preceded by full colonisation, with the exception of the positive control P. chrysosporium. This was demonstrated further by isolate G9o where the bituminous coal pieces were simultaneously compared against two types of lignite coal for colonisation and degradation (Figure 3.16). Indeed, both lignites (Loy Yang and Kalimantan) were more heavily colonised than the bituminous coals and resulted in solubilisation of the lower-ranked coal. This was also shown by Penicillium sp. G5o (Appendix V). The results demonstrate a relationship between colonisation and degradation of coal by these isolates, whereby a substrate that is more susceptible to degradation (lignite) is colonised faster than the less amenable one (bituminous coal).

The isolates chosen for lignite solubilisation were also tested further for oxidative enzyme production. The production of oxidative enzymes such as manganese peroxidase (MnP), lignin peroxidase (LiP) and laccase have been associated with humic acid and coal degradation (Granit et al. 2007; Hofrichter et al. 1997b; Hofrichter and Fritsche 1997; Wilmann and Fakoussa 1997b).

Based on the results on MnCl2, ABTS and reactive dye oxidation (Table 3.4), the isolates differed in the time taken to oxidise the substrates. Both G9o and

G5o had the ability to oxidise MnCl2 and ABTS (Figures 3.12-3.13), which indicated the presence of MnP and laccase enzymes, respectively. The two isolates could also decolourise the dyes, indicating the ability to cleave phenolic complexes, although G5o was more efficient in degrading the Reactive Violet 5 than G9o (Figure 3.14). Isolate Y7e however, could only decolourise the
122 Remazol Brilliant Blue R dye but was quicker than G5o and Y7e (Figure 3.16). The results imply that the isolates produced oxidative enzymes that can be potentially applied for coal degradation, although their mechanisms to degrade coal differ from one another. However, the lack of production of these enzymes does not necessarily mean a lack of coal degrading capacity, as there are other mechanisms such as alkaline substances and chelators that fungi may use to degrade coal (Hofrichter and Fakoussa 2001).

In this study, a filamentous bituminous coal-degrading fungi Fusarium sp. G9o was isolated from an Australian environment. The isolate also showed lignite solubilisation and oxidative enzyme production. Furthermore, preferential coal attachment was observed, which was shown to link to coal degradation. The positive correlation observed in this study between colonisation and degradation, although not applied to all coal-degrading fungi, is an important fundamental aspect worthy of further investigation.

The successful isolation provides a useful application to the microbial transformation of coal to methane. Further characterisation of this isolate is necessary prior to its application to the wider field. Mechanisms of coal degradation, and more fundamentally, the cell attachment and colonisation of coal that precedes coal degradation, are addressed in Chapter 4.

123 4 Characterisation of the fungus Fusarium oxysporum G9o on the attachment and degradation of coal

4.1 Introduction

In the previous chapter, the isolation of coal degrading microorganisms revealed that the fungus Fusarium sp. G9o was able to degrade high-ranked bituminous coal. It was the only isolate that showed this attribute, and therefore was chosen as a model organism for studying the mechanisms of fungal attachment on coal and coal biodegradation.

Ubiquitous in nature, F. oxysporum is particularly common in soil and the rhizosphere (Fravel et al. 2002; Rodriguez-Galvez and Mendgen 1995). Plant pathogenic and non-pathogenic strains have been isolated (Gordon and Martyn 1997; Macia-Vicente et al. 2008; Vu et al. 2006). Furthermore, this fungus was found to degrade fractions of wheat straw, specifically those of hemicellulosic, cellulosic and lignin origins (Rodriguez et al. 1996), and also lignin-related phenyl-propanoid acids as a sole carbon source (Falcon et al. 1995). Interestingly, under low oxygen conditions, F. oxysporum has been found to attack low molecular weight polyaromatic hydrocarbons (PAHs), where ligninolytic enzyme production was observed (Silva et al. 2009). These abilities and the evidence presented in the previous chapter indicate that F. oxysporum is a good candidate for aerobic coal degradation, particularly at the initial stages of microbial coal fragmentation.

The ability of F. oxysporum to degrade coal has been documented. For example, Holker et al. (1995) isolated an F. oxysporum strain from an open cast coal-mining area in Cologne, Germany, that showed solubilisation of hydrogen peroxide-treated lignite occurring within 24 hours of incubation. The authors suggested several possible mechanisms by which F. oxysporum degrades coal (Holker et al. 1999). Further, ligninolytic enzyme production by F. oxysporum

124 that paralleled those found in P. chrysosporium, a well-known coal-degrading fungus, has also been demonstrated (Monkemann et al. 1996). These studies provide essential information to understand the relationship between F. oxysporum and coal, however, the amount of literature concerning this subject remains scarce.

To date, there is no evidence in the literature that F. oxysporum can degrade a higher-ranked (i.e. bituminous) coal, and the use of analytical chemistry methods to give insights on its coal-degrading mechanism has not been applied. More fundamentally, cell attachment to coal has not been investigated thoroughly for any coal-degrading fungus. Thus, this study aims to extend the existing research on F. oxysporum and coal degradation, with an emphasis on fungal attachment on coal, to better understand its coal-colonising and degrading properties and potential biotechnological applications.


4.2 Material and Methods
4.2.1 Molecular investigation on species identification of Fusarium sp. G9o

The taxonomic identity of isolate G9o was investigated using primer sets targeting the 18S, internal transcribed spacer (ITS) and the large subunit (LSU) regions. Primer sets NS1/FR1 (White et al. 1990), ITS1F/ITS4 (White et al. 1990) and LR0R/LR16 (Yoo and Eom 2012) were used to target the DNA regions respectively. In the previous chapter, the ITS3/ITS4 primer pair was used to target the ITS2 region of G9o after being unable to amplify the whole ITS region with the ITS1/ITS4 primer pair. Despite the success in sequencing the ITS2 region, the sequence length was inadequate to obtain the fungal species identity. Therefore further modification of the ITS PCR protocol, in addition to targeting other regions (18S and LSU) were applied.

125 The G9o DNA was extracted according to the method outlined in Section 3.2.8.1. For primer sequences, refer to Appendix II. All PCR procedures (except for the thermal cycling profile) were conducted as outlined in Section 3.2.8.2. The thermal cycling PCR profile and reference for the amplification is as follows:

• NS1/FR1 (White et al. 1990): Initial denaturation at 95 oC for 3 min; 35 cycles of denaturation at 94 oC for 30 s, annealing at 57 oC for 30 s, extension at 72 oC for 105 s, followed by a final extension at 72 oC for 2 min. • ITS1F/ITS4 (Granit et al. 2007): Initial denaturation at 94 oC for 4 min; 32 cycles of denaturation at 94 oC for 45 s, annealing at 62 oC for 30 s, extension at 72 oC for 45 s, followed by a final extension at 72 oC for 7 min. • LR0R/LR16 (Yoo and Eom 2012): Initial denaturation at 94 oC for 3 min; 36 cycles of denaturation at 94 oC for 30 s, annealing at 54 oC for 30 s, extension at 72 oC for 1 min, followed by a final extension at 72 oC for 10 min.

Amplified products were further sequenced and identified according to procedures outlined in Section 3.2.8.3 differing only by the forward primer used. The ITS fragment was compared phylogenetically to sequences of known species in the GenBank database of the National Center for Biotechnology Information (NCBI) by using the BLASTn program. Phylogenetic trees were analysed using the Phylogeny.fr program (Dereeper et al. 2008), which includes alignments using ClustalW (Thompson et al. 1994), Gblocks curation (to eliminate poor alignments) (Castresana 2000) and tree construction using the BioNJ method (Gascuel 1997). The phylogenetic tree was visualised using the tree rendering program TreeDyn. Comparable sequences for the ITS region were selected from initial BLASTn searches and GenBank database on several known coal degrading fungi.

126 4.2.2 Direct observation and Scanning Electron Microscopy (SEM) of bituminous coal colonised by F. oxysporum G9o
Untreated bituminous coal pieces colonised by G9o from previous coal degradation analyses (Figure 3.12) were further investigated based on direct observation and SEM. The coal pieces were gently rinsed in phosphate buffered saline (PBS) three times before following a series of SEM preparation procedures involving ethanol dehydration, hexamethyldisilizane drying and gold sputtering outlined in Section 2.2.6.
4.2.3 Contact angle measurements and surface thermodynamics of adhesion of F. oxysporum G9o on coal
Contact angle measurements of water, diiodomethane and formamide on G9o hyphal mat surfaces were conducted to measure G9o surface hydrophobicity. The surface free energy and free energy of adhesion between G9o and coal surfaces were calculated (see Section 2.2.8). The fungal surface was prepared in accordance to a method developed by Chau et al. (2009) with modifications. The fungal surface was prepared by growing it on a thin layer of agar on a microscope glass slide.

To prepare the agar, a 25 x 75 mm glass microscope slide sterilised with 70% w/v ethanol, was spread evenly with approximately 2 ml of sterile Sabouraud media using a micropipette. The agar slide was kept in a sterile 90 mm petri dish and left to cool before use.

A liquid suspension of G9o was prepared by scraping a 1-week old grown fungal mat using a 10-μL inoculating loop and transferring it into 100 μL of phosphate buffered saline (PBS) in a 1.5 ml Eppendorf tube. The tube was vortexed for 5-7 seconds before inoculating approximately 25 μL on the centre of the agar using a micropipette. This was to ensure that the fungus grew from a single point and spread to both sides of the agar. The inoculum was left to dry and incubated in the dark at 22 oC. The slide was observed every 2-3 days for growth during 2 weeks. Locations of growth for 2 weeks, 1 week and 2 days of

127 the fungi on the agar were noted and used to perform the contact angle measurements. Approximately 10 μL of each liquid was used to measure the contact angle using the static sessile drop method (Section 2.2.8). Three independent replicates were used, and the measurements were conducted twice for each replicate.
4.2.4 Fourier Transform Infrared (FTIR) analysis of coal incubated with F. oxysporum G9o
Coal pieces that were analysed for colonisation and degradation based on direct observation (Figure 3.12) were also examined for FTIR. All four coal types, i.e. untreated bituminous, treated bituminous, Loy Yang lignite and Kalimantan lignite that were exposed to the fungus were used in this analysis. The coal pieces were dried in a 60 oC oven overnight to remove any moisture and were subsequently ground with pestle and mortar to form a powder. Each powdered coal sample was then mixed with oven-dried potassium bromide (1:100) before being pressed with a 13 mm PerkinElmer evacuable pellet die (Beaconsfield, UK) and Specac 10 ton hydraulic press (Kent, UK). The coal samples were individually prepared to prevent cross contamination. Unmodified coal samples coming from the same batch as the G9o-exposed coal were also included in the preparation (including drying) and analysis.

The pellets were analysed using a Perkin Elmer Spectrum100 FTIR spectrometer fitted with a deuterated triglycine sulfate (DTGS) detector (PerkinElmer, Beaconsfield, UK). The infrared (IR) spectra were processed using the PerkinElmer Spectrum 6 software. Stackplots of the average of 64 spectra were prepared. The data was corrected for baselines and the effect of atmosphere.
4.2.5 Analysis of F. oxysporum G9o coal degradation in liquid culture

G9o biomass was pre-grown in a Sabouraud medium (Appendix I) at 28 oC in the dark for 48 hours (exponential phase) at 70 rpm on a rotary shaker (AVM Laboratory Service Pty. Ltd., Ryde, Australia). The biomass was then washed
128 three times in a glutamate medium without glucose (Appendix I) by centrifugation at 2,000 x g for 15 min. The washed biomass served as the inoculum for the assay. Approximately 2 ml (64 mg of dry weight) of the biomass was used to inoculate 80 ml of glutamate medium with 1% w/v glucose and 0.25% w/v of powdered coal in a 250 ml conical flask. The content was mixed by swirling the flask five times prior to sampling. Two coal types were used: untreated bituminous coal and Loy Yang lignite coal, which were chosen as representative of the high and low coal rank, respectively. Non-inoculated controls for each coal type were also included. Three independent replicates of each test were utilised. The cultures and controls were incubated in the dark at 28 oC for 37 days with no agitation.

When sampling, approximately 4 ml of culture was transferred into a 5 ml tube and centrifuged at 2,500 x g for 10 min to separate the fungal biomass and/or unsolubilised coal from the supernatant. The supernatant was then filtered through a 0.45 μm Millipore filter (Merck Millipore, USA). Approximately 700 μL of the filtered supernatant was transferred into a 1 ml spectrophotometric cuvette and measured at 300, 350, 400, 450 and 600 nm using a SmartSpec Plus spectrophotometer (Bio-Rad Laboratories Inc., California, USA). The glutamate medium served as a blank for the analysis. Samplings were conducted every 2 days (Day 0 included) for the first 2 weeks and subsequently once a week on average.
4.2.5 Extracellular oxidative enzyme activity analysis of F. oxysporum G9o

In Chapter 3 (Section 3.3.5), isolate G9o was reported to produce enzymes oxidising phenolic complexes (i.e. MnCl2, ABTS and azo dyes). In this chapter, the activity of manganese peroxidase (MnP), lignin peroxidase (LiP) and laccase in G9o were tested in the presence of coal. Supernatants of the cultures used to test coal degradation in liquid (Section 4.2.7) were used and analysed every 2 days over 2 weeks. All enzyme tests were conducted using a Cary 100 Bio UV-Visibile Spectrophotometer (Agilent Technologies, California,

129 USA). The Cary WinUV software under Enzyme Kinetics was used to analyse the data in real-time.

Manganese peroxidase (MnP) activity was determined using 3-methyl-2- benzothiazolinone hydrazone (MBTH) and 3-(dimethylamino)benzoic acid (DMAB) as substrates for the enzyme (Castillo et al. 1994). The reaction mixture contained 70 μM MBTH, 1 mM DMAB (in absolute ethanol), 300 μM manganese sulfate (MnSO4) and 50 μM hydrogen peroxide (H202) in 125 mM sodium tartrate buffer. The presence of manganese peroxidase was detected by the oxidation of MBTH, which then reacted with DMAB to produce an azo dye with a blue coloration. Absorbance was measured at 590 nm (extinction -1 -1 coefficient, ε590= 53,000 M cm ). This reaction is based on the oxidative 2+ coupling of MBTH and DMAB in the presence of H202, Mn and MnP.

Lignin peroxidase (LiP) activity was assayed using veratryl alcohol as substrate (Tien and Kirk 1984). The reaction mixture contained 2 mM veratryl alcohol

(distilled), and 0.5 mM H202 in 50 mM sodium tartrate buffer pH 3. The reaction is based on the oxidation of veratryl alcohol to veratryl aldehyde, which absorbs -1 -1 light at 310 nm (ε310= 9300 M cm ).

Laccase activity was measured using 2,2’-azinobis(3-ethylbenzothazoline-6- sulfonic acid) (ABTS) as substrate (Giardina et al. 2000). The reaction mixture contained 2.5 mM ABTS in 100 mM sodium citrate buffer pH 3. The substrate

ABTS is oxidised and gives a dark green color measured at 420 nm (ε420= 36,000 M-1 cm-1).

All enzyme activities were measured at 25 °C and expressed in International Units (IU), where one unit of enzyme activity is defined as the amount of enzyme that oxidises one µmole of substrate per minute.

130 4.2.6 Statistical analyses
The student t-test (one-tailed distribution, paired) was employed using Microsoft Excel 2011 to determine statistical significance in the difference between two samples of interest.

4.3 Results
4.3.1 Species identification of Fusarium sp. through 18S, ITS and LSU sequencing

A series of molecular identification tests on Fusarium sp. G9o targeted the 18S, ITS and LSU regions of the fungal rDNA. Only the ITS and LSU could be successfully amplified. The NS1/FR1 primers for the 18S region did not generate a product despite extensive PCR optimisation. Results from BLASTn queries against the GenBank database revealed that sequences from both the ITS and LSU of isolate G9o were closest to those of Fusarium oxysporum (99% match identity, 0.0 Expect (E)-value).

A phylogenetic tree using the ITS region of F. oxysporum G9o was constructed (Figure 4.1). F. oxysporum G9o were analysed against known sequences of F. oxysporum strains and several known coal degrading fungi. It is shown that F. oxysporum G9o is almost identical to a list of known F. oxysporum strains, namely F. oxysporum ES12 (accession no. DQ489299), F. oxysporum A4 (accession no. JF779674) and F. oxysporum forma specialis (f. sp.) ciceris (accession no. JN400697). F. oxysporum G9o was also compared against a list of known coal-degrading fungi from Penicillium sp. (Penicillium sp 8a accession no. AY391833 and P. chrysogenum ES16 accession no. DQ438910), Phanerochaete sp. (Phanerochaete Y6 accession no. DQ438910 and P. chrysosporium 776 accesion no. AY854086) and Trametes sp (Trametes sp. M23 accession no. DQ408582). Among these genera, F. oxysporum G9o is found to be closest to Penicillium sp.

131 98

100

100

99 100

Figure 4.1 Phylogenetic tree generated by BioNJ method based on ITS rDNA sequences of Fusarium oxysporum G9o and sequences from GenBank. The strain name and accession number (except for G9o) were included in the figure. Bootstrap values of > 95% from a sample of 1000 replicates are shown in each branch.

4.3.2 Direct observation and Scanning Electron Microscopy (SEM) of bituminous coal colonised by F. oxysporum G9o
Based on an earlier colonisation study (Figure 3.12),
a unique observation was made on the untreated bituminous coal colonised by G9o (Figure 4.2). At the start of incubation, the coal pieces remained intact (since fragile coal pieces would break easily during coal preparation). However, after approximately a month, fine fractures on the coal pieces were observed. The fungal colonisation at the edge the fine fracture lines was also more apparent. After 3 months of incubation with G9o, the initially intact coal pieces were physically fragmented.

Day 25

Day 52 Figure 4.2 Untreated bituminous coal pieces incubated with G9o on Day 25 and 52. The initially intact coal pieces were colonised by the fungus especially on the fine lines (full arrows) but eventually were physically fragmented (dashed arrows) over time.

132 The untreated bituminous coal pieces incubated with G9o were imaged using SEM to observe the fungal cell morphology and its mechanism of coal colonisation. The fungal morphology consisted of extensive and branching mycelia, which intertwined and formed a mat covering the coal surface (Figure 4.3 A). Coccobacillus-shaped cells of approximately 2 x 5 μm could also be observed attaching mostly on the tip of the hyphae and on the coal surface (Figure 4.3 B and 4.3 C respectively). These entities were most likely to be microconidia of the fungus, as confirmed in a previous SEM study of F. oxysporum (Alves Mde and Pozza 2009). In addition to these cells, inter-joining globose-shaped cells with punctate surfaces of approximately 9 x 11 μm were observed along the fungal hyphae (Figure 4.3 D). These cells were characteristic of chlamydospores of the fungus (Ohara and Tsuge 2004).

A B

C D

Figure 4.3 Representative SEM micrographs of F. oxysporum G9o cell morphology. A) complex hyphal network intertwining across the coal surface. B) Cocco-bacillus shaped cells (full arrow) attaching to the hyphal tips and C) on the coal surface. D) Inter-joining and globose-shaped cells (dashed arrows) were observed among the hyphae.

133
Furthermore, it was obvious from the micrographs that the coal surface was not only covered wholly by the hyphae, but also by another layer of fungal cells that penetrated through the fine coal cracks (Figure 4.4). Some parts of the coal were also observed to be loosely attached to the main coal piece but were still supported by the fungal hyphae.

C F F C C F

C C F F

C

C

C

F F

C

Figure 4.4 Representative micrographs of coal surfaces (C) exposed to G9o colonisation. Some of the fungal biomass (F) could be seen penetrating through the fine coal cracks. There were also some loosely attached coal pieces (arrows), where the fungal network could be seen underneath.
134
4.3.3 Contact angle measurements and surface thermodynamics of adhesion between F. oxysporum G9o and coal
The contact angles of water, formamide and diiodomethane were measured on G9o surfaces to determine the fungal surface hydrophobicity and free energy of adhesion on coal. The fungus was grown on agar slides for 2 weeks and its surface measured on Days 2, 7 and 14.

Figure 4.5 shows the water contact angle of the fungal surface over the incubation period. The water contact angles of the untreated bituminous coal and P. fluorescens were also included for comparison. The contact angles increased gradually as the fungus aged from Days 2, 7 and 14, starting from 63o up to 107o, which was close to the contact angle of untreated bituminous coal (approximately 120o). The difference in contact angles between Days 2 and 14 was significant (P<0.0001).
140

120 ) W

100 θ

80

60

40 Water contact angle ( Water 20

0 Day 2 Day 7 Day 14 Untreated P. fluorescens bituminous coal

Figure 4.5 Water contact angle of G9o surface on Days 2, 7 and 14 in comparison to those of untreated bituminous coal and P. fluorescens. As the days increase, the water contact angle of G9o also increases, approaching close to that of the untreated bituminous coal on Day 14. Error bars represent standard deviation of at least three independent replicates.

135 Table 4.1 summarises the contact angles of all liquids (water, formamide and diiodomethane) and surface free energy components of G9o on days 2, 7 and 14. The values for bituminous coal and lignite from an earlier study (Chapter 2) were also included for comparison. On average, the total surface free energy (γTOT) of G9o at all stages was much lower in comparison to that of coal and P. fluorescens (Table 2.6). However, as the fungus aged, its electron accepting capacity (γ +) increased, whilst the reverse was true for its electron donating capacity (γ -). A remarkably high γ - was observed for G9o on day 2, which was higher by 5 and 35 times that of days 7 and 14, respectively.

Based on the contact angle and surface free energy data, the free energy of adhesion (∆GAdh) between G9o and coal was calculated (Table 4.2). In general, the ∆GAdh was negative between G9o and coal. The least negative ∆GAdh values 2 were between G9o and untreated bituminous coal (on average -27 mJ/m ). The

∆GAdh between G9o and treated bituminous coal doubled with an average of -46 mJ/m2 across all days. These values were also similar against lignite for G9o on days 7 and 14. Interestingly, the ∆GAdh on day 2 between G9o and lignite was extremely low (-82 mJ/m2) but increased as the fungus grew. This trend could be coupled to the electron donating capacity γ - of G9o. As lignite had the highest electron accepting capacity γ + among all coal types, this contributed to its strong interaction with G9o on day 2, which had the highest opposing electron charge.

136

W F Table 4.1 Contact angle and calculated surface energy components of G9o on days 2, 7 and 14. The contact angles of water (θ ), formamide (θ ) and Di TOT diiodomethane (θ ) were measured on G9o surface. These values were used to calculate the total surface energy (γ ) and its components: Lifshitz-van der LW + - AB Waals (γ ) and electron accepting (γ ) and donating (γ ) capacities that make up the acid-base (γ ) interaction. The values for coal from an earlier study were also included for comparison. ‘Bituminous’ refers to untreated bituminous coal and ‘Treated bituminous’ refers to nitric-acid treated bituminous coal. ‘±’ - refers to standard deviation of three independent replicates. The bolded value highlights the relatively high electron donating capacity (γ ) of G9o on day 2.

-1 Surface Contact angle θ (o) Surface energy components (mN m )

θ W θ F θ Di γ LW γ + γ - γ TOT

G9o Day 2 63.1 ± 5 107.8 ± 4.3 106.5 ± 3.8 6.5 0 96.2 6.5

G9o Day 7 88 ± 8.5 96.3 ± 10.6 112.8 ± 2.3 4.8 0.7 21 12.7

G9o Day 14 106.7 ± 4.6 97 ± 5.6 107.3 ± 3.3 6.3 1.4 2.7 10.1

Bituminous 118.2 ± 3.1 101.7 ± 6.8 57.9 ± 9.6 29.8 0 0.4 29.8

Treated 24.7 ± 4.8 18.6 ± 5.5 20.3 ± 6.8 48.2 0.1 48.6 53.3

bituminous

Lignite 85.4 ± 3.6 16.2 ± 3.1 16 ± 3.5 48.9 5.5 0 48.9

137 Table 4.2 Total free energy of adhesion (∆GAdh) for F. oxysporum G9o against untreated bituminous, nitric acid-treated bituminous and lignite coals across days 2, 7 and 14 of incubation. ‘Bituminous’ refers to untreated bituminous coal and ‘Treated bituminous’ refers to nitric acid-treated bituminous coal. The bolded value highlights the relatively low ∆GAdh between - lignite and G9o on day 2, which corresponds to the latter high electron donating capacity (γ ) in Table 4.1.

2 Coal surface Total free energy of adhesion ∆GAdh (mJ/m )

F. oxysporum G9o cell surface

Day 2 Day 7 Day 14

Bituminous -28 -25 -29

Treated -42 -45 -52

bituminous

Lignite -82 -52 -43

4.3.4 Fourier Transform Infrared (FTIR) analysis of different coal types colonised and/or degraded by F. oxysporum G9o
Figures 4.6 – 4.7 correspond to the representative FTIR spectra of coal modified by G9o. The major differences were observed in the spectral range between 450-1800 cm-1. Each G9o-modified coal spectrum (‘b’) was compared against its unmodified coal spectrum (‘a’).
A common peak of interest across the coal samples was at 1262 cm-1, which corresponds to a C-O vibration (Painter et al. 1980). Changes in this peak’s relative intensity was observed against that of another peak at approximately 1640 cm-1, which corresponds to the C=C ring deformation mode from the

138 aromatic material in coal. The 1640 cm-1 peak was present in every spectrum and not expected to change in intensity due to a large proportion of aromatic material found in all coals. The relative intensity of the 1262 cm-1 peak in the G9o-modified untreated coal spectrum was approximately 7-fold higher than its unmodified version. In contrast, very little increase (1.3-fold) of the 1262 cm-1 relative peak intensity was observed in the G9o-modified treated coal against the unmodified control. Interestingly, this peak was observed in the unmodified lignites but was substantially reduced in intensity (approximately 7-fold) after exposure to G9o. This shows that the untreated coal and both lignite spectra had the most substantial changes in peak intensity at 1262 cm-1 compared to the treated coal after fungal exposure.

For both untreated and treated bituminous coals (Figure 4.6), similarities in peak reduction (relative intensity against the 1640 cm-1 peak) were observed in the G9o-modified coal spectra. Peaks positioned around 1117, 1013, 913, 537/8 and 473/4 cm-1 were either substantially reduced or disappeared after G9o modification. In particular, peaks at 538 cm-1 and 474 cm-1 were more intense in the modified treated coal than the untreated coal. However these peaks were likely non-carbon bonds e.g. silicon dioxide containing bonds (i.e. 474, 538, 1013 cm-1 assignment to Si-O) (Gomez-Serrano et al. 2003) and minerals such as kaolin (913 cm-1 assignment to Al-OH) (Miller et al. 2012) and -1 anhydrite (1117 cm assignment to CaSO4) (Estep et al. 1968). There were reduced peaks unique to the G9o-modified coal spectrum that relate to carbon or alcohol-containing groups not observed in the modified treated coal spectrum. These were 749 cm-1 and 1436 cm-1, which can be assigned to aromatic C-H plane deformation vibration (Painter and Rhoads 1981) and C=C, C-O or OH bonds in organic matter (Gomez-Serrano et al. 2003), respectively. These examples show that changes in carbon-related peaks were more evident in the modified untreated coal than the modified treated coal spectra.

For both Loy Yang and Kalimantan lignite (Figure 4.7), apart from the loss of the 1262 cm-1 peak in the G9o-modified coal, the only other similarity both coals shared was the reduction of the 1714 cm-1 peak, which corresponds to a C=O vibration (Fredericks et al. 1983; Painter et al. 1980). This peak was also
139 observed in the treated bituminous coal spectrum, however it did not show any obvious decrease in relative intensity. The other substantial change of peak intensity was at 801 cm-1 in the Loy Yang lignite spectrum, which can be assigned to kaolin (Si-O) (Gomez-Serrano et al. 2003; Miller et al. 2012).

A a

1640

b %T

1640

-1 cm

B a

1640

% b

1640

-1 cm

Figure 4.6 Representative FTIR spectra of A) Untreated bituminous and B) nitric acid-treated bituminous coal modified by G9o (b) against the unmodified control (a). The full arrows correspond to peaks that were present in the unmodified coal but reduced or disappeared after incubation with G9o, whilst the dashed arrows show peaks present in the G9o modified coal but not present or in less intensity in the original coal.
140 a A

1640 b %T

818

1640

-1
cm

a B

b %T 1640 1403

1640

-1 cm
Figure 4.7 Representative FTIR spectra of A) Loy Yang lignite and B) Kalimantan lignite modified by G9o (b) against the unmodified coal (a). The full arrows correspond to peaks that were present in the unmodified coal but reduced or disappeared after incubation with G9o.

141 4.3.5 Coal degradation by F. oxysporum G9o in submerged conditions
F. oxysporum G9o was tested for growth and degradation of coal in liquid culture. The fungus was incubated in glutamate medium with powdered untreated bituminous coal and Loy Yang lignite for 37 days at 28 oC. Filtrates of culture supernatants, including those of the G9o-free and coal-free controls, were measured for absorbance at specific wavelengths. The filtrates were also used for the extracellular enzymes assay (section 4.3.6).

After a lag period of 2-3 days, G9o was observed to grow fully submerged in the glutamate medium with a small amount of biomass on the medium-air interface. The growth looked like a mixture of filamentous biomass and gelatinous material, which increased in proportion throughout the incubation period. Furthermore, some of the biomass was observed to attach strongly onto the coal particles over time. At the end of the incubation, most of the coal powder was observed to accumulate with the biomass and form a gelatinous mixture shown in Figure 4.8.

A

G9o G9o + L G9o + B

B

Figure 4.8 A) Representative F. oxysporum G9o liquid cultures in glutamate medium with 1% w/v glucose and no coal (G9o) or 0.25% w/v lignite coal (G9o + L) or 0.25 % w/v untreated bituminous coal (G9o +B); B) A zoomed-in photo of G9o biomass and lignite coal which was observed to attach and clump together (arrow) within a gelatinous material by the end of incubation.
142 After centrifugation and filtration, yellow-coloured supernatants (Figure 4.9) were observed in the G9o-lignite cultures that became darker at each sampling time. This was followed to a lesser extent by the G9o-bituminous coal cultures. Some light colouring was also produced in the lignite-only supernatants. The G9o-only and bituminous coal-only supernatants remained clear until the end of incubation.

G9o G9o + Lignite G9o + Bit. coal Lignite Bit. coal

Figure 4.9 Supernatants of G9o with coal (lignite or untreated bituminous) and controls (G9o only and coal only). The supernatants (in triplicates) were centrifuged and filtered to obtain biomass-free and coal-free supernatants. Yellow-coloured supernatants were observed in the G9o-lignite and G9o-bituminous coal cultures that became darker at each sampling time. ‘Bit. coal’ refers to bituminous coal.

The culture supernatants were measured for absorbance at 300, 350, 400, 450 and 650 nm (Figure 4.10). Absorbance at these wavelengths has been previously used to indicate coal solubilisation (Cohen and Gabriele 1982; Hofrichter et al. 1997a; Holker et al. 1995; Laborda et al. 1997). The G9o and lignite culture supernatants showed the most significant increase in the whole incubation period (P<0.0001). The increase started to appear by the fourth day of incubation (300-450 nm) and became exponential (400-650 nm) after 30 days. These increases were not observed in other samples, including the G9o- free and coal-free controls. The G9o-lignite culture was also the only sample that showed an increase at 650 nm.

The G9o and bituminous coal culture supernatants also showed an increase in absorbance at 300-450 nm, although to a lesser and slower extent than the

143 G9o-lignite supernatants. An increase in absorbance in G9o-bituminous supernatants appeared at least 6 days after G9o-lignite (at 400 nm). The increase was significantly higher than the coal-free and G9o-free controls (P<0.05) for all wavelengths tested.

The controls (G9o-only and lignite-only) showed some increase in absorbance, but in all cases were significantly lower than the main tests (P<0.05). The increase in the controls also appeared much slower, e.g. lignite-only control showed an increase approximately 2 weeks later than G9o-lignite supernatant at 300 nm. G9o only-control supernatant showed an even more delayed increase, i.e. appearing only after 20 days of incubation. The bituminous coal control (i.e. without G9o) did not show any increase throughout the incubation period across all wavelengths.

A 3 G9o 2.5 G9o + Lignite G9o + Bituminous coal 2 Lignite Bituminous coal 300 1.5

1 Abs 0.5

0 0 5 10 15 20 25 30 35 40 Day

B 3.5 G9o 3 G9o + Lignite 2.5 G9o + Bituminous coal 2 Lignite

350 Bituminous coal 1.5

Abs 1 0.5 0 0 5 10 15 20 25 30 35 40 Day
144 C 2 G9o G9o+ Lignite 1.5 G9o + Bituminous coal Lignite

400 1 Bituminous coal Abs 0.5

0 0 5 10 15 20 25 30 35 40 Day

D 2 G9o G9o + Lignite 1.5 G9o + Bituminous coal Lignite

450 1 Bituminous coal Abs 0.5

0 0 5 10 15 20 25 30 35 40 Day

E 1.5 G9o G9o+ Lignite G9o + Bituminous coal 1 Lignite Bituminous coal 650 Abs 0.5

0 0 5 10 15 20 25 30 35 40 Day Figure 4.10 Absorbances at A) 300, B) 350, C) 400, D) 450 and E) 650 nm of G9o supernatants with and without coal for 37 days. Supernatants of the coal controls (Lignite and Bituminous coal) were also measured. Across all wavelengths, the supernatant from G9o with lignite showed the most increase, followed by G9o with bituminous coal. Error bars represent standard deviation of three independent replicates.

145
4.3.6 Extracellular oxidative enzyme activity of F. oxysporum G9o in the presence of coal
Supernatants that were tested for coal solubilsation at different wavelengths (section 4.3.5) were also analysed for extracellular enzyme activity of manganese peroxidase (MnP), lignin peroxidase (LiP) and laccase over 14 days. Overall, no indication of increasing enzyme activities in the presence of coal was observed (Figure 4.11). In fact, the G9o culture without coal had higher enzymatic activity than G9o with coal. The fungus alone showed an increase in activity for all enzymes between 0-4 days but gradually decreased thereafter. Laccase displayed the highest activity (79.12 mU/ml), followed by MnP (5.2 mU/ml) and LiP (0.5 mU/ml).

More specifically, no differences were observed between the G9o-coal cultures and coal controls in MnP activity. Except for the G9o control, enzyme activity was maintained at 1-2 mU/ml. This value was considered baseline and therefore was not a positive indication of MnP activity in G9o with coal. A gradual increase of LiP production was shown in G9o-coal cultures between Days 2-8. At the end of incubation, both the G9o-bituminous and G9o-lignite cultures had more significant LiP activity than their respective coal controls (P<0.001). However, it is noted that this increase, measured at 310 nm, was similar to those seen in the previous degradation tests measured at 300 nm (Figure 4.10). Hence, water-soluble coal components likely interfered with the enzyme analysis and therefore the increase observed in G9o-coal cultures was not considered as positive indication of LiP activity. Finally, laccase activity was present in all G9o cultures including those containing coal. However, the enzyme activity decreased by approximately 2-fold in G9o-lignite and 4-fold in G9o-bituminous cultures from the G9o control. The coal (G9o-free) controls showed no indication of laccase activity. Overall, the presence of coal in G9o cultures did not seem to stimulate the activity of all three enzymes.

146 8 A MnP G9o G9o + Lignite 6 G9o + Bituminous coal Lignite Bituminous coal 4 mU/ml

2

0 0 2 4 6 8 10 12 14 Day
0.8 B LiP G9o G9o + Lignite G9o + Bituminous coal 0.6 Lignite Bituminous coal

0.4 mU/ml

0.2

0 0 2 4 6 8 10 12 14 Day
C 100 Laccase G9o G9o + Lignite 80 G9o + Bituminous coal Lignite 60 Bituminous coal

40 mU/ml

20

0 0 2 4 6 8 10 12 14 Day

Figure 4.11 Manganese peroxidase (A), lignin peroxidase (B) and laccase (C) extracellular enzyme activity (mU/ml) by G9o, with and without coal (lignite or bituminous coal) during 14 days of incubation. Coal controls, i.e. lignite and bituminous coal without the fungus, were also included. Error bars represent standard deviation for three independent replicates.

147 4.4 Discussion

In this chapter, isolate G9o obtained from the previous isolation work (Chapter 3) was identified as the fungus Fusarium oxysporum and was characterised for its ability to colonise and degrade coal. Direct observations, SEM analysis and contact angle measurements were conducted to examine the isolate’s mechanisms of coal attachment and colonisation. Its coal degrading ability was investigated through FTIR, UV-visible spectrophotometry and enzymatic analysis.

The ITS sequence in Fusarium oxysporum G9o was compared against those of other strains. It was shown that G9o was found to be similar to F. oxysporum ES12 (accession no. DQ489299) and F. oxysporum A4 (accession no. JF779674). These strains were both isolated from soil and have been linked to low rank coal and high molecular weight PAHs degradation, respectively (Medaura et al. 2011; Silva-Stenico et al. 2006). G9o was also similar to F. oxysporum f. sp. ciceris (accession no. JN400697), a known plant pathogen. This strain has been shown to degrade phenolic complexes in plants, e.g. pectin molecules, using certain extracellular enzymes (Perez-Artes and Tena 1989).

The colonisation of bituminous coal by G9o resulted in apparent alterations to the coal that would otherwise be physically intact. At the end of incubation with the fungus, bituminous coal was fragmented into smaller pieces (Figure 4.2). As a result of the impaction of plant materials during the coalification process, coal consists of naturally wavy fracture lines (i.e. cleats) that can crack when enough mechanical pressure is applied. Thus, it is probable that the fungus followed this “path of least resistance” for colonisation and degradation, and consequently the coal pieces split along existing fractures. Similarly, Hofrichter et al. (1997b) and Bublitz et al. (1994) observed that colonisation by fungi caused macroscopic fractures of coal. Despite this observation, it is uncertain whether the physical coal fragmentation observed was a result of an active separation by the fungus, or merely due to the mechanic effects of hyphal

148 colonisation, also observed on some other inorganic materials such as bricks and concrete (Gaylarde and Morton 1999; Gu et al. 1998). Regardless of the motivation, the fragmenting ability observed in G9o is advantageous to the fungus as it increases the surface area of coal, hence leading to a more effective biodegradation.

Based on SEM observations it was evident that G9o was able to not only colonise the whole coal surface, but also to penetrate through the coal cracks and fissures (Figure 4.4). Filamentous fungi, including Fusarium sp., have been shown to cause erosion of hard surfaces e.g. rocks, minerals and concretes through hyphae penetration (Burford et al. 2003; Gaylarde and Morton 1999; Giannantonio et al. 2009; Gu et al. 1998; Warscheid and Braams 2000). This can be achieved through exerting a high amount of turgor (hydrostatic) pressure from the hyphae, which enables effective ‘burrowing’ into the solid substrate to occur (Burford et al. 2003; Money and Howard 1996). The penetrating fungus is further influenced by the substratum features, including grooves, cracks, ridges and pores, which facilitate the direction of hyphal extension and penetration (Burford et al. 2003; Watts et al. 1998). This occurrence, termed ‘thigmotropism’ or contact guidance, is a known feature of filamentous fungi that enables them to spatially explore the outer and inner surface of substratum (Jacobs et al. 2002; Watts et al. 1998). Thus, it is very likely that G9o utilised its hyphae based on this mechanism to penetrate through the pores and fine cracks of coal, which eventually fragmented to smaller pieces.

In addition to biomechanical mechanisms, building-deteriorating fungi have been shown to produce metabolites that caused further erosion of the colonised material, which enabled deeper hyphal penetration (Gaylarde and Morton 1999; Gu et al. 1998). Fungi are able to produce a wide range of organic acids, including acetic, carboxylic, citric and oxalic acids (Adams et al. 1992; Dutton and Evans 1996; Gadd 1999; Sunesson et al. 1995), which may play a role in the chemical attack of mineral surfaces (Gu et al. 1998; Muller et al. 1995). A Fusarium sp., in particular, has been shown to solubilise concrete through extensive biofilm formation and organic acids excretion, which resulted in an
149 increased permeability and porosity of the material (Gu et al. 1998). In addition to hard surfaces, phytopathogenic fungi have also been shown to penetrate plant cell walls and produce high amount of enzymes (e.g. glucanase) in the apical region of the hyphae during penetration (Bastmeyer et al. 2002).

The above examples show that colonisation of solid substrates by filamentous fungi includes both biomechanical (i.e. hyphae penetration) and biochemical (i.e. metabolite production) mechanisms occurring simultaneously rather than as separate events. This suggests that G9o and other coal-degrading fungi may follow a similar pathway in their interaction with coal. It is known that coal can be degraded through several mechanisms, i.e. depolymerisation, solubilisation and utilisation, which depend on production of various substances such as oxidative enzymes and chelating agents (Hofrichter and Fakoussa 2001). Therefore, it is likely that the substance(s) are produced simultaneously as coal is being extensively colonised, although no studies have previously attempted to demonstrate this relationship. Further investigation is therefore necessary to address this link.

The physico-chemical attachment of G9o on coal was studied through contact angle measurements. Overall, G9o exhibited a hydrophobic cell surface (> 60o, as suggested by Smits et al. (2003)) independent of its age (Figure 4.5). However, as G9o grew older, its surface hydrophobicity significantly increased (P<0.0001). Similarly, Chau et al. (2009) observed that fungi became more hydrophobic as they increased in age. A likely reason for this is sporulation and/or hydrophobin production (an adhesive material produced by filamentous fungi), which increase in some fungi as they age (Chau et al. 2009; Nakari- Setala et al. 2002; Smits et al. 2003). In light of this, Chau et al. (2009) described a new class of fungi, ‘Chronos-amphiphilic’ (i.e. Greek: time-loving both), based on fungi that have shifting hydrophobicity over time. Thus, F. oxysporum G9o can also be categorised as such, since there was a significant difference in hydrophobicity between the young and older culture of G9o over time. The ‘chronos-amphiphilic’ nature of G9o would also mean that its

150 adhesion to a hydrophobic surface (i.e. coal) would likely increase as G9o hydrophobicity increases through time.

TOT The total surface free energy (γ ) of G9o and free energy of adhesion (∆GAdh) between the fungus and coal were calculated (Tables 4.1 - 4.2). For favourable adhesion to occur, the ∆GAdh value between two surfaces must be negative. A number of factors contribute to this, including hydrophobicity, γ TOT and/or components of γ TOT of the two interacting surfaces. Overall, regardless of its age, G9o exhibited a relatively low γ TOT, which, together with high hydrophobicity, resulted in a favourable adhesion with all coal types. In contrast, earlier studies (Chapter 2) indicate that P. fluorescens was more hydrophilic and had high γ TOT that led to a smaller net difference with coal (high γ TOT) (see Figure 2.11 and Table 2.5). Therefore, although P. fluorescens showed favourable adhesion on to the untreated bituminous coal and lignite, G9o appeared to adhere twice as strongly as a result of the latter higher hydrophobicity and lower γ TOT. The more favourable attachment on coal by fungi may explain their higher prominence in coal degradation over bacteria. It is likely that G9o produced certain hydrophobins or hydrophobin-like material in its spores, both in aerial and submerged environments, which influenced its surface properties and higher likelihood of attachment to coal. It may be that the gelatinous material observed in the liquid culture (Figure 4.8) constituted this adhesive material, which resulted in the cell-coal aggregation. Although hydrophobin production has not been demonstrated by a F. oxysporum strain, several studies have indicated its presence in other Fusarium species (Fuchs et al. 2004; Sarlin et al. 2005). Further investigation is needed to prove this in G9o.

The isolate G9o showed thermodynamically more favourable adhesion towards the treated bituminous coal than the untreated one. A two-fold increase in adhesion between G9o and treated coal was calculated compared to G9o and untreated coal. This is likely due to a higher γ TOT of the treated coal than the untreated, which resulted in a larger net difference when interacting with G9o and therefore stronger adhesion forces.

151

However, based on macroscopic observations, it was clear that G9o preferred colonising the untreated over the treated bituminous coal. Very little to no cell attachment occurred on the latter, which conflicted with the ∆GAdh results. This implied that the fungus actively rejected colonising the treated bituminous coal surface and was not governed entirely by physico-chemical factors. It is uncertain as to what caused this colonisation preference by G9o. Chemotaxis in fungi was suggested by Jones (1994) as one of the primary adhesion mechanisms prior to active attachment. Hence, the untreated coal may be attractive for G9o to attach to chemotactically, which then prompted an active attachment (e.g. adhesin production) to occur.

In contrast, the altered composition on the treated coal (due to nitric acid treatment) may cause negative chemotaxis behaviour by G9o, which suggests that some coal molecules, resulting from the nitric acid treatment, were toxic to the fungus. The nitric acid treatment of coal results in a chemical incorporation of elements from nitric acid into coal molecules (Kinney and Ockert 1956). Apart from the addition of nitric acid to unsaturated structures in coal, nitric acid cleavage of other structures such as phenolic ether linkages could also occur, which leads to the formation of nitrophenols. Nitrophenols have been known as a toxicant to some microorganisms including fungi (Liu et al. 2011), and hence may be equally toxic to G9o that caused it to repel adhesion. However, it has been recently shown that Trametes versicolor, a known coal degrader (Cohen et al. 1990), was able to degrade 2-nitrophenol completely (Yemendzhiev et al. 2012). Thus, this suggests that the toxicity of treated coal may only apply to some fungi, and that the treated coal biodegradability was as a result of the ability of these fungi to utilise nitro aromatic compounds as a sole carbon source. The result also may explain the different colonising preferences on the untreated and treated bituminous coal by the fungal isolates observed in Chapter 3 (Figure 3.3).

The attachment to lignite by G9o was thermodynamically favourable. This was in agreement with the earlier observation where lignite was the most intensely colonised coal (Figure 3.12). Lignite adhesion by G9o was double the strength
152 of G9o-untreated bituminous coal adhesion. This implies that the favoured attachment on lignite gave the fungus an advantage to degrade it more effectively. Whether there is a link between rapid colonisation and effective degradation is still uncertain, however, a more favourable attachment based on physico-chemical adhesion would indeed be advantageous to coal degradation. Whilst lignite is known to be a less challenging substrate to degrade from a chemical perspective, it can also be seen as a less challenging substratum to initially attach to and colonise by filamentous fungi. Such generalisation should be exercised with caution, since not all fungi exhibit similar surface properties (i.e. high hydrophobicity and low surface energy) as F. oxysporum. Further analyses on cell surface properties of other coal-degrading fungi need to be conducted to reach a conclusion.

The most thermodynamically favourable adhesion in this study was between the least matured G9o and lignite (Table 4.2). Indeed, G9o on Day 2 had the highest degree of physico-chemical interaction with lignite, surpassing that of older cultures and other coal surfaces by at least two-fold. The relationship between G9o and lignite was similar to that of P. fluorescens and lignite (see Table 2.6 and Figure 2.12 for P. fluorescens studies), which indicates the superiority of lignite in terms of adhesion across different microbes. However, it is important to note that in the colonisation and degradation assay (Chapter 3), the isolates, including G9o, were exposed to both lignite and bituminous coal pieces only after 1.5 to 2 weeks of incubation, so G9o was not in its earliest growth phase. Nevertheless, the results show that contact between the fungus and lignite at an early stage of growth would make a significant difference in the amount of biomass attached on the coal compared to in a later growth stage. This is an important consideration in the context of biotechnological applications of the fungus to degrade coal.

Based on infrared spectrometry, F. oxysporum G9o showed positive indications of untreated bituminous coal and lignite degradation (Figures 4.6 - 4.7). Clear changes in the coal spectrum strongly suggest that chemical modification of

153 coal was due to the fungal presence, which confirms the ability of G9o to modify coal.

A main structural change in coal as a result of G9o modification lies at 1262 cm- 1, which corresponds to a C-O vibration (Painter et al. 1980). The peak appeared in the untreated bituminous coal exposed to G9o, suggesting oxidation of coal by the fungus. The peak position matches that expected for ester C-O or aromatic ether C-O (Socrates 2001), which has been proposed by Gao et al. (2012). Interestingly, the same peak was also observed in the unmodified lignite coal spectra (Figure 4.7). Lignites are naturally highly oxidised and would contain a diverse range of oxygen-based functional groups including those containing the C-O bond. In contrast to the untreated bituminous coal, the peak disappeared in the modified lignite spectra, which implied that G9o modified the C-O bond during incubation. The presence of the C-O bond in the bituminous coal and absence in lignite after G9o incubation suggests that there is a continuum of coal degradation by G9o. Depending on the coal type (or ‘phase’), the G9o mechanism of C-O attack seems to include oxidation and possibly cleaving reactions of the C-O bond, which occur over time. The initial oxidation of bituminous coal could be seen as a “reversion” to a lower-ranked coal (structurally similar to lignite) followed by a further C-O oxidation or cleaving activity as observed in the modified lignite. A hypothetical example of such a chemical pathway is given in Figure 4.12, where a compound in the unmodified coal (a) is oxidised to become a phenol residue (b), explaining the presence of the 1262 cm-1 peak. The C-O bond may be cleaved over time by the fungus, or the phenol (b) is further oxidised to quinone, which explains the peak disappearance.

154 (a) (b) (c) Figure 4.12 A possible pathway for oxidation of a hypothetical coal residue. Compound (a) constitutes an unmodified coal, which is oxidised to phenol (b) to give a C-O bond observed at 1262 cm-1 in the IR spectrum. Further oxidation of (b) leads to quinone (c), which causes the 1262 cm-1 peak to disappear.

An important question arises from this analysis: does chemical modification by G9o correlate with its colonisation on coal? Earlier, it was observed that G9o preferred colonising untreated bituminous coal to treated coal, which is in conflict with thermodynamic data. It was apparent from the 1262 cm-1 C-O peak that the untreated coal was more oxidised than the treated coal, and that changes to the assigned aromatic C-H bond at 749 cm-1 (Painter and Rhoads 1981) and C=C, C-O or OH bond at 1436 cm-1 (Gomez-Serrano et al. 2003) were observed only in the former. There were observable changes to peaks in the G9o-modified treated coal spectrum, however, these peaks probably belonged to bonds in silica and clay minerals such as kaolin and CaSO4 (Estep et al. 1968; Gomez-Serrano et al. 2003; Miller et al. 2012), suggesting that changes to these peaks were more likely due to natural weathering than biodegradation effects by the fungus. These changes were also observed in the modified untreated coal, although to a lesser extent. This may be due to the coal surface being less exposed to oxygen (due to complete colonisation) than the treated coal (i.e. where colonisation did not occur) over three months of incubation. Furthermore, changes to the 1714 cm-1 peak assigned to the C=O bond (Fredericks et al. 1983; Painter et al. 1980) were apparent only in both lignites but not the treated coal after exposure to G9o. This implies that modification of C=O bond occurred in only lignites (both fully colonised by G9o) and not the treated coal despite its presence in the latter. These examples indicate that there were hardly any changes to the treated bituminous coal
155 resulting exposure to G9o, which correlates with its lack of colonisation by the fungus. Hence, this suggests that colonisation was necessary for coal degradation to occur by G9o.

Apart from the degradation observed in the aerial environment, F. oxysporum G9o also showed ability to solubilise coal in the aqueous phase (Figures 4.8 – 4.10). The yellowish color observed in the G9o-coal supernatants (Figure 4.9) was indicative of water-soluble components in coal, e.g. humic acids or phenolics leaching into the medium (Hofrichter and Fritsche 1997). Similar to the non-aqueous experiments, lignite was substantially degraded in liquid media based on the spectrophotometry analysis. The degradation of bituminous coal was also apparent (although slower than that of lignite), which showed the capacity of the fungus to degrade a higher-ranked coal both in aerial and submerged conditions.

Fusarium oxysporum G9o could grow fully submerged and in close contact with coal (Figure 4.8). The fungus exhibited a different morphology in liquid, where gelatinous material was evident mid-way through incubation. Since G9o mycelia were highly hydrophobic, it was likely that the viscous material was exopolymeric substance (EPS) that helped the fungus to stay submerged. This observation was documented in previous studies on coal degradation by fungi in the aqueous phase (Achi and Emeruwa 1993; Igbinigie et al. 2010; Laborda et al. 1997). The EPS production is advantageous from a coal degradation perspective as aggregation resulted in direct contact between fungal cells and coal. Although it has been shown that cell-free supernatants were able to degrade coal, e.g. through their extracellular enzymatic system, cell to coal contact has been demonstrated as an important parameter for effective degradation (Wilmann and Fakoussa 1997b). Thus, the ability of G9o to adhere to coal and degrade it in the aqueous phase is useful especially for field applications where coal is submerged in groundwater.
In conjunction with the spectrophotometry analyses, the G9o culture supernatants were analysed for extracellular enzymes production in the absence and presence of coal (Figure 4.11). Overall, there was no positive

156 impact of coal on the production of manganese peroxidase (MnP), lignin peroxidase (LiP) and laccase. In the presence of coal, an absence or decrease in the enzyme activity was observed, which indicated a potential inhibitory effect of coal on the assays. Previous studies have shown that peroxidases were competitively inhibited by humic acid or water-soluble components from coal, which led to a decrease in the enzymatic activity (Hofrichter and Fritsche 1997; Ralph and Catcheside 1994; Wondrack et al. 1989). Thus, it is possible that the enzymes in the G9o-coal cultures were produced at a similar (or higher) rate than in the G9o control, but were inhibited by coal complexes in the medium. Furthermore, the presence of glutamate (a component of the growth medium) has been known to suppress the enzymatic activities of LiP and laccase (Holker et al. 1999; Kirk and Farrell 1987). This may have been true for LiP, as it showed very low activity in all cultures, but not true for laccase, which showed high activity (although lower in the presence of coal).

Physico-chemical interactions between coal and enzymes may also be responsible for enzymatic inhibition. Coal, particularly pulverised coal, tends to adhere to surfaces. It is likely that the extracellular enzymes adhered to the fine coal particles that were removed from the supernatant for the enzymatic tests. Although this was not tested as part of this work, an experiment investigating the adhesion of proteins to coal was conducted in our laboratory, where only less than 50% of proteins added to medium containing powdered coal was recovered (Gutierrez-Zamora 2013). Thus, enzymes were likely to have adhered to coal in a similar manner.

The enzymatic analysis suggests extracellular MnP, LiP or laccase do not play a role in coal degradation by F. oxysporum G9o. Even if these enzymes did contribute to coal degradation, the effect would be based on secondary metabolite production, since G9o produced the enzymes in the absence of coal. It is not known what metabolites activated the synthesis of these enzymes, particularly laccase, in cultures with no coal. F. oxysporum has been shown to degrade lignite coal based on other mechanisms, such as alkalinisation of the medium and chelators (Holker et al., 1999), although extensive work is necessary to investigate this further, particularly for bituminous coal
157 degradation. Based on the FTIR analysis conducted in this study, G9o showed an array of coal chemical modifications, which may reflect diverse mechanisms of degradation.
In summary, this study was conducted to characterise the mechanism of attachment, colonisation and degradation of coal by F. oxysporum G9o. The fungus extended its hyphae following the natural fissures of coal ultimately colonising the entire coal surface. The penetration mechanism observed through SEM and a significantly hydrophobic cell surface of G9o showed its advantages in colonising coal over P. fluorescens, which may explain the higher prevalence of filamentous fungi in degrading coal compared with bacteria. Surface thermodynamic analyses showed that the interaction between fungi and coal was favourable. However, depending on the growth stage and the type of coal, varying degrees of thermodynamically favourable interactions were observed. In addition, adhesion between G9o and coal was governed not only by thermodynamics, but also by chemical and/or biological factors.

Importantly, the FTIR and UV-visible spectrophotometric analyses showed that G9o was capable of degrading bituminous coal and lignite, which confirmed the direct observation on coal degradation by this fungus. This study is the first to show bituminous coal degradation by F. oxysporum, which has previously been linked only to low rank coal degradation. The FTIR results also indicated a possible correlation between colonisation and coal modification by G9o, which suggests that cell attachment to coal is necessary for effective degradation to occur. The fungus also showed its ability to grow and degrade coal in an aqueous phase, an advantageous feature for field applications. The coal degradation by G9o did not seem to be influenced by oxidative enzymes, implying that other coal degrading mechanisms such as alkalinisation and chelation were utilised. Further work on degradation mechanisms is thus necessary to completely characterise the coal degrading ability of Fusarium oxysporum G9o.

158 Overall, the attachment and degradation studies conducted provide fundamental insights into the interactions between fungi and coal, which are useful in the potential application of these microbes in the field.

159 5 A microbial community analysis of bituminous coal buried in soil from the Lithgow State Coal Mine, New South Wales

5.1 Introduction

A large proportion of biofilm studies have focused on single-species biofilms (Allison et al. 1998; Danese et al. 2000; Davies et al. 1998; Klausen et al. 2003; Tolker-Nielsen et al. 2000; Webb et al. 2003) to investigate various fundamental aspects in cell attachment to and colonisation of surfaces. Whilst pure isolate studies remain an important field in biofilm research, they do not necessarily reflect the true conditions that occur in natural environments. Microorganisms rarely live in isolation in natural settings, but exist in complex and heterogeneous communities as biofilms (Burmolle et al. 2006). Simplified models have been used to study various aspects governing the natural-existing biofilms, which include mixed-species interactions e.g. symbiosis, competition, predation (Burmolle et al. 2006; Kawarai et al. 2007; Rao et al. 2005; Rypien et al. 2010) and their response to different environmental factors e.g. shear stress (Besemer et al. 2007; Zhang et al. 2013). Part of the essential tools in studying the impact of these factors are through analysing changes in the communities and identifying the role of individual populations in the community. This can be achieved by employing microbial community fingerprinting techniques such as Denaturing Gradient Gel Electrophoresis (DGGE) (Muyzer et al. 1993) or Terminal-Restriction Fragment Length Polymorphism (T-RFLP) (Liu et al. 1997), and next-generation sequencing methods such as 454 pyrosequencing (Goldberg et al. 2006) or Illumina sequencing (Qin et al. 2010). Utilising these technologies enable us to address specific questions towards understanding the significance of biofilms in the environment.

In this chapter, microbial community analysis was applied to investigate the role of coal as a substratum for cell attachment and biofilm formation in a complex
160 multi-species environment. Forest soil, specifically from the vicinity of the Lithgow State Coal Mine (LSCM), New South Wales, was chosen as the environment in which coal was exposed. Soils are usually associated with high microbial diversity (Gans et al. 2005; Torsvik et al. 1990) and soil microbes have been known to degrade a variety of hydrocarbons and natural organic matter like cellulose, humus and lignin as well as pollutants such as oil and chlorinated compounds (Bouchez et al. 1995; Ghazali et al. 2004; Thompson et al. 1998). This makes soil a suitable environment to select specific microbial communities with the potential to adhere to coal surfaces, as coal is rich in carbon and complex organic matter. Thus, the hypothesis is that coal plays the role of a ‘biotrap’ in selecting unique microorganisms that have the ability to adhere and take advantage of this complex carbon source.

There were two main aims in this study: 1) to compare the soil community adhering to coal with the original soil community over time. The expectation was that the coal community composition would be different from that in soil, as coal is potentially an attractive but selective carbon substrate. T-RFLP fingerprinting was employed to observe the community differences. 2) To identify key microorganisms from soil found to be abundant on coal. A majority of microbes that are dominant on coal are expected to be significantly different from the dominant microbes in soil due to several differences (e.g. substrate types) between the two niches. A 454 pyrosequencing analysis was utilised to achieve this aim.

5.2 Material and methods
5.2.1 Experimental set up

Surface soil (topsoil) from a depth of approximately 20 cm deep was obtained in the forest vicinity of the Lithgow State Coal Mine (LSCM), New South Wales, Australia in October 2011. The topsoil horizon of this soil contained fragmented pieces of coal artificially deposited from previous mining activities. The soil was brought to the laboratory and stored in the dark at room temperature (22 oC).

161 Immediately before use, the soil was sieved to obtain particles less than 2 mm in diameter, mixed thoroughly and transferred into a pot 25 x 60 x 27 cm where soil depth was approximately 20 cm. Untreated and nitric-acid treated coal pieces of approximately 2 x 1.5 x 1 cm each were randomly placed in a designated area at 10-12 cm deep. As both coal types have demonstrated differences in chemical composition and preferential colonisation (Chapter 3), it is hypothesised that there will be differences in the community composition attached to each coal type. The soil and coal mixture was incubated at 22 oC in dark for 27 weeks.

The moisture content was determined through dry weight measurements. Soil moisture was maintained between 20-22% (grams of water [gram dry weight of soil]-1) by using chlorine-free water, which was recollected and reused for watering to minimise nutrient loss in soil.

5.2.2 Coal sampling and DNA extraction

Sampling of the coal pieces was conducted at weeks 0, 2, 4, 10, 17 and 27. At each time point, four samples (replicates) of the untreated and treated coal were randomly selected and removed from the soil. The coal pieces were lightly vortexed in phosphate buffer solution (PBS) for 5-7 s and gently rinsed in PBS for three times to remove non-adhered soil particles. To obtain adhered biomass on coal, the coal pieces were scraped using sterile cell scrapers (Sarstedt, Nümbrecht, Germany). Approximately 500 μL of PBS was used to rinse the cell scraper and coal piece to obtain the scraped and loosely attached material. In addition to coal samples, four soil sample replicates (500 mg each) that were in close proximity with the extracted coal pieces in the soil were used as reference for the microbial community analyses.

The scraped biomass (500 μL) and soil sample (500 mg) underwent DNA extraction using the method described by Li et al. (2008) with modifications: 1 ml of the extraction buffer (100 mM Tris HCl pH 8, 100 mM EDTA pH 8 and 1.5 M NaCl in MilliQ water) was aliquoted into each tube containing the scraped

162 biomass or soil sample and 200 μL of zirconia/silica beads (BioSpec Products, Inc., USA). Tubes were placed in a FastPrep machine (FP120, Thermo Savant, Canada) and shaken at 5.5 m/s for 45 s and 15 with an interval of 1 min where the tubes were placed on ice. The tubes were centrifuged at 4,500 x g for 1 min before adding 200 μL of 20% sodium dodecyl sulfate (SDS) and incubated at 65 oC for an hour in a water bath. The tubes were centrifuged again at 4,500 x g for 5 min and the supernatant was transferred into a sterile tube. Half a volume of the supernatant was mixed with 30% PEG-8000 (in 1.6 M NaCl solution) and tubes were left incubating at room temperature overnight. The tubes were centrifuged at 16,000 x g for 20 min and the pellets were dissolved in 100 μL of TE buffer and 100 μL of 1 M potassium acetate solution was added before placing all samples on ice for 5 min. The tubes were centrifuged at 16,000 x g for 10 min at 4 oC and the supernatant was mixed with one volume of phenol: chloroform: isoamyl alcohol (25:24:1) solution. Centrifugation at 16,000 x g for 3 min proceeded and the aqueous phase was retrieved and mixed with chloroform: isoamyl alcohol (24:1) before centrifuging again at 16,000 x g for 3 min. The aqueous phase was retrieved and mixed with 0.6 volume of isopropanol and 1.3 μL of 15 mg/mL GlycoBlue (Life Technologies Inc., USA) and incubated overnight at 4 oC. The tubes were centrifuged at 16,000 x g for 20 min at room temperature and pellets were washed with 70% ethanol before reconstituting with 30 μL TE buffer. DNA yields were quantified using a Qubit® fluorometer (Life Technologies Australia Pty Ltd., Australia).

5.2.3 Bacterial 16S rRNA gene amplification of the scraped biomass on coal and LSCM soil

DNA amplification targeting the 16S rRNA gene using the 27F-FAM and 530R primers (refer to Appendix II for primer sequences) was conducted for all samples.

The PCR mixture (50 μL total volume) for each sample consisted of 25 μL of 2x Promega PCR Master Mix (Promega Corp., USA) which contained 50 units/ml of Taq DNA polymerase supplied in a proprietary reaction buffer (pH 8.5), 400

163 μM dATP, 400 μM dGTP, 400 μM dCTP, 400 μM dTTP and 3 mM MgCl2, 1 μL of 10 μM each forward and reverse primer, 21 μL of molecular biological grade water and 2 μL of the extracted DNA. The amplification was conducted for two times each sample to achieve a 100 μL total volume of amplified DNA.

Using an MJ Mini Personal Thermal Cycler (Bio-Rad Laboratories Inc., USA), the thermal cycling PCR profile for the amplification was as follows: Initial denaturation at 94 oC for 3 min; 30 cycles of denaturation at 94 oC for 30 s, annealing at 55 oC for 30 s, extension at 72 oC for 30 s followed by a final extension at 72 oC for 3 min.

All PCR products were verified using electrophoresis on a 1.5% agarose gel and stained with SYBR® Gold nucleic acid gel stain (Molecular Probes, Inc., USA). Products were visualized by UV transillumination and digitally photographed with a Molecular Imager® Gel Doc™ XR System (Bio-Rad Laboratories Inc., USA).

PCR products were then purified using a DNA Clean & ConcentratorTM-5 kit (Zymo Research Corp., USA), verified by electrophoresis for DNA presence and quality inspection and quantified using a Qubit® 2.0 fluorometer (Life Technologies Australia Pty Ltd., Australia).

5.2.4 Terminal restriction fragment length polymorphism (T-RFLP) of 16S rRNA genes extracted from biomass in soil and on coal

T-RFLP was used as a method for analysing microbial communities on coal and soil by comparing the profiles of restriction fragments (T-RFs) of each sample. In this study the amplified and purified DNA for each sample was digested with the restriction enzyme RsaI that cleaves DNA at the sequence 5’-GT|AC-3’ giving blunt ends. Negative (no DNA) and positive (Escherichia coli DNA) controls were included.

164 The restriction digestion mixture for each sample contained the following: 5-10 ng/μL purified DNA, 400 μg/mL RsaI restriction endonuclease and 1x of NE Buffer 4 (New England Biolabs, Inc., USA), and molecular biology grade water to top up a final volume of 25 μL. In cases where the DNA concentration was too low, the maximum volume of DNA sample was used without dilution.

The digestion procedure was conducted at 37 oC for 3 h and stopped at 65 oC for 20 min to inactivate the enzyme activity. Digested products were then purified using a DNA Clean & ConcentratorTM-5 kit (Zymo Research Corp., USA) and eluted with molecular biology grade water before sent for fragment analysis using an Applied Biosystems 3730 DNA Analyzer (Life Technologies Corp., USA) at the Ramaciotti Centre for Gene Function Analysis (University of New South Wales, Sydney, Australia). The Applied Biosystems GeneScan™ - 500 LIZ® (Life Technologies Corp., USA) was used as an internal lane size standard.

Electropherograms of the data were analysed using Applied Biosystems Peak Scanner 1.0 (Life Technologies, Inc., USA) and T-REX (Culman et al. 2009) software to prepare the data for statistical analysis.

5.2.5 454 pyrosequencing and quality filtering

To identify key microbes from soil that adhered to coal, selected DNA samples from weeks 17 and 27 for both coal and soil were sequenced using a Roche Genome Sequencer FLX 454 (Roche Diagnostics Corp., USA) at the Research and Testing Laboratory (Texas, USA). The 926wF and 1392R primer pair (see Appendix II for sequences and reference) was used to target the 16S rRNA genes (archaea and bacteria) and 18S rRNA genes (eukaryotic). The raw sequencing data were processed for quality filtering, aligned and sub-sampling using the 454 Standard Operating Procedure (SOP) of the Mothur software (Schloss et al. 2009; Schloss et al. 2011). The ribosomal database project (RDP) SILVA databases were used for taxonomical identification.

165 5.2.6 Statistical analyses for T-RFLP and pyrosequencing data

Statistical analyses of the T-RFLP fragments and pyrosequencing data were conducted using PRIMER 6 with PERMANOVA+ software (PRIMER-E Ltd, Ivybridge, UK). Permutational multivariate analysis of variance (PERMANOVA) was used to determine the statistical significance between groups of samples (i.e. differences in microbial community composition between soil and on coal surface). The ‘similarity percentages’ (SIMPER) analysis was used to determine the percentage contributions of fragments/species/operational taxonomy units (OTUs) that contributed most to the dissimilarity between two groups of samples. Non-Metric multidimensional scaling (nMDS) plots were generated to present similarities or dissimilarities between groups of samples based on the calculation of distances using the Bray-Curtis measure of similarity. The data was transformed via log (x+1) transformation.

5.2.7 Scanning Electron Microscopy of coal pieces exposed to soil

Coal samples subject to soil were observed for surface colonisation using Scanning Electron Microscopy (SEM). At the end of incubation time (over week 27), the untreated and treated coal samples were extracted, vortexed lightly in PBS for 5-7 sec and rinsed gently for three times in PBS. The coal sample underwent similar SEM procedures of glutaraldehyde fixation, ethanol dehydration, HMDS drying and gold sputtering as outlined in Section 2.2.4 before imaging using an ESEM Quanta 500 microscope (FEI, Oregon, USA) at 10 kV.

5.3 Results

5.3.1 DNA extraction and amplification of biomass on coal

Soil biomass that adhered to coal was scraped from the coal surface and subjected to DNA extraction and amplification of rRNA genes. Overall, the

166 average amplified DNA yield of the scraped biomass from both coal types was low (< 1 ng/µL) compared to that of soil (6.7 ± 3 ng/µL), particularly at weeks 0 - 10. However, the amplicon concentration from coal was observed to increase particularly in Weeks 17 and 27 as shown by a representative agarose gel (Figure 5.1). There was more DNA (i.e. approximately two times at week 17 and three times greater at week 27) in the untreated coal sample than in the treated one.

M Week 0 Week 2 Week 4 Week 10 Week 17 Week 27 Negative 1 2 3 4

Figure 5.1 Representative agarose gel illustrating amplified DNA from the scraped biomass (SB) of the untreated coal (Lane 1) and treated coal (Lane 3) against their reference soil samples (Lane 2 and 4 respectively) over the sampling time (week). The DNA yields for the SB samples were lower in comparison to those of soil, however they started to increase in weeks 17 and 27, particularly from the untreated coal sample (full arrow). Very faint bands could be seen for SB of the treated coal sample (dashed arrow) in the final two weeks of incubation. ‘Negative’ refers to negative control (i.e. no DNA template) and M refers to a low range DNA marker (GeneRuler 1kb DNA ladder, Thermo Scientific Inc., USA).

5.3.2 Terminal restriction fragment length polymorphism (T-RFLP) of DNA extracted from soil and biomass on coal
The 16S rRNA genes extracted from soil and coal surfaces were subject to T- RFLP analysis to obtain bacterial community fingerprints from both sources over time. Despite the low DNA yield in coal samples particularly for weeks 0 – 10 for the untreated coal and weeks 0 – 27 for the treated coal, fragments could still be detected albeit in much lower intensity than those of soil (see Appendix VI for T-RFLP electropherograms for all samples). Relevant peaks could be

167 observed in soil and coal that were absent in both the negative (no template) and positive (E. coli) control.

Figure 5.2 shows the non-metric multidimensional scaling (nMDS) plots based on the distance (differences) calculated in the T-RF profiles of each sample. Overall, the community compositions (derived from the T-RFs fingerprint) on the untreated and treated coal were observed to be significantly different against the soil community throughout the 27 weeks of incubation (P=0.0001). No significant differences were found between soil samples over time (P=0.37) suggesting that the soil community remained relatively static during the incubation period. Nevertheless, the community composition on coal became less different than soil over time. Starting from weeks 10-17, the coal samples showed a converging trend moving towards the soil plots. This was more obvious for the untreated coal samples, where the untreated coal plots seemed to be closer to soil than the treated coal. However, there were still significant differences in composition between both coal types and soil at the end of the incubation (P=0.01).

The coal replicates were distinct from each other from weeks 0 – 10 (Figure 5.2). This is further evident in Figure 5.3, where a lower average similarity percentage among coal replicates was observed from week 0 (20-30%) compared to the soil replicates that were at least 2.5 times higher in similarity. However, the similarity increased substantially for the untreated coal replicates to approximately 70% by week 27, which was almost similar to those of coal. This was followed by the treated coal replicates to a lesser extent (i.e. approximately 50%). There was a fluctuation in the similarity percentage within the treated coal replicates at weeks 2 and 4, which corresponded to the fluctuating distance from the soil sample community observed in the MDS plot. However this fluctuation stabilised and the similarity increased gradually after week 4.

168

A: Untreated coal

B: Treated coal

Figure 5.2 Non-metric multidimensional scaling (nMDS) plots of the T-RFs of scraped biomass of A) untreated coal and B) treated coal samples against their respective soil reference samples. The plots were generated by calculating distances using the Bray-Curtis measure of similarity. The data was transformed using log (x + 1) transformation. ‘Soil0-27’ and ‘ScrapedCoal0-27’ refer to the soil and scraped biomass on coal, respectively, at 0, 2, 4, 10, 17 and 27 weeks of incubation. Significant differences (P=0.0001) were found between the overall 27-week community composition on coal and in soil for both the untreated and treated coal. Despite the converging trend of coal plots moving closer to the soil plots, significant differences were still observed between coal and soil at weeks 17 and 27 (P=0.01).

169 100

80

60

40 Untreated coal Average similarity (%) Average 20 Treated coal Soil (Untreated coal) Soil (treated coal) 0 0 5 10 15 20 25 Weeks
Figure 5.3 Average similarity (%) between the community composition among replicates for each sample (Untreated and treated coal and their respective soil samples) over 27 weeks of incubation.

Direct comparison between individual T-RFLP fingerprints of coal and soil revealed some differences in the presence and relative intensity of specific fragments. Figure 5.4 shows a representative T-RFLP electropherogram for each sample at Week 27. Whilst both soil and coal exhibited similar T-RF patterns, several fragments (e.g. 243, 307 and 524 bps) were higher in intensity in the untreated coal than soil. There were also fragments, e.g. 74 bp, substantially reduced in the untreated and treated coal but present in soil. A substantial reduction in intensity of another fragment (i.e. 524 bp) was also observed in the treated coal but not in soil. Importantly however, the exact trend was not consistent for all coal replicates. There was higher variability in the presence or relative intensity of unique fragments in other coal replicates against those of soil, which is further observed in the pyrosequencing results (section 5.3.3). Nevertheless, individual T-RLFP profile comparison between soil and untreated coal showed unique differences in soil and coal communities.

170 Negative

106 420 Soil (Untreated)

74 524

106 Soil (Treated) 420 74 524

524 Untreated coal 106 420

243 307

106 420 Treated coal

Figure 5.4 Representative 16S rDNA T-RFLP electropherograms of soil and coal at week 27 (see Appendix VI for all profiles). The x-axes represent fragment length whilst the y-axes indicate fluorescence intensity of fragments. All profiles were adjusted to the same intensity (y- axis) scale except for Treated coal, which produced lower intensity fragments resulting from a low DNA concentration in the sample. The general fragment pattern nevertheless was similar across both samples. Unique fragments were observed in the untreated coal sample (full arrows) that were not apparent or less abundant in soil. Several fragments observed in soil were substantially reduced in relative intensity in the untreated coal and/or treated coal samples (dashed arrows). The peak observed in the negative (template-free) sample corresponded to a fragment less than 30 bp, i.e. the minimum fragment generated, and therefore was considered an artefact.
171
5.3.3 Pyrosequencing results

Coal and soil DNA samples specifically from weeks 17 and 27 (where most abundant DNA were found) were analysed for pyrosequencing to identify dominant microbes on coal that were different in terms of presence/absence and/or relative abundance from those in soil. As all the soil samples showed high similarity in microbial composition and abundance (P=0.37), only the soil from the proximity of the untreated coal was used. A total of 148,297 sequences were retrieved. Quality filtering removed poor quality sequences, chimeras and overlapping regions after alignment, resulting in 8,622 sequences. The trimmed sequences were taxonomically assigned using the SILVA 16S and 18S rRNA gene databases, and further clustered into Operational Taxonomic Units (OTUs). One OTU was defined by sequences that were above 97% similarity with sequences deposited in the databases.

A total of 3,470 bacterial/archaeal OTUs and 337 eukaryotic OTUs were classified. The first ten most abundant OTUs, or those above the top 2.5% abundance in each sample were used for comparison. The OTUs were processed in silico to the species level using Mothur, however due to the inherent short sequence lengths of pyrosequencing, the OTUs were taxonomically allocated to its lowest identifiable taxonomic rank (the lowest being genus). There were instances where unclassified bacteria or archaea OTUs were found. In cases where two or more OTUs shared the same lowest classification, a number was assigned to each of them (e.g. Bacillus sp. 1, Bacillus sp. 2 etc.).

5.3.3.1 Archaeal and bacterial OTUs
Figure 5.5 illustrates an overview of the top ten most abundant OTUs across all samples (for raw data see Appendix VII). A significant difference of abundant OTUs was observed between coal and soil samples at weeks 17 and 27 (P=0.0001), which is consistent with the T-RFLP analysis (Figure 5.2). In

172 contrast to coal, soil samples showed more similarity in abundant OTUs among its replicates and across different time points. For example, OTUs identified as ‘Acidobacterium sp. Gp2-2’, ‘Alphaproteobacteria 1’, ‘Gammaproteobacteria 2’, ‘Nocaridioiceae 1’, ‘Rhizobiales 1’ and ‘Unclassified bacteria 1’ were found in one or more soil replicates at both time points. However, a number of unique abundant OTUs in soil were also observed. Soil replicate S1wk27 showed the highest number of unique abundant OTUs, illustrated in the nMDS plot of distances in OTUs in Figure 5.6.

Different from soil, coal samples contained more unique abundant OTUs (Figure 5.5). Although a small number of abundant OTUs was shared between coal and soil (e.g. ‘Acidobacterium sp. Gp1-1’, ‘Alphaproteobacteria 1’ and ‘Unclassified bacteria 1’ in the treated coal and soil samples at week 17), a larger number of OTUs in coal samples was not found abundant in soil at any time point. Further, many of the OTUs in each coal sample were not only different between coal types (i.e. untreated and treated), but were also independent of replicates and time. This was shown further by the nMDS plot of OTUs in each sample (Figure 5.6), where differences in OTUs between coal samples were larger than between soil samples.

Despite a low number of OTUs shared between coal samples, several abundant OTUs were identified. At week 27, ‘Acidobacterium sp. Gp1-3’, ‘Peptococcaceae_1’ and ‘Phenylobacterium sp. 3’ were found abundant on the treated coal, and ‘Rhizomicrobium sp. 1’ on the untreated coal. ‘Acidobacterium sp. Gp1-3’ present on the treated coal at week 27 was also found abundant at week 17. There were also abundant OTUs, e.g. ‘Rhizobium sp. 2’, Solirubrobacter sp. and Staphylococcus sp. at week 27, shared between both the untreated and treated coal.

173 110

Verrumicrobia subdiv. 3 sp. Unclassified bacteria 10 100

Unclassified bacteria 1 Staphylococcus sp. 90 Streptococcus sp. Solirubrobacter

80 Rhizobiales 1 Rhizomicrobium sp. 2 Pseudolabrys sp. Rhizomicrobium sp. 1 70 Phenylobacterium sp. 3 Peptococcaceae_1 Pasteuria sp.

60 Nocardioidaceae 1

50

Gammaproteobacteria 2 Gammaproteobacteria 1 40 Burkholderia

Individual OTUs (1-101) 30 Alphaproteobacteria 1 20 Acidobacterium sp. Gp2-2 Acidobacterium sp. Gp2-1 Acidobacterium sp. Gp1-3 10 Acidobacterium sp. Gp1- 1 0

T1 
T2 
U1 
U2 
S1 
S2 
S3
T1
T2
T3 

U1 U2 
U3 
S1 
S2 

S3 
-10 wk17 wk17 wk17 wk17 wk17 wk17 wk17 wk27 wk27 wk27 wk27 wk27 wk27 wk27 wk27 wk27

Figure 5.5 Overview of 101 individual OTUs (y-axis) across individual coal and soil samples (x-axis) at weeks 17 and 27. Each circle corresponds to one OTU (see Table 5.1 for OTU names). The 10 most abundant OTUs are displayed for each sample. Grey circles correspond to unique OTUs found in respective samples. Assorted coloured circles denote shared OTUs between two or more samples. T= Treated coal, U= Untreated coal, S= Soil, wk17= Week 17 and wk27= Week 27. Dashed lines separate each set of sample replicates. Double dashed line separates samples into weeks 17 and 27. Circle area signifies the abundance of each OTU.

174

Table 5.1 Operational taxonomic units (OTUs) and their corresponding y-axes label in Figure 5.5.

Operational Y-axis Taxonomic Unit (OTU) Y-axis OTU Y-axis OTU Y-axis OTU Y-axis OTU Phenylobacterium 1 Abiotrophia sp. 23 Alphaproteobacteria 3 45 Hymenobacter sp. 67 sp. 3 89 Unclassified bacteria 1 Phenylobacterium 2 Acetobacteraceae 24 Alphaproteobacteria 4 46 Kitasatospora sp. 68 sp. 4 90 Unclassified bacteria 2 3 Acidimicrobiales 1 25 Bacillales 1 47 Ktedonobacter sp. 69 Planctomycetaceae 91 Unclassified bacteria 3 Porphyrobacter sp. 4 Acidimicrobiales 2 26 Bacillales 2 48 Ktedonobacterales 70 1 92 Unclassified bacteria 4 Acidobacterium sp. Gp1- Methanococcoides sp. Porphyrobacter sp. 5 1 27 Bacillales 3 49 1 71 2 93 Unclassified bacteria 5 Acidobacterium sp. Gp1- Methanococcoides sp. Porphyrobacter sp. 6 2 28 Bacillales 4 50 2 72 3 94 Unclassified bacteria 6 Acidobacterium sp. Gp1- Methanococcoides sp. Propionibacterium 7 3 29 Bacillus sp. 51 3 73 sp. 95 Unclassified bacteria 7 Acidobacterium sp. Gp1- Methanococcoides sp. 8 4 30 Betaproteobacteria 52 4 74 Pseudolabrys sp. 96 Unclassified bacteria 8 Acidobacterium sp. Gp1- Methanococcoides sp. 9 5 31 Bradyrhizobium sp. 1 53 5 75 Pseudomonas sp. 97 Unclassified bacteria 9 Acidobacterium sp. Gp2- Unclassified bacteria 10 1 32 Bradyrhizobium sp. 2 54 Microbacteriaceae 76 Rheinheimera sp. 98 10 Acidobacterium sp. Gp2- 11 2 33 Brevundimonas sp. 55 Microbacterium sp. 77 Rhizobiales 1 99 Variovorax sp. Acidobacterium sp. Gp2- Verrumicrobia 12 3 34 Burkholderia sp. 56 Nocardioidaceae 1 78 Rhizobiales 2 100 subdivision 3 sp. Rhizomicrobium sp. 13 Acidobacterium sp. Gp6 35 Caldilinea 57 Nocardioidaceae 2 79 1 101 Xanthomonadaceae Acidobacterium sp. Rhizomicrobium sp. 14 Gp10 36 Chitinophagaceae 58 OP11 sp. 80 2 15 Acidovorax sp. 37 Cloacibacterium sp. 59 Oxalobacteraceae 1 81 Sinobacteraceae 16 Actinobacteria 1 38 Gammaproteobacteria 1 60 Oxalobacteraceae 2 82 Solirubrobacter sp. 17 Actinobacteria 2 39 Gammaproteobacteria 2 61 Paenibacillaceae_1 83 Staphylococcus sp. 18 Actinomycetales 1 40 Gammaproteobacteria 3 62 Pasteuria sp. 84 Streptococcus sp. 19 Actinomycetales 2 41 Gammaproteobacteria 4 63 Peptococcaceae_1 85 Tepidimonas sp. Thermomonosporac 20 Actinomycetales 3 42 Halolactibacillus sp. 1 64 Petrimonas sp. 86 eae Phenylobacterium sp. 21 Alphaproteobacteria 1 43 Halolactibacillus sp. 2 65 1 87 Tumebacillus sp. 1 Phenylobacterium sp. 22 Alphaproteobacteria 2 44 Helcobacillus sp. 66 2 88 Tumebacillus sp. 2
175

U2 wk17

T1
wk17

U1 wk17 S1 wk27

Figure 5.6 A non-metric multidimensional scaling (nMDS) plot of OTUs of coal and soil samples. Each treated coal, untreated coal and soil sample was represented by green triangle, blue square dark blue inverted triangle, respectively. A greater dissimilarity was observed among coal samples than soil, as noted by the larger distance area (dashed line) covered by the former. A number of samples (labelled symbols) contributed to most of the differences within each group.

More notable OTUs were observed in coal samples. A particular genus, Phenylobacterium, was observed in all three replicates of the treated coal at week 27 (Figure 5.7). The genus was also found abundant in an untreated coal replicate at week 17. No soil samples at any time point showed a high abundance of this genus, although the genus could be found in very low abundance in soil (data not shown). The high abundance of this genus on coal samples thus shows its preferential attachment and colonisation on coal.

176 40

35 Treated 1 Treated 2 30 Treated 3 25

20

15

OTU abundance (%) 10

5

0

PhenylobacteriumPhenylobacterium sp. 2 sp. 3 Phenylobacterium sp. 4

Figure 5.7 Abundance of OTUs (%) in the treated coal replicates (Treated 1, 2 and 3) at week 27. All replicates showed the presence of Phenylobacterium sp. (arrow and in bold) in different proportions.

177 Furthermore, a group of OTUs from the same genus or family showed dominance in several untreated coal samples. The most prominent example came from the archaea Methanococcoides sp. and bacteria Porphyrobacter sp., which dominated an untreated coal replicate at week 17 (Figure 5.8). The coal sample contained five different Methanococcoides and three Porphyrobacter species, which cumulatively dominated 75% and 19% of the sample’s most abundant population, respectively. These OTUs were not found abundant in soil. Methanococcoides OTUs were the only archaea found on coal with a very low abundance of the genus found in soil (the total archaeal abundance was < 0.01%). The only other archaea found in soil was from the phylum Crenarchaeota at very low abundance.

Methanococcoides Methanococcoides Methanococcoides sp. 3 sp.5 sp. 4 2% 2% Verrumicrobia 2% subdivision 3 sp. Porphyrobacter sp. 3% 3 2% Brevundimonas sp. 3% Porphyrobacter sp. 2 3%

Methanococcoides sp. 2 11% Methanococcoides sp. 1 Porphyrobacter 58% sp. 1 14%

'Untreated 1' Week 17

Figure 5.8 Proportion of the most abundant OTUs in ‘Untreated 1 Week 17’ coal sample, where Methanococcoides sp. and Porphorybacter sp. (in bold) dominated the sample population. ‘Methanococcoides sp. 1’ was present over half of the total most abundant population.

Along with Phenylobacterium sp., the high abundance and frequency of Methanococcoides and Porphyrobacterium genera on coal suggest that they were among the dominant coal colonisers in this study.

178 5.3.3.2 Fungal OTUs

The number of sequences obtained for the 18S rDNA OTUs on coal and in soil samples were ten fold lower than those for archaea and bacteria. In addition, many OTUs were unclassified and/or exhibited very low abundance. Exceptions were samples from the untreated coal at week 17 that showed a relatively high abundance of OTUs (Figure 5.9). One sample was heavily dominated by Philoporus sp., which constituted 99% of the total sample population. The genus was also found in other samples including soil, although at a much lesser proportion. The remaining 1% of the population on the coal sample was Blakeslea sp. Another untreated coal replicate showed similarly high abundance of OTUs, however, these OTUs were unclassified.

Figure 5.9 Fungal OTUs of coal and soil samples at week 17 and 27. Samples were labelled by the first letter of its name, week and replicate (e.g. treated coal replicate 1, untreated coal replicate 1 and soil replicate 1 at week 17 are T17_1, U17_1 and S17_1, respectively). Each colour denotes an OTU. Both replicates of the untreated coal at week 17 exhibited relatively higher abundance than other samples. The untreated coal replicate 2 specifically showed a very high proportion (99%) of Pilophorus sp. The most abundant OTUs in the untreated coal replicate 1, however, consisted of unclassified eukaryotic OTUs.

179 5.3.4 Scanning Electron Microscopy of biomass on coal

Untreated and treated coal samples exposed to soil were visualised through SEM at the end of Week 27. Prior to imaging, the coal pieces were briefly vortexed and rinsed in PBS medium to remove non-adhered particles from the coal surface.

In general, aggregated soil particles of varying sizes could be seen attaching on the coal surface (Figures 5.10-5.11). Whilst there were soil aggregates adhering to smoother areas of coal, other aggregates could be seen attaching on areas that were more hollow and concave. There was a noticeably larger surface area covered with soil particles on the untreated coal compared to the treated coal. These aggregates were also larger in size (maximum approximately 2 x 3 mm) than on the treated coal (max. approx. 0.1 x 0.12 mm).

Figure 5.10 Soil aggregate attachment (arrows) on the untreated coal.

180

A B

Figure 5.11 A) Soil aggregate attachment on the treated coal (arrow). B) A blown up image of a soil aggregate in (A) (dashed box), which consisted of complex and heterogeneous material.

The typical appearance of biofilms (i.e. cell aggregates with extrapolymeric substances) was not apparent on coal. The soil aggregates in particular (e.g. Figure 5.11 B) contained a complex and heterogeneous set of materials that made it challenging to locate and discern microbial cells from inorganic components of soil (e.g. salts and minerals). However, certain entities that were characteristic of filamentous fungal and bacterial cells (width of approximately 1-2 μm) could be located either directly on the coal surface or soil aggregates (Figure 5.12). Spore-like structures (approximately 10 x 20 μm or 4 x 2.5 μm) could also be observed.

181 A B

C D

E F

Figure 5.12 Microbial-looking entities (arrows and/or dashed boxes) observed on coal and across the soil aggregates: Filamentous fungal hyphae on a soil aggregate (A) and coal surface (B); elongated bacterial cell on coal (C, zoomed image D) and spores in the soil aggregate (E and F).

182 5.4 Discussion

This chapter investigated mixed-community microbial attachment on coal from a molecular ecological perspective. Soil obtained from the Lithgow State Coal Mine was chosen as an environment to which coal was exposed. The community that adhered to coal was compared against that in soil to observe if there were any differences in the community composition, and also if there were any unique populations that utilised coal as a substratum for attachment. Terminal restriction fragment length polymorphism (T-RFLP) and pyrosequencing techniques were used to achieve these aims.

Based on the T-RFLP analysis, the overall community composition on both the untreated and treated coal was found to be significantly different (P=0.001) from that in soil (Figure 5.2). Significant differences between coal and soil communities were observed until the end of incubation (week 27) despite the soil and coal communities became more similar over time. These differences were also shown to be significant in the pyrosequencing analysis of coal and soil samples at weeks 17 and 27 (Figures 5.5 and 5.6).

Differences in community composition between coal and soil were more apparent at earlier time points (weeks 0-10). Here, a low average similarity in community composition was also observed between the coal replicates (Figure 5.3). It was clear, however, that as time progressed, community composition between coal and soil became more similar and the average similarity between replicates increased. This highly coincided with the amount of DNA on coal samples that also increased over time (Figure 5.1). The log (x+1) data transformation was applied to favour the statistical analysis based on community composition (presence of T-RFs) rather than abundance (intensity of T-RFs), but it was not completely successful due to a large difference in the amount of DNA between coal and soil. Hence, the significant difference in composition observed between coal and soil at weeks 0 – 10 were probably influenced by abundance rather than composition. This problem may be have been alleviated by dilution of soil extracts to match coal extract concentrations, however this approach was not taken to avoid loss of information. Nonetheless, the information obtained from
183 extracts at later incubation times (where DNA was more abundant) is statistically sound.

The lack of DNA on coal samples at early incubation implied scarcity in biomass found on the coal surface. This shows that the number of attached cells on the coal surface was low, since non-adhered particles were removed from the coal surface prior to DNA extraction. Although the DNA amount was undetectable at weeks 0 – 10, the increasing similarity between replicates indirectly indicated that the amount gradually increased over time. Thus, this implies that the number of cells attached to coal increased over time and became detectable through DNA quantification by week 17. Given this observation, the rate for cell attachment on coal was shown to be extremely low in contrast to other biofilm studies where cells were observed to form biofilms within 24 hours to one week of incubation (Agladze et al. 2003; Burmolle et al. 2007; Hinsa and O'Toole 2006; Tolker- Nielsen et al. 2000; Webb et al. 2003).

Many biofilm studies have been conducted under conditions that favour growth and biofilm formation. One such condition is nutrient availability, where cells are given adequate (accessible) carbon and nutrient sources in a medium to grow in large numbers. This is in line with the viewpoint of Costerton et al. (1995), where biofilm formation would often occur in carbon- and nutrient-rich media that allow cells to be more metabolically active. As excess nutrients were not available in this study, soil microbes had to depend on the sources present in the soil (e.g. humic acids, N and P). The absence in excess nutrients may have created an environment where extensive and rapid biofilm formation on coal surfaces was not favoured, thus resulting in comparatively low number of cells attaching to coal over time.

There was also an apparent difference in the attachment of microbes on the untreated and treated coal, as proven by the lower amount of DNA and soil aggregation on the latter (Figure 5.11). This may have also contributed to a significant difference in community composition on the treated coal and in soil at week 27, as shown by its lower abundance than that on untreated coal. This difference may reflect the different surface properties of both coal types and the
184 free energy of adhesion ΔGAdh between coal and cells as previously described (Chapter 2 and 3). Whilst both coal types were thermodynamically favourable for attachment by hydrophobic fungi, adhesion was only favourable on the untreated coal by hydrophilic bacteria. Thus, although cell attachment occurred on the treated coal, it was likely that the adhered cells took more energy (hence time) to overcome the energy barrier for cell attachment. Hence, a ‘time lag’ was observed in the treated coal attachment compared to the untreated coal.

The pyrosequencing data revealed a high frequency of unique abundant OTUs on coal across replicates and different time points (Figures 5.5-5.6). This was in contrast to soil, where a large number (70%) of abundant OTUs were shared between the soil samples at both time points. The reason for the high variability of OTUs among the coal samples is unclear. Since high reproducibility of OTUs was shown in the soil samples, the high variability in coal samples was not likely due to errors in the pyrosequencing reactions. The pyrosequencing results were also processed appropriately to ensure only high quality sequences were used.

However, depending on the coal type, different proportions of coal macerals, i.e. vitrinite, liptinite and inertinite, can be present. Hence, the varying components in coal itself could potentially attract different group of microbes for attachment, leading to the variability observed in microbial composition. Another reason that may have contributed to the high variability was the spatially varying proportion of the lower abundant microbes in soil. The majority of the dominant soil microbes were relatively stable and homogeneous, however, this may not necessarily be true for soil microbes of very low abundance that were found on coal. This creates a sub-heterogeneity environment in the otherwise homogenous soil, where exposure of these low abundant microbes to coal would depend on random chance. Because each coal sample was placed at varying locations and depths (± 2 cm), differences in the presence and/or abundance of the low abundant microbes in soil at a given microenvironment likely contributed to the high variability of OTUs among coal samples.

Despite the variable outcomes, pyrosequencing analyses revealed several microbes that dominated colonisation on coal. These include members of
185 Phenylobacterium, Methanococcoides and Porphyrobacter genera, which were found on coal samples in high frequency and abundance (Figures 5.7-5.8).

Phenylobacterium species are strict aerobes belonging to the alphaproteobacteria class commonly isolated from soil (Eberspächer and Lingens 2006; Islam et al. 2011; Jakobs-Schönwandt et al. 2010; Sánchez-Peinado et al. 2010; Tiago et al. 2005). Interestingly, previous studies have linked this species with degradation of several compounds including Chloridazon, a known herbicide (Mayer et al. 1985; Muller et al. 1982), N-cyclohexyldiazenium dioxide (HDO) (Jakobs-Schönwandt et al. 2010), p-nitrophenyl butyrate (Lee et al. 2010) and linear alkylbenzene sulfonate (Sánchez-Peinado et al. 2010). The examples show a strong association between Phenylobacterium and the biodegradation of aromatic compounds. Thus, coal, which also contains a high amount of aromatic compounds, may have been chemically attractive to a number of Phenylobacterium species that resulted in their dominance on coal. Further, because this genus was more frequently found in the treated than untreated coal, it was likely that the Phenylobacterium group were attracted to nitric acid-treated coal compounds such as nitrophenols (Kinney and Ockert 1956).

Interestingly, some individual coal samples were heavily dominated by a specific OTU. This was not observed in many other samples, including soil. The genera Methanococcoides and Porphyrobacter on the ‘Untreated 1’ coal replicate at week 17 (Figure 5.8) were found at high frequency and abundance, which suggests a link between these microbes and coal. The dominance and/or presence of these specific groups were not observed on other coal replicates, which, as discussed earlier, was likely due to the maceral selective pressure or spatial sub-heterogeneity in soil.

Methanococcoides spp. are methylotrophic methanogens that have been previously isolated from anaerobic water sediments in cold temperatures (> 0 oC) (Franzmann et al. 1992; Singh et al. 2005; Sowers and Ferry 1983). Despite the ability to grow in cold environments, they were found to grow optimally at warmer temperatures (24 – 30 oC), which is consistent with the more recent findings of Methanococcoides species in a tropical mangrove (Lyimo et al. 2009) and mud
186 volcano (Lazar et al. 2011). Unlike hydrogenotrophic and acetoclastic methanogens, Methanococcoides species do not depend on hydrogen, carbon dioxide or acetate as substrates for methanogenesis; instead, methylamines or methanol are preferred (Franzmann et al. 1992; Singh et al. 2005; Sowers and Ferry 1983). The latter is the less dominant pathway, accounting for approximately 5-10% of methanogenesis in the environment (Penger et al. 2012). This is true in the case of methanogenesis on coal beds, where the majority of methanogens that have been found are hydrogenotrophic and acetoclastic (Games et al. 1978; Li et al. 2008; Penner et al. 2010; Strąpoć et al. 2011b; Strąpoć et al. 2011a). Recently, however, a few studies have shown that methylotrophic methanogenesis occurs in coal seams worldwide, indicated by several factors including a large presence of known methylotrophic methanogens (e.g. Methanolobus sp.) (Doerfert et al. 2009; Guo et al. 2012; Shimizu et al. 2007a; Wawrik et al. 2012). Thus, although Methanococcoides has yet to be identified as one of the key players in coal bed methane, the high abundance of this genus on coal in this study is potentially indicative of its association with coal through methylotrophic methanogenesis.

Studies on methylotrophic methanogenesis in coal beds suggested that the presence of syntrophic bacteria was likely the main reason for the degradation of coal to simpler substrates, such as aliphatic and aromatic monomers, butyrate, alcohols and methanol, which act as precursors for methylotrophic methanogenesis. Bacterial biodegradation of lignin components in coal, in particular, would result in production of methanol through the transformation of methoxylated aromatic compounds, which are building blocks of lignin (Donnelly and Dagley 1980; Schink and Zeikus 1982). Thus, it is highly probable that a proportion of the bacteria found on coal, alongside Methanococcoides, acted in syntrophy by degrading coal and providing the by-products as substrates for methanogenesis.

One example of synthrophic bacteria was Porphorybacterium spp., which was highly abundant on the Methanococcoides-containing coal sample. These bacteria have been known to degrade a number of organic and aromatic compounds, including biphenyls, dibenzofuran (Hiraishi et al. 2002) and high
187 molecular weight polyaromatic hydrocarbons (PAHs) such as naphthalene and pyrene (Gauthier et al. 2003), all of which are naturally found in coal. These bacteria are also phylogenetically closely linked to Sphingomonas spp., which are known to degrade a plethora of aromatic compounds, including PAHs (Eriksson et al. 2003; Kästner et al. 1998; Viñas et al. 2005), biphenyls (Peng et al. 1998; Pieper 2005), phenols (McAllister et al. 1996; Tanghe et al. 1999), azo dyes (Keck et al. 1997; Rau et al. 2002) and dioxins (Wittich et al. 1992). Thus, this suggests that Porphyrobacter spp. could potentially degrade components in coal with methanol as a by-product that can feed into methanogenesis, which have been previously suggested by Strapoć et al. 2010, 2011, 2011b). Interestingly, however, Porphyrobacter spp. are strictly aerobic, whereas Methanococcoides spp. are strict anaerobes. This suggests that the two groups were spatially segregated, or that the bacteria protected the methanogens from oxygen by consuming the latter. Oxic/Anoxic interphases may have also been present in this microniche, which was supported by the presence and abundance of facultative anaerobic bacteria such as Brevundimonas sp.. The potential association between these bacteria and archaea provide an interesting basis for further research.

Other groups that showed high abundance on coal include ‘Acidobacterium sp. Gp1-3’ and ‘Peptococcaceae_1 on the treated coal, Streptococcus sp. and ‘Rhizomicrobium sp. 1’ on the untreated coal, and ‘Rhizomicrobium sp. 2’, Solirubrobacter sp. and Staphylococcus sp. on both coal types (Figure 5.5). Except for Streptococcus and Rhizomicrobium, all the genera or phyla above have been associated with coal through direct isolation (Staphylococcus sp. (Pokorny et al. 2005), Acidobacterium sp. (Brofft et al. 2002)) or discovered as part of a coal bed methane community (Peptococcaceae (Shimizu et al. 2007b), Solirubrobacter sp. (Tang et al. 2012)). The lack of abundance of these species in soil, and their repetitive presence and abundance on coal suggests that they may have benefited from coal to some extent. These species may have used coal as substratum for attachment and/or as a carbon source. This was not tested directly, however the inclusion of inert substratum controls similar to coal in shape and topography (e.g. small rocks) may be useful in future experiments

188 to distinguish species that would use coal not only for attachment but also as substrate for growth.

The pyrosequencing results did not provide sufficient information to discern most of the fungal OTUs obtained in each sample (Figure 5.9). Whilst sequence data was obtained, most of the OTUs were very low in abundance and could not be classified accurately. The universal primers used here targeted bacteria, archaea and eukaryotes. However, in this case, the sensitivity for fungal sequences was rather low. Further studies on similar samples may require fungal specific (e.g. 18S or ITS) primers instead to have a clearer picture of fungal populations in the coal/soil interphase. Nonetheless, one fungal species, Pilophorus sp., was abundant in one of the untreated coal replicates at week 17. Pilophorus spp. are fungi that form lichen associations and belong to the Cladoniaceae family of the Ascomycota (Wang et al. 2010; Wedin et al. 2000). There is no previous evidence showing any direct association of this fungus with coal. Further, because of the role it plays as lichen, it is likely that Pilophorus sp. used coal as a substratum for attachment and stability.

The untreated and treated coal pieces were imaged using Scanning Electron Microscopy (SEM) at week 27 to observe the microbial attachment and colonisation on coal surfaces (Figures 5.10-5.12). The coal pieces were vortexed and rinsed in a medium to ensure that only adhered particles were observed in the micrographs. Overall, both coal surfaces contained soil aggregates consisting of complex and heterogeneous particles. The ‘typical’ biofilm appearance consisting of high volume of clustered cells joined together by exopolymeric substances (EPS) (see Figure 2.2 for example) was not observed. This was expected to some extent since no additional nutrients were added in this experimental set up.

Some microbial-looking cells were observed across the coal surface and soil aggregates (Figure 5.12). However, these cells were scarce and did not reflect the high DNA amount and sequencing depth obtained at week 27. Thus, it is likely that most cells existed within the soil aggregates rather than as separate entities. Cells may have formed complex clusters with soil organic and inorganic
189 particles. Although SEM was useful in observing the coal surface at high resolution, it was challenging to identify cells among the aggregates due to the complex and heterogeneous nature of soil. Complementary techniques such as fluorescence in-situ hybridization (FISH) that utilise specific bacterial and fungal probes may be useful to identify cells in aggregates.

In summary, the microbial community analysis on bituminous coal buried in soil showed significant differences between the community adhering to coal and the original soil community, with the former becoming increasingly similar to the latter over time. Cell attachment on coal did not occur rapidly and there was a lag of several weeks before coal was colonised, which may have resulted from low inorganic nutrient availability in the environment and thermodynamic energy barrier from coal. While soil communities were stable over time, coal communities differed markedly. Variation in community composition in coal samples may have resulted from the inherent heterogeneity of coal’s maceral distribution and sub-heterogeneity of low abundant microbes in soil. Nonetheless, over time coal acted as a positive selective force for a group of microorganisms, which include members from the Phenylobacterium, Methanococcoides and Porphyrobacterium genera, where their high abundance and frequency on coal suggest their dominance in bituminous coal colonisation. This study provides a basis for further research into the effects of coal in natural environments and its potential role as a selective pressure for attachment, biofilm formation and potential biodegradation in situ.

190 6 General discussion

Since the discovery of coal-degrading microorganisms was made over the past century, studies on microbial degradation of coal are increasingly of interest to microbiologists today. The fact that coal, a solid and low-bioavailable hydrocarbon, could be modified by certain groups of microorganisms provides various possibilities in the biotechnological applications of coal utilisation. These microbe-driven applications create an alternative to conventional coal processing, which often involves energy-intensive means (e.g. coal combustion) to achieve desired outcomes. Whilst the latter approach is currently necessary to sustain the increasing global energy demand, the negative impacts these methods have on the environment have led to efforts to achieve a more environmentally sustainable utilisation of the world’s natural resources. Indeed, a promising ‘greener’ solution to coal utilisation is the application of coal- degrading microorganisms, which have been demonstrated in several coal processing regimes, including coal desulphurisation and valuable mineral recoveries (Acharya et al. 2001; Ohmura et al. 1993b; Pawlik et al. 2004; Uhl et al. 1989; Vijayalakshmi and Raichur 2002; Vijayalakshmi and Raichur 2003).

A current interest in utilising coal is the capturing of methane from coal beds. Once thought only as a hazard to coal miners, coal bed methane (CBM) is now regarded as a valuable energy resource. Discoveries of biogenic methane from various coal beds worldwide have led to the understanding that microorganisms substantially contribute to the conversion of coal to methane. Different stages of the coal-to-methane bioconversion process have been identified, with the rate- limiting step being the initial fragmentation of the coal macromolecule (see Section 1.2.4 and Figure 1.2). Originally an anaerobic process in situ, the bioconversion of coal to methane can be accelerated via aerobic coal degradation that targets the initial coal fragmentation. Although studies on aerobic coal biodegradation have been ongoing, it is only very recently that its potential application to methane generation of coal is recognised (Haider et al. 2013). Indeed, this creates an exciting platform for the use of aerobic coal
191 biodegradation, which has been successful in liquid fuel generation and land restorations (Hofrichter and Fakoussa 2001).

Studies on the aerobic biodegradation of coal often involve the isolation of novel aerobic coal-degrading microorganisms and/or their possible degradation mechanism, which can be compared to other hydrocarbon degradation studies including that of crude oil. However, only the latter has recognised the importance of cell attachment and biofilm formation to degradation, evident from the various studies conducted (Abbasnezhad et al. 2011; Abbasnezhad et al. 2008; Barathi and Vasudevan 2001; Chakraborty and Mukherji 2010; Johnsen and Karlson 2004; Prabhu and Phale 2003; Rosenberg et al. 1980; Rosenberg and Rosenberg 1981; Rosenberg et al. 1982). This is in stark contrast to aerobic coal biodegradation, where no cell attachment and biofilm studies on coal have been investigated in detail, despite the frequent observations suggesting its importance in coal degradation (Achi and Emeruwa 1993; Cohen and Gabriele 1982; Hofrichter et al. 1997b; Monistrol and Laborda 1994; Mukasa-Mugerwa et al. 2011). Thus, the lack of fundamental knowledge on cell attachment to coal prompted this study to be the first to explore in detail the role of coal as a substratum for cell attachment and colonisation and its link to coal degradation.

6.1 Coal as substratum for cell attachment and colonisation
When viewed as a substratum for cell attachment, coal exhibits unique surface properties that are different from the conventional substrata used in previous biofilm studies (e.g. glass and polystyrene). Coal, in its raw and untreated state, has a relatively more hydrophobic surface (Chapter 2), which is typical for a hydrocarbon and in line with other substances such as crude oil. Substratum surface hydrophobicity is indeed a highly influential factor that attracts cells of varying hydrophobicity to attach and colonise (Chakraborty and Mukherji 2010; Das et al. 2011; Pringle and Fletcher 1983; Webb et al. 1999). This is in agreement with the results obtained in this study (Chapters 2 and 4), where coal, particularly in its untreated state, showed relatively higher hydrophobicity

192 and more favourable microbial cell attachment for both bacteria and fungi. Therefore, coal in its natural state is a suitable substratum for cell attachment, which is advantageous for field applications, where costly and difficult alterations to coal (e.g. oxidising treatment) can be avoided.

Despite the favourable surface characteristics of coal, the degree and total outcome of cell attachment and colonisation on coal depend on other factors, which take into account other physico-chemical, environmental and biological parameters. This creates more complexity in determining whether or not favourable cell attachment on coal would occur.

A significant physico-chemical influence to cell attachment on coal is the acid- base interactions between two surfaces (Chapters 2 and 4). This is in agreement with the van Oss theory (1986), where a substantial portion of the physico-chemical influences in cell-substratum interaction comes from the acid- base (AB) interaction. It was further shown that acid-base interactions have a far greater influence on cell adhesion than van der Waals (LW) forces, particularly when more hydrophobic surfaces are involved (Das et al. 2010). The electron donating and accepting capacities that constitute the AB interaction between two surfaces play an important role, where greater electron give-and-take between coal and cells results in more thermodynamically favourable adhesion (Chapters 2 and 4). Hence, hydrophobicity as well as surface free energy, particularly the electron donating-accepting capacities of both surfaces, represent important aspects of the physico-chemical interactions between cells and coal. As each coal type possesses different hydrophobicity and surface energy properties (Chapter 2), this results in varying cell adhesion strength depending on the coal type (Chapters 2 and 4).

Whilst the untreated coal surface is physico-chemically favourable for cell attachment, mixed results were obtained for treated bituminous coal (Chapters 2 and 4). Treated coal has been considered in previous studies as a more amenable substrate for degradation due to its more oxidised state (Laborda et al. 1999; Machnikowska et al. 2002; Maka et al. 1989; Strandberg and Lewis 1987a; Wondrack et al. 1989). In regard to cell attachment, however, it seems
193 that most of the treated bituminous coal used in this study (i.e. nitric acid- and peroxidase-treated) did not promote favourable attachment for bacterial cells or cells with low hydrophobicity. In fact, delayed colonisation was observed in a natural environment where bacteria predominate (Chapter 5). The thermodynamically unfavourable properties of the nitric acid and peroxidase- treated coal surface were contributed from their markedly reduced hydrophobicity and higher electron donating capacity than untreated coal. These two main properties make the treated coal far less attractive for adhesion by bacterial cells that have similar low hydrophobicity and high electron donating components. On this account, treated bituminous coal is a less suitable substratum for the attachment of bacterial cells than untreated bituminous coal or lignite.

In contrast to bacteria, filamentous fungi showed a thermodynamically favourable attachment to treated coal due to their higher hydrophobicity and lower electron donating capacity or total surface free energy (Chapter 4). Despite the favourable adhesion thermodynamics, certain elements present on the treated coal resulting from chemical modification repelled certain fungi from attaching (Chapters 3 and 4). The repulsion was perhaps due to a negative chemotaxis resulting from potential toxicity of the modified coal surface. Notwithstanding, it is important to acknowledge that certain groups of microbes, including bacteria, are attracted to, or unaffected by, the treated coal surface, which prompts cell adhesion to occur despite the unfavourable adhesion thermodynamics (Chapter 5). These cells may use biological means, e.g. EPS and cell appendages, in order to overcome the physico-chemical barrier and attach to the treated coal surface (Boks et al. 2008; Quirynen and Bollen 1995). The above results show that whilst physico-chemical factors play a significant role in determining initial cell adhesion to coal, other factors may override their dominance.

Favourable cell adhesion on coal is determined by the surface properties not only of coal but also those of cells. It was shown here that higher hydrophobic cell surface promotes stronger cell attachment to coal (Chapter 4), which is in agreement with previous cell attachment studies of other hydrocarbons and
194 surfaces (An and Friedman 1998; Chakraborty and Mukherji 2010; Ding 2009; Van der Mei et al. 1995; Zita and Hermansson 1997). This can be attributed to the characteristics of the cell surface, where hydrophobic components such as hydrophobins, lipopolysaccharides (LPS) and expolymeric substances (EPS) are present (Chau et al. 2009; Smits et al. 2003; Van Loosdrecht et al. 1987b). Thus, higher surface hydrophobicity poses an advantage for cells in adhering to the coal surface. As filamentous fungi are shown to be generally more hydrophobic than bacteria based on contact angle measurements (Chau et al. 2009; Sharma and Rao 2002; Smits et al. 2003), it can be speculated that the high hydrophobicity of filamentous fungal surfaces contributes to the higher frequency of coal degradation found in the literature, which has been largely attributed to the production of coal-degrading substances (e.g. ligninolytic enzymes).
Cell morphology plays another important role in determining the extent of cell colonisation on coal. Coal as a substratum contains various structures resulting from its uneven topography, which promote adhesion of both filamentous and non-filamentous cells. Fine fissures, holes and ridges naturally present on the coal surface may act as ‘refuge’ for non-filamentous cells, i.e. mostly bacteria, to attach into and form aggregates (Chapter 2). Bacterial cell aggregation depends largely on the size of the cell and concavity of the surface, where cells adhere strongly due to higher cell ‘attachment points’ available on the surface (Scardino et al. 2006). Thus, the small and compact cell size characteristic of bacteria may give them an advantage to utilise the rougher structures of coal for cell attachment and biofilm formation. Further work involving the measurement of surface roughness of coal is necessary to determine the extent of its influence on cell attachment.

As shown in this study, filamentous fungi could also utilise the rough structures of coal to substantially colonise the outer and inner coal surface (Chapters 3 and 4). In comparison to bacteria, filamentous fungi could extend their cells (i.e. hyphae) and penetrate deep into the fine fissures of coal, resulting in significant physical alterations to the coal structure. Penetration increases the contact surface area between coal and fungal cells. This could not be easily achieved

195 by non-filamentous cells, which are predominantly bacteria. Although bacteria have some morphological advantages in coal surface colonisation, the ability of filamentous fungi to colonise both the inner and outer coal surface makes them morphologically more dominant as coal colonisers. This, along with the greater surface hydrophobicity in fungi, could partly explain the dominance of filamentous fungi in coal degradation.

An important parameter for substantial and rapid colonisation by both bacteria and fungi on coal is nutrient availability. Cells in a nutrient-rich medium grow considerably more compared to cells in nutrient-limited medium (Chapter 2). The latter is especially true in natural environments, where oligotrophic conditions limit the extent of natural biofilm formation compared to when excess carbon is available (Chapter 5). Although coal is rich in carbon, its low bioavailability and recalcitrance makes it a difficult source to obtain energy. This results in coal, especially the higher ranks (e.g. bituminous coal), acting like an inert substratum, rather than a substratum that is rich in carbon and energy. Nevertheless, various microbial modifications (e.g. surfactant production) by coal-degrading microbes eventually cause changes to the coal surface properties, thereby making it less inert and more carbon-available over time. This prompts further colonisation by microbes due to an increased availability of carbon coming from coal. Thus, since limited carbon in the medium does not cause significant reduction in the initial cell attachment (Chapter 2), and because carbon can be liberated from coal through modification by the attached cells, substantial microbial colonisation on coal is possible without the presence of external carbon source in the medium. However, a considerably longer time is expected to achieve the same extent of colonisation as to when more accessible carbon is present.
The above discussion summarises key findings on the role of coal as substratum for cell attachment obtained from this study. It is evident that physico-chemical factors primarily determine cell attachment to coal. Secondary factors that determine cell attachment to coal are coal, cell and/or environment- specific.

196



6.2 Linking cell attachment and colonisation with degradation of coal
Previous studies have shown the relative importance of cell attachment and colonisation on the degradation of hydrocarbons (García
Junco et al. 2001; Johnsen and Karlson 2004; Rosenberg and Rosenberg 1981; Rosenberg et al. 1982; Wick et al. 2002). In relation to coal degradation, a question thus arises: Is cell attachment and colonisation of coal absolutely necessary for effective coal degradation? Whilst Abbasnezhad et al. (2011) have argued that cell adhesion to a hydrocarbon is indeed critical towards an effective microbial carbon uptake, its dependency relies on several main factors, including hydrocarbon bioavailability and the strength of the microbial degradation system. Indeed, coal is a very low bioavailable substrate for biodegradation, thus requiring microbes to employ various strategies to surpass the bioavailability barrier in order to degrade coal and/or access its carbon. Cell attachment and colonisation on coal can be seen as one of the strategies in overcoming the low bioavailability of coal, which therefore enhances coal degradation (Chapter 4).

However, it occurs that the cell attachment strategy is dependent on the degrading microorganism itself. Assuming that all cells are able attach and colonise surfaces (in physically permissible conditions), attachment and full colonisation on coal is not required by certain microbes (e.g. Phanerochaete chrysosporium) prior to biodegradation (Chapter 3). It is possible that microbes that rely less on coal adhesion possess more efficient extracellular degrading system (e.g. stronger enzymes, surfactants), such that adhesion to coal becomes unnecessary. Environmental factors (e.g. pH, temperature, water availability) further enhance the effectiveness of the degrading system and decrease the need for cell attachment by these microbes.

197 The variability in cell adhesion or the lack of it in coal degradation strengthens the notion that cell adhesion is not an absolute requirement for coal degradation to occur; rather, it acts as one of the mechanisms in coal degradation. Whilst cell attachment may not be completely necessary for coal degradation for all coal-degrading microorganisms, it nonetheless a necessary strategy for many coal degraders to effectively degrade coal. Additional work is required to further demonstrate this claim, including cell-free coal degradation analyses, and genetic manipulation of the adhesion factor in cells and its impact to coal degradation.
6.3 Fusarium oxysporum G9o: A suitable candidate in the application of coal bioconversion to methane?

The application of aerobic coal degrading microorganisms is a promising biotechnology tool towards utilising vast coal resources for methanogenesis. Despite the ongoing research on aerobic coal degradation and coal methanogenesis, the two have been studied as separate research entities; no previous research has attempted to combine the two until very recently (Haider et al. 2013). Field work applications with the aim to enhance in situ methane production from coal reservoirs using native microorganisms still requires substantial research an laboratory testing. A starting point is finding native coal- degrading isolates that are suitable for the application. In this study, isolates obtained in the vicinity of the bituminous coal reserves of interest (Lithgow State Coal Mine, NSW) were screened for coal degradation. Whilst several fungi were able to degrade low-rank coal, only one isolate, Fusarium oxysporum G9o, was able to demonstrate bituminous coal degradation (Chapters 3 and 4).

Several key coal-degrading characteristics were shown by F. oxysporum G9o that are advantageous for in situ application of the isolate. G9o demonstrated preferential colonisation and degradation of untreated bituminous coal (Chapters 3 and 4), which is a major advantage for fieldwork applications, since costly and laborious coal pretreatments can be avoided. Growth on and degradation of coal by G9o occurred successfully at room temperature (22 oC),

198 thus not requiring a warmer condition (28 – 30 oC) usually associated with coal degradation (Achi and Emeruwa 1993; Cohen and Gabriele 1982; Cohen et al. 1990; Gupta et al. 1990; Igbinigie et al. 2008; Laborda et al. 1999). Furthermore, G9o also showed the ability to solubilise bituminous coal both in aerial and aqueous conditions (Chapter 4). These features of G9o are useful in field applications and therefore makes G9o an isolate of commercial interest.

Despite the advantages shown by G9o in coal degradation, several limitations prevail. A major shortcoming is the relatively long duration (i.e. at least 3 months) for observable coal degradation by G9o to occur under room temperature. The delay is a setback to the artificially enhanced coal bioconversion process; thus, there is a clear need to further optimise conditions for more rapid and effective degradation by G9o. Another limitation by G9o degradation is that, like other coal degrading fungi, G9o requires extra nutrients and carbon sources for effective growth on and degradation of coal. Although G9o was not tested for degradation under limiting carbon and nutrient conditions in this study, several tests outside this study have confirmed this limitation. Thus, for successful application of G9o in situ, nutrients need to be administered to meet necessary growth and degradation requirements.

Other parameters also need to be considered when applying G9o to fieldwork. This includes its ability to withstand anaerobic conditions when required (e.g. during transitioning into anaerobic phase) and to cooperate with other microbes that are/may be present in the coal seam. Equally important, the coal fragmented products by G9o need to be suitable for the next phase in coal degradation. The mechanism of coal degradation by G9o is still unknown despite the attempts made in this study (Chapter 4). Elucidating mechanisms would help optimise degradation. Overall, G9o is promising for coal bioconversion, but a considerable amount of work is required before this fungus can be applied to coal conversion to methane.

199 6.4 Future directions
Key findings in this study have unveiled several important observations on the impact of physicochemical, biological and environmental factors on cell adhesion and biofilm formation on coal. As this study is among the first to investigate these matters in the context of aerobic coal degradation, a considerable amount of work in the above three facets is still needed to fully understand the role of cell attachment and colonisation on coal. A few examples of future developments to this study are given below.

In addition to surface hydrophobicity, surface energy and adhesion thermodynamics studied earlier, another physico-chemical factor that needs to be addressed is the impact of coal surface topography on cell adhesion. Surface roughness has been shown to be influential in cell attachment and biofilm formation (Riedewald 2006; Shellenberger and Logan 2002). As mentioned earlier, coal surface is naturally uneven and contains several features contributing to its surface roughness, in addition to its smoother areas. Thus, investigations on the heterogeneity of coal topography may reveal differences in adhesion on the smoother and rougher areas of the coal surface. Atomic Force Microscopy (AFM) can be employed to measure surface roughness of different types of coal and measure the relative adhesion strength of a particular cell along the coal surface. This is performed by moving a cell- adhered cantilever across the coal surface in close proximity to obtain profiling of the substratum topography and cell adhesion strength.

Physico-chemical analyses in this study have revealed possible biological roles in influencing not only the irreversible cell adhesion but also the initial cell attachment to coal. Thus, in conjunction with physico-chemical studies, biological aspects governing cell adhesion on coal need to be further investigated. Both bacteria and fungi produce various adhesins, including exopolymeric substances (EPS), hydrophobins, lipopolysaccharides (LPS), fimbrae and flagella, which have important roles in cell adhesion. Genotypic and phenotypic characterisations of the adhesion factors, including experiments

200 involving knockout mutants may be useful in knowing the relative impact of each adhesion factor on the cell attachment to coal.

Further investigation is needed on the negative chemotaxis behaviour exhibited by certain microbes on coal. Possible toxicity of coal, particularly of the treated coal (but not excluding untreated coal), may be responsible in repelling microbes from attaching to the coal surface. Direct analyses on the effect of coal toxicity on cell adhesion are difficult as coal is chemically heterogeneous; certain components in coal may or may not be toxic to microbes. Therefore, testing the effects of certain coal compounds that may be dominant (e.g. n- nitrophenol in nitric acid-treated coal) in isolation may be useful to determine the extent of the overall coal toxicity.

6.5 Concluding remarks

The microbial cell attachment and colonisation of coal is an important but neglected aspect in previous coal biodegradation studies. Its importance has been acknowledged in the past; however, no single study has been conducted in detail to investigate the fundamental attributes of cell adhesion and biofilm formation on coal. This study explored this aspect of coal microbiology, focusing on several key attributes governing the attachment to and colonisation of coal. Important discoveries on cell adhesion and colonisation on coal were demonstrated, although many other aspects are to be further developed in the future.

Key findings presented in this study will contribute to a new body of knowledge that benefits the biotechnological application of coal utilisation, while other aspects in realising this potential are still underway. More fundamentally, a new research area has been created, which serves as a novel starting point in further understanding the complexity of microbial interactions with coal.

201 References

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Appendices

Appendix I: Media recipes and composition

Sabouraud medium For a 1 L solution add: 800 ml MilliQ water, 10 g peptone, 40 g glucose and 15 g agar (if solid media is required). Mix well and adjust pH to 5.6. Add water to 1 L and autoclave.

Modified minimal 9 (M9) medium (pH 5.6) with trace element solution (TES) and bituminous coal for isolation studies For making 1 L of coal agar, autoclave 15 g of agar in 800 ml of water. After slightly cooled and in constant stirring, add 100 ml of sterile M9 salt solution

(10x), 10 ml of TES (100x), 2 ml of filter sterilised 1 M MgSO4, 100 μl of filter sterilised CaCl2 and 10 g of untreated ground bituminous coal. Mix well and adjust to 1 L. Transfer 25 ml of the mixture into a 90 mm petri dish and cool. For 6x TES, add 60 ml instead of 10 ml TES. For coal silica, substitute agar with 134 g of Na2SiO2.9H20 and follow instructions given in Section 3.2.6.

BIII fungal medium (pH 5.6) (adapted from Kirk et al., 1986) with 6x TES and bituminous coal for isolation studies For making 1 L of coal agar, autoclave 15 g of agar in 600 ml of water. After slightly cooled and in constant stirring, add 100 ml of pre-autoclaved 2 g/L ammonium tartrate, 100 ml of pre-autoclaved 20 g/L KH2PO4, 1 ml of filter- sterilised 5 g/L MgSO4.7H20, 100 μl of filter sterilised CaCl2, 1 ml of 1 g/L thiamine.HCl, 60 ml of TES and 10 g of untreated ground bituminous coal. Mix well and adjust to 1 L. Transfer 25 ml of the mixture into a 90 mm petri dish and cool. For coal silica, substitute agar with 134 g of Na2SiO2.9H20 and follow instructions given in Section 3.2.6.

231 Modified M9 salts (10x) pH 5.6 composition

Content Amount (g/L)

Na2HPO4 0.59

NaH2PO4.H20 13.23 NaCl 5.0

NH4Cl 10.0

Glutamate medium pH 5.6 composition (adapted from Igbinigie et al., 2008)

Content Amount (g/L) Sodium L-glutamate 1.6 K2HPO4 2.5 KH2PO4 10.2 NaNO3 2.4 MgSO4.7H2O 0.4 KCl 0.4

Trace element solution (TES) 100 x composition

Content Amount (mg/L) Nitriloacetic acid 15.0

MgSO4.7H20 30.0

MnSO4.H20 5.0 NaCl 10.0

FeSO4.H20 5.0

CoCl2 1.0

CaCl2 1.0

ZnSO4.7H20 1.0

CuSO4.5H20 0.1

AlK(SO4).12H20 0.8

H3BO3 0.1

NaMoO4.2H20 0.9

232 Appendix II: Primer sequences

Primer target Primer name Sequence 5’-3’ References Fungal ITS1, ITS2 ITS1 TCC GTA GGT GAA CCT GCG G (White et al. 1990) and 5.8S rRNA ITS4 TCC TCC GCT TAT TGA TAT GC regions ITS1-F CTT GGT CAT TTA GAG GAA GTA A (Gardes and Bruns 1993) Fungal ITS1 region ITS1 TCC GTA GGT GAA CCT GCG G (Kumar and Shukla 2005) ITS2 GCT GCG TTC TTC ATC GAT GC A portion of fungal ITS3 GCA TCG ATG AAG AAC GCA GC (Kumar and Shukla 2005) 5.8S and ITS2 ITS4 TCC TCC GCT TAT TGA TAT GC regions 18S region NS1 GTA GTC ATA TGC TTG TCT C (White et al. 1990) NS7 AIC CAT TCA ATC GGT AIT Large subunit region LR0R ACC CGC TGA ACT TAA GC (Klonowska et al. 2003) (LSU) LR16 TTC CAC CCA AAC ACT CG A portion of 16S 27F GAG TTT GAT YMT GGC TC (Frias-Lopez et al. 2002) rRNA region 519R GWA TTA CCG CGG CKG CTG (Hutter et al. 2003) 16S rRNA region 926wF AAA CTY AAA KGA ATT GRC GG (Mason et al. 2012) 1392R ACG GGC GGT GTG TRC (Lane 1991)

233 Appendix III: SEM micrographs of P. fluorescens on coal a) Representative images of cells attaching on bituminous coal (left) and scraped coal (right) that removed approximately 90% of the cells:

b) Representative microcolonies of P. fluorescens on bituminous coal in the absence of external carbon and nitrogen source (i.e. glucose and CAA):

234

235

236

Appendix IV: ATR-FTIR peak-fitting for peroxidase interaction with coal

ATR-FTIR Peak fitting results for peroxidase interaction with coal: (a) 0 mg/mL, (b) 0.0024 mg/mL, (c) 0.024 mg/mL, and (d) 0.24 mg/mL.

a)

b)

237 c)

d)

238

Appendix V: G5o colonisation and degradation of coal

U K

P

T L

U
K

P

T

L

Colonisation and degradation analysis of bituminous and lignite coals by Penicillium sp. G5o. More intense colonisation was observed with Kalimantan (K) (circle outlined) and Loy Yang (L) (box outlined) lignites than the untreated (U) and treated (T) bituminous coals. This resulted in solubilisation of the lignite noted by the brown pigments in the agar directly underneath the coal piece (arrow).
239 Appendix VI: Raw T-RFLP electropherograms (Chapter 5)

Negative sample

Positive sample (Escherichia coli)

240

Soil (untreated coal) week 0 – (Four replicates each week)

241

Soil (untreated coal) week 2

242

Soil (untreated coal) week 4

243

Soil (untreated coal) week 10

244

Soil (untreated coal) week 17

245

Soil (untreated coal) week 27

246

Biomass on untreated coal week 0 – (Four replicates each week)

247

Biomass on untreated coal week 2

248

Biomass on untreated coal week 4

249

Biomass on untreated coal week 10

250

Biomass on untreated coal week 17

251

Biomass on untreated coal week 27

252

Soil (treated coal) week 0 – (Four replicates each week)

253

Soil (treated coal) week 2

254

Soil (treated coal) week 4

255

Soil (treated coal) week 10

256

Soil (treated coal) week 17

257 Soil (treated coal) week 27

258

Biomass on treated coal week 0 – (Four replicates each week)

259

Biomass on treated coal week 2

260

Biomass on treated coal week 4

261

Biomass on treated coal week 10

262

Biomass on treated coal week 17

263

Biomass on treated coal week 27

264 Appendix VII: Raw Operational Taxanomic Unit (OTU) data (sub-sampled) from Chapter 5. T= Treated coal, U=Untreated coal, S= Soil, Wk= Week. Each sample contained at least two replicates (e.g. T1 Wk17, T2 Wk17). Operational Taxonomic T1 T2 U1 U2 S1 S2 S3 T1 T2 T3 U1 U2 U3 S1 S2 S3 Unit (OTU) Wk17 Wk17 Wk17 Wk17 Wk17 Wk17 Wk17 Wk27 Wk27 Wk27 Wk27 Wk27 Wk27 Wk27 Wk27 Wk27 Abiotrophia sp. 0 0 0 0 0 0 0 0 0 0 0 186 0 0 0 0 Acetobacteraceae 0 0 0 0 0 0 0 0 0 0 0 27 0 0 0 0 Acidimicrobiales 1 0 0 0 0 0 0 0 0 0 0 0 0 0 39 0 0 Acidimicrobiales 2 0 0 0 0 0 0 0 0 0 0 0 0 0 27 0 0 Acidobacterium sp. Gp1-1 35 0 0 0 11 0 11 0 0 0 0 0 0 0 11 20 Acidobacterium sp. Gp1-2 0 0 0 0 0 0 0 0 0 0 0 0 0 44 0 0 Acidobacterium sp. Gp1-3 99 0 0 0 0 0 0 41 94 0 0 0 0 0 0 0 Acidobacterium sp. Gp1-4 0 0 0 0 0 10 0 0 0 0 0 0 0 0 0 0 Acidobacterium sp. Gp1-5 0 0 0 0 0 0 0 0 0 0 0 10 0 0 0 0 Acidobacterium sp. Gp2-1 0 0 0 0 11 10 0 0 0 0 0 13 0 0 0 0 Acidobacterium sp. Gp2-2 0 0 0 0 12 0 8 0 0 0 0 0 0 0 0 21 Acidobacterium sp. Gp2-3 0 0 0 0 0 0 0 16 0 0 0 0 0 0 0 0 Acidobacterium sp. Gp6 0 0 0 0 0 0 0 0 0 0 0 0 0 0 12 0 Acidobacterium sp. Gp10 0 0 0 0 0 0 0 0 0 0 16 0 0 0 0 0 Acidovorax sp. 56 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Actinobacteria 1 0 0 0 16 0 0 0 0 0 0 0 0 0 0 0 0 Actinobacteria 2 0 0 0 0 0 0 0 0 0 0 14 0 0 0 0 0 Actinomycetales 1 0 20 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Actinomycetales 2 0 0 0 0 0 0 0 0 0 0 0 0 0 27 0 0 Actinomycetales 3 0 0 0 0 0 0 0 0 0 203 0 0 0 0 0 0 Alphaproteobacteria 1 17 18 0 0 9 14 17 0 0 0 0 0 0 37 11 19 Alphaproteobacteria 2 65 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Alphaproteobacteria 3 0 0 0 0 0 0 0 0 0 0 0 45 0 0 0 0 Alphaproteobacteria 4 0 0 0 0 0 12 0 0 0 0 0 0 0 0 0 0 Bacillales 1 0 0 0 0 0 0 0 0 0 0 0 0 31 0 0 0 Bacillales 2 0 0 0 0 0 0 0 0 0 0 0 0 22 0 0 0 Bacillales 3 0 0 0 0 0 0 0 0 0 0 0 0 14 0 0 0 Bacillales 4 0 16 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Bacillus sp. 166 0 0 11 0 0 0 0 0 0 0 0 0 0 0 0 Betaproteobacteria 0 0 0 0 0 0 0 0 53 0 0 0 0 0 0 0 Bradyrhizobium sp. 1 0 0 0 0 0 14 0 0 0 0 0 0 0 0 0 0 Bradyrhizobium sp. 2 0 0 0 0 0 0 10 0 0 0 0 0 0 0 0 9 Brevundimonas sp. 0 0 17 0 0 0 0 0 0 0 0 0 0 0 0 0 Burkholderia sp. 0 0 0 0 14 13 9 0 0 0 0 0 0 0 12 13 Caldilinea 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 9

265 Chitinophagaceae 97 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Cloacibacterium sp. 0 60 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Gammaproteobacteria 1 0 0 0 0 30 19 33 0 0 87 0 0 0 0 45 0 Gammaproteobacteria 2 0 0 0 0 25 19 18 0 0 0 0 0 0 0 32 49 Gammaproteobacteria 3 0 0 0 10 0 0 0 0 0 0 0 0 0 0 0 0 Gammaproteobacteria 4 0 0 0 0 0 0 0 0 0 0 0 0 11 0 0 0 Halolactibacillus sp. 1 0 0 0 0 0 0 0 0 0 0 0 0 49 0 0 0 Halolactibacillus sp. 2 0 0 0 0 0 0 0 0 0 0 0 0 21 0 0 0 Helcobacillus sp. 0 0 0 0 0 0 0 0 0 0 0 40 0 0 0 0 Hymenobacter sp. 0 0 0 11 0 0 0 0 0 0 0 0 0 0 0 0 Kitasatospora sp. 0 0 0 0 0 0 0 0 40 0 0 0 0 0 0 0 Ktedonobacter sp. 0 0 0 0 0 0 0 16 0 0 0 0 0 0 0 0 Ktedonobacterales 0 0 0 0 0 0 0 34 0 0 0 0 0 0 0 0 Methanococcoides sp. 1 0 0 345 0 0 0 0 0 0 0 0 0 0 0 0 0 Methanococcoides sp. 2 0 0 66 0 0 0 0 0 0 0 0 0 0 0 0 0 Methanococcoides sp. 3 0 0 13 0 0 0 0 0 0 0 0 0 0 0 0 0 Methanococcoides sp. 4 0 0 14 0 0 0 0 0 0 0 0 0 0 0 0 0 Methanococcoides sp. 5 0 0 13 0 0 0 0 0 0 0 0 0 0 0 0 0 Microbacteriaceae 0 0 0 0 0 0 0 48 0 0 0 0 0 0 0 0 Microbacterium sp. 0 15 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Nocardioidaceae 1 0 0 0 0 16 13 10 0 0 0 0 0 0 0 23 22 Nocardioidaceae 2 0 0 0 0 0 0 0 0 0 0 43 0 0 0 0 0 OP11 sp. 0 0 0 0 0 0 0 0 0 47 0 0 0 0 0 0 Oxalobacteraceae 1 0 0 0 0 0 0 0 0 0 14 0 0 0 0 0 0 Oxalobacteraceae 2 0 0 0 0 0 0 0 0 0 0 0 0 15 0 13 0 Paenibacillaceae_1 0 0 0 0 0 0 0 0 0 0 0 0 0 32 0 0 Pasteuria sp. 0 0 0 0 0 0 0 0 0 0 42 0 0 0 14 9 Peptococcaceae_1 0 0 0 0 0 0 0 0 94 21 0 0 0 0 0 0 Petrimonas sp. 0 0 0 0 0 0 0 63 0 0 0 0 0 0 0 0 Phenylobacterium sp. 1 0 0 0 27 0 0 0 0 0 0 0 0 0 0 0 0 Phenylobacterium sp. 2 0 0 0 0 0 0 0 199 0 0 0 0 0 0 0 0 Phenylobacterium sp. 3 0 0 0 0 0 0 0 60 40 0 0 0 0 0 0 0 Phenylobacterium sp. 4 0 0 0 0 0 0 0 0 0 13 0 0 0 0 0 0 Planctomycetaceae 0 0 0 14 0 0 0 0 0 0 0 0 13 0 0 0 Porphyrobacter sp. 1 0 0 82 0 0 0 0 0 0 0 0 0 0 0 0 0 Porphyrobacter sp. 2 0 0 20 0 0 0 0 0 0 0 0 0 0 0 0 0 Porphyrobacter sp. 3 0 0 9 0 0 0 0 0 0 0 0 0 0 0 0 0 Propionibacterium sp. 0 0 0 57 0 0 0 60 0 0 0 0 0 0 0 0 Pseudolabrys sp. 0 0 0 0 0 0 0 0 0 40 40 0 0 30 13 0 Pseudomonas sp. 0 0 0 0 0 0 0 0 38 0 0 0 0 0 0 0 Rheinheimera sp. 0 59 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Rhizobiales 1 0 0 0 0 10 14 12 0 0 0 0 0 0 0 0 0 Rhizobiales 2 0 0 0 0 0 0 0 15 0 0 0 0 0 0 0 0 Rhizomicrobium sp. 1 0 0 0 0 0 0 0 0 0 0 43 12 0 0 0 0
266 Rhizomicrobium sp. 2 0 0 0 8 0 0 0 0 71 0 90 0 0 0 0 0 Sinobacteraceae 0 0 0 0 0 0 0 0 0 0 0 0 0 50 0 0 Solirubrobacter sp. 0 0 0 0 0 0 0 0 54 97 14 0 0 0 0 0 Staphylococcus sp. 0 0 0 0 0 0 0 0 40 0 0 194 21 0 0 0 Streptococcus sp. 0 0 0 0 0 0 0 0 0 0 0 137 11 0 0 0 Tepidimonas sp. 0 62 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Thermomonosporaceae 0 0 0 8 0 0 0 0 0 0 0 0 0 0 0 0 Tumebacillus sp. 1 0 24 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Tumebacillus sp. 2 0 0 0 0 0 0 0 0 0 0 16 0 0 0 0 0 Unclassified bacteria 1 0 33 0 0 20 0 9 0 0 0 0 0 0 27 0 0 Unclassified bacteria 2 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 9 Unclassified bacteria 3 0 0 0 0 0 0 0 0 0 0 0 0 0 38 0 0 Unclassified bacteria 4 75 0 0 0 0 0 0 0 0 0 0 0 0 24 0 0 Unclassified bacteria 5 20 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Unclassified bacteria 6 49 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Unclassified bacteria 7 0 20 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Unclassified bacteria 8 0 0 0 28 0 0 0 0 0 0 0 0 0 0 0 0 Unclassified bacteria 9 0 0 0 0 0 0 0 0 68 0 0 0 0 0 0 0 Unclassified bacteria 10 0 0 0 0 0 0 0 0 0 0 20 0 0 0 0 0 Variovorax sp. 0 0 0 0 0 0 0 0 0 30 0 0 0 0 0 0 Verrumicrobia subdivision 0 0 15 0 0 0 0 0 0 0 0 0 0 0 0 0 Xanthomonadaceae 0 0 0 0 0 0 0 0 0 61 0 0 0 0 0 0

267