Streptomyces as a Source of Geosmin and 2-methylisoborneol Associated Taste and Odour Episodes in Drinking Water Reservoirs
Elise Anne Asquith BEnvScMgt (Hons)
A thesis submitted to the University of Newcastle, Australia, in fulfilment of requirements for admission to the degree of Doctor of Philosophy
February 2015
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DECLARATION
The thesis contains no material which has been accepted for the award of any other degree or diploma in any university or other tertiary institution and, to the best of my knowledge and belief, contains no material previously published or written by another person, except where due reference has been made in the text. I give consent to the final version of my thesis being made available worldwide when deposited in the University’s Digital
Repository, subject to the provisions of the Copyright Act 1968.
………………………………….. Elise Anne Asquith
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ACKNOWLEDGEMENTS There are a number of individuals who have been of immense support during my PhD candidature who I wish to acknowledge. It has been a challenging and enduring experience, but the end result has to be recognised as a great sense of academic achievement and personal gratification.
I would like to express my deep appreciation and gratitude to my supervisors. Dr Craig Evans has undoubtedly been the most important person guiding my research over the past three years and has been a tremendous mentor for me. I am truly grateful for his advice, patience and support. In particular, I wish to thank him for accompanying me on all of my visits to Grahamstown and Chichester Reservoirs and generously dedicating much time to reviewing my thesis. Thanks also goes to my other supervisors. To Associate Professor Phillip Geary for his support and assistance in the revision of the thesis and Professor Hugh Dunstan who provided brilliant and novel ideas which have been of immense importance in directing much of this research.
Hunter Water Corporation provided the financial support for conducting this research and I especially would like to thank Mr Bruce Cole for proposing the interesting research topic of this thesis. I also would like to acknowledge The University of Newcastle, Australia, for providing me with a Postgraduate Research Scholarship.
I wish to thank all the staff and students associated with the Dunstan-Roberts Laboratory. Dr Margaret MacDonald for her continuous encouragement, Mr Tony Rothkirch for his technical assistance and Nicole, Marcus, and Mousa for their friendship and support during my PhD. I am also very appreciative of the generosity of researchers from the Reproductive Science Laboratory, especially Dr Matthew Jobbling and Dr Ben Curry for their assistance in the molecular analyses conducted in this research.
Luke, my loving husband, needs to be recognised as the most significant individual who has been patient and understanding, has supported me and given me continuous encouragement to complete my PhD. I also would like to acknowledge the important role of my parents, Angela and Ralph in getting me to this level of my education. They have given me every opportunity in life and have always encouraged me to reach my fullest potential. Words cannot express how grateful I am to them. Of course my siblings Nivi, Melissa and Joe deserve a mention and I thank them for pretending to show interest in my research over the years. Thanks goes to my canine companion Cooper who has sat loyally and patiently by my side during the writing of this thesis, keeping me company and taking me for much needed runs to clear my mind. II
“An understanding of the natural world and what's in it is a source of not only a great curiosity, but great fulfillment”
Sir David Attenborough
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TABLE OF CONTENTS
DECLARATION ...... I ACKNOWLEDGEMENTS ...... II TABLE OF CONTENTS ...... IV LIST OF FIGURES ...... XI LIST OF TABLES ...... XVIII ABBREVIATIONS ...... XIX ABSTRACT ...... XXI PUBLICATIONS ...... XXIV
CHAPTER 1 - INTRODUCTION ...... 1
CHAPTER 2 - STREPTOMYCES AND THE OCCURRENCE OF GEOSMIN AND 2-METHYLISOBORNEOL IN AQUATIC ENVIRONMENTS ...... 7 2.1 Introduction ...... 7
2.2 Actinobacteria and the genus Streptomyces: What are they? ...... 8
2.3 The ecological significance of actinobacteria ...... 10
2.4 Geosmin and 2-MIB in drinking water supplies ...... 11
2.4.1 Non-actinobacterial producers of geosmin and 2-MIB ...... 13
2.4.2 Geosmin and 2-MIB production by Streptomyces and other actinobacteria ...... 14
2.4.2.1 Conditions of production ...... 15
2.4.2.2 Pathways of biosynthesis ...... 18
2.4.2.3 Purpose of biosynthesis ...... 21
2.5 A terrestrial origin of Streptomyces and T&O compounds? ...... 22
2.6 Streptomyces and T&O production in aquatic environments ...... 23
2.6.1 The water mass ...... 27
2.6.2 Vegetation ...... 28
2.6.3 Sediments and muds ...... 29
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2.6.4 Other aquatic habitats ...... 31
2.7 Conclusions ...... 33
CHAPTER 3 - EXPLORATORY MULTIVARIATE MODELLING OF PAST GEOSMIN AND 2-METHYLISOBORNEOL EPISODES IN GRAHAMSTOWN AND CHICHESTER DRINKING WATER RESERVOIRS ...... 35 3.1 Introduction ...... 35
3.2 Methods ...... 39
3.2.1 Description of the drinking water reservoirs ...... 39
3.2.2 Water quality data sets ...... 41
3.2.3 Statistical analysis ...... 43
3.3 Results ...... 44
3.3.1 Overview of PLS models ...... 44
3.3.2 PLS models for geosmin in Grahamstown Reservoir ...... 47
3.3.3 PLS models for 2-MIB in Grahamstown Reservoir ...... 53
3.3.4 PLS models for geosmin in Chichester Reservoir ...... 56
3.4 Discussion ...... 62
3.5 Conclusions ...... 69
CHAPTER 4 - STREPTOMYCES IN DRINKING WATER RESERVOIRS: DISTRIBUTION, ACTIVITY AND POTENTIAL CONTRIBUTION TO TASTE AND ODOUR EPISODES INVOLVING GEOSMIN AND 2-METHYLISOBORNEOL ...... 71 4.1 Introduction ...... 71
4.2 Materials and methods ...... 79
4.2.1 Sampling sites and collection ...... 79
4.2.2 Extraction of DNA from environmental samples ...... 84
4.2.3 Quantitative polymerase chain reaction (qPCR) protocol ...... 87 V
4.2.4 Enumeration and isolation of Streptomyces...... 91
4.2.5 Identification of Streptomyces-like isolates ...... 96
4.2.5.1 Extraction of DNA from Streptomyces-like isolates ...... 97
4.2.5.2 Amplification of Streptomyces-specific 16S rRNA sequences ...... 97
4.2.5.3 Sequencing of Streptomyces-like isolates ...... 97
4.2.5.4 Phylogenetic analysis ...... 98
4.2.6 Geosmin and 2-MIB production by Streptomyces-like isolates ...... 99
4.2.7 Production of geosmin and 2-MIB by Streptomyces on environmental substrates ...... 103
4.2.8 Data analysis ...... 106
4.3 Results ...... 107
4.3.1 Efficiency of cell lysis methods to differentiate between Streptomyces vegetative cells and spores ...... 107
4.3.2 Evaluation of the qPCR assay ...... 110
4.3.3 Detection and quantification of Streptomyces in environmental samples ...... 112
4.3.3.1 Streptomyces abundance and activity in water samples ...... 112
4.3.3.2 Streptomyces abundance and activity in solid environmental substrates ... 116
4.3.4 Enumeration of Streptomyces on selective growth media and a comparison with qPCR-determined abundance ...... 122
4.3.5 Streptomyces-like isolates: identification and ability to produce geosmin and 2-MIB...... 124
4.3.6 Geosmin and 2-MIB production by Streptomyces on environmental substrates ...... 132
4.3.7 Summary of key findings ...... 134
4.4 Discussion ...... 135
4.4.1 The potential of Streptomyces to contribute to T&O ...... 135
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4.4.2 The ‘wash-in’ of Streptomyces and their T&O secondary metabolites from marginal environments into drinking water supplies ...... 136
4.4.3 The potential for in situ production of T&O secondary metabolites by Streptomyces in the water mass ...... 145
4.4.4 Molecular versus culture-dependent detection and quantification of Streptomyces ...... 150
4.4.5 Streptomyces isolates: identification and geosmin and 2-MIB producing ability ...... 152
4.5 Conclusions ...... 154
CHAPTER 5 - STREPTOMYCES IN THE MARGINAL SEDIMENTS OF DRINKING WATER RESERVOIRS: A SIGNIFICANT SOURCE OF GEOSMIN AND 2-METHYLISOBORNEOL DURING WATER LEVEL RECESSION? ...... 158 5.1 Introduction ...... 158
5.2 Materials and methods ...... 160
5.2.1 Microorganism ...... 160
5.2.2 Experimental design ...... 161
5.2.3 Dissolved oxygen measurements ...... 163
5.2.4 Geosmin and 2-MIB measurements ...... 163
5.2.5 Quantification of Streptomyces in sediment ...... 164
5.2.6 Statistical analyses ...... 165
5.3 Results ...... 166
5.3.1 Geosmin and 2-MIB production in exposed and submerged sediment ...... 166
5.3.2 Dynamics of Streptomyces population densities in exposed and submerged sediment ...... 169
5.3.3 Dissolved oxygen concentration at the surface of submerged sediment ...... 170
5.4 Discussion ...... 171
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5.4.1 Geosmin and 2-MIB production by Streptomyces in marginal sediments: a comparison between exposed and submerged conditions ...... 171
5.4.2 Geosmin and 2-MIB production by Streptomyces under anaerobic conditions in submerged sediments ...... 177
5.5 Conclusions ...... 180
CHAPTER 6 -MORPHOLOGICAL DIFFERENTIATION AND THE INFLUENCE OF ENVIRONMENTAL FACTORS ON GEOSMIN AND 2-METHYLISOBORNEOL PRODUCTION BY STREPTOMYCES SPP...... 182 6.1 Introduction ...... 182
6.2 Materials and methods ...... 186
6.2.1 Microorganisms ...... 186
6.2.2 Full factorial experimental design ...... 187
6.2.3 Media composition and culture conditions ...... 192
6.2.3.1 Experiment 1: The effect of carbon, nitrogen and phosphorous concentration ...... 192
6.2.3.2 Experiment 2: The effect of temperature, NaCl concentration and pH ..... 193
6.2.3.3 Experiment 3: The effect of calcium, magnesium and potassium concentration ...... 193
6.2.3.4 Experiment 4: The effect of iron, zinc, copper and manganese concentration ...... 194
6.2.4 Biomass measurements and culture extraction of T&O metabolites ...... 194
6.2.5 Geosmin and 2-MIB analysis ...... 195
6.2.6 Microscopic assessment of morphological differentiation ...... 195
6.2.7 Analysis and interpretation of the results ...... 197
6.3 Results ...... 197
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6.3.1 Experiment 1: The effects of carbon, nitrogen and phosphorous concentration on geosmin and 2-MIB production by Streptomyces spp...... 198
6.3.2 Experiment 2: The effects of temperature, pH and NaCl concentration on geosmin and 2-MIB production by Streptomyces spp...... 204
6.3.3 Experiment 3: The effects of calcium, potassium and magnesium concentration on geosmin and 2-MIB production by Streptomyces spp...... 210
6.3.4 Experiment 4: The effects of iron, zinc, copper and manganese concentration on geosmin and 2-MIB production by Streptomyces spp...... 216
6.3.5 The relationship between morphological differentiation and T&O metabolite production by Streptomyces spp...... 225
6.3.6 Summary of key findings ...... 232
6.4 Discussion ...... 234
6.4.1 The relationship between geosmin and 2-MIB and morphological differentiation in Streptomyces spp...... 235
6.4.2 Carbon, phosphorous and nitrogen in relation to geosmin and 2-MIB production by Streptomyces spp...... 237
6.4.3 Other physico-chemical parameters in relation to geosmin and 2-MIB production by Streptomyces spp...... 243
6.4.3.1 Physico-chemical factors inhibiting T&O metabolite production by Streptomyces spp...... 244
6.4.3.2 Physico-chemical factors stimulating T&O metabolite production by Streptomyces spp...... 249
6.5 Conclusions ...... 251
CHAPTER 7 - BIOSYNTHESIS OF GEOSMIN AND 2-METHYLISOBORNEOL BY STREPTOMYCES: WHY? ...... 254 7.1 Introduction ...... 254
7.2 Materials and methods ...... 260 IX
7.2.1 Microorganisms ...... 260
7.2.2 Co-culture experiments: the effect of interaction with other microorganisms on the production of geosmin and 2-MIB by Streptomyces spp...... 261
7.2.3 Antimicrobial assays of geosmin and 2-MIB ...... 263
7.2.4 Statistical analysis ...... 265
7.3 Results ...... 265
7.3.1 The effect of interaction with other microorganisms on the production of geosmin and 2-MIB by Streptomyces spp...... 265
7.3.2 Antimicrobial activity of geosmin and 2-MIB ...... 274
7.4 Discussion ...... 275
7.5 Conclusions ...... 284
CHAPTER 8: CONCLUSIONS AND RECOMMENDATIONS FOR FUTURE RESEARCH ...... 286
REFERENCES ...... 293 APPENDICES ...... 311
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LIST OF FIGURES
Figure Brief description Page 2.1 Sporulating colony and a photomicrograph of a Streptomyces sp. 9
2.2 Development of mycelium in Streptomyces 10
2.3 Simplified scheme of geosmin and 2-MIB biosynthesis in Streptomyces 19
3.1 Geosmin and 2-MIB concentrations in Grahamstown Reservoir from 1998 36 to 2012 3.2 Geosmin and 2-MIB concentrations in Chichester Reservoir from 2002 to 37 2012 3.3 Map showing the location of Grahamstown and Chichester Reservoirs 40
3.4 Loading column plot for all geosmin data in Grahamstown Reservoir 48
3.5 Loading column plot for the first geosmin event in Grahamstown Reservoir 48
3.6 Loading column plot for the second geosmin event in Grahamstown 49 Reservoir 3.7 Loading column plot for the third geosmin event in Grahamstown 49 Reservoir 3.8 Loading column plot for the fourth geosmin event in Grahamstown 50 Reservoir 3.9 Loading column plot for the fifth geosmin event in Grahamstown Reservoir 50
3.10 Loading column plot for the sixth geosmin event in Grahamstown 51 Reservoir 3.11 Loading column plot for the seventh geosmin event in Grahamstown 51 Reservoir 3.12 Grahamstown Reservoir geosmin and 2-MIB concentration and Anabaena 52 abundance
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3.13 Grahamstown Reservoir geosmin, 2-MIB and DO concentration 52
3.14 Loading column plot for all 2-MIB data for Grahamstown Reservoir 54
3.15 Loading column plot for the first 2-MIB event in Grahamstown Reservoir 54
3.16 Loading column plot for the second 2-MIB event in Grahamstown 55 Reservoir 3.17 Loading column plot for the third 2-MIB event in Grahamstown Reservoir 55
3.18 Grahamstown Reservoir geosmin and 2-MIB concentration and water level 56
3.19 Grahamstown Reservoir geosmin, 2-MIB and abundance of three 57 cyanobacteria 3.20 Loading column plot for all geosmin data in Chichester Reservoir 58
3.21 Loading column plot for the first geosmin event in Chichester Reservoir 58
3.22 Loading column plot for the second geosmin event in Chichester Reservoir 59
3.23 Loading column plot for the third geosmin event in Chichester Reservoir 59
3.24 Loading column plot for the fourth geosmin event in Chichester Reservoir 60
3.25 Chichester Reservoir geosmin and 2-MIB concentration and Anabaena 60 abundance 3.26 Geosmin and 2-MIB concentrations with silica concentration, turbidity and 61 water level in Chichester Reservoir 3.27 Seasonal presence of geosmin and 2-MIB in Grahamstown Reservoir from 64 1998 till 2012 3.28 Seasonal presence of geosmin and 2-MIB in Chichester Reservoir from 64 2002 till 2012 4.1 Schematic of sampling protocol for the marginal and open water sampling 80 sites
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4.2 Grahamstown Reservoir sampling locations 81
4.3 Chichester Reservoir sampling locations 81
4.4 Comparison of marginal sampling site in Chichester Reservoir during wet 83 and dry conditions 4.5 Comparison of marginal sampling site in Grahamstown Reservoir during 83 wet and dry conditions 4.6 Colonies of Streptomyces spp. growing on starch-casein agar 95
4.7 Mass spectra of the two analytes geosmin and 2-MIB and internal standard 102
4.8 Gas chromatogram showing the retention time of 2-MIB, biphenyl-d10 and 103 geosmin 4.9 Calibration curves of geosmin and 2-MIB analytical standards 103
4.10 Solid media prepared using environmental substrates for examining 105 Streptomyces spp. growth and geosmin and 2-MIB production. 4.11 Comparison of DNA yield obtained between the two cell lysis methods for 109 DNA extraction of spores and vegetative cell suspensions 4.12 Spore and vegetative cell suspensions prepared from cultures of S. 109 antibioticus and S. coelicolor A3(2) 4.13 Gel electrophoresis of the PCR product obtained by amplification of 110 Streptomyces spp. DNA using the primer pair targeting 16S rRNA 4.14 Standard curves for the qPCR assay using Streptomyces 16S rRNA 111 targeting primers 4.15 Mean cell densities of Streptomyces in Chichester Reservoir water samples 113 based on qPCR 4.16 Mean cell densities of Streptomyces in Grahamstown Reservoir water 114 samples based on qPCR 4.17 Mean cell densities of Streptomyces in Chichester Reservoir solid substrate 117 samples based on qPCR 4.18 Mean cell densities of Streptomyces in Grahamstown Reservoir solid 118 substrate samples based on qPCR
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4.19 Comparison of Streptomyces-like colonies enumerated on SCA and AIA 123 from Chichester environmental samples to the qPCR data 4.20 Comparison of Streptomyces-like colonies enumerated on SCA and AIA 123 from Grahamstown environmental samples to the qPCR data 4.21 Phylogenetic relationship between isolates and related sequences based on 126 16S rRNA gene sequences 4.22 Gel electrophoresis of the PCR product amplified with the Streptomyces- 127 specific 16S rRNA primers using DNA extracted from the 23 isolates 4.23 Production yields of geosmin and 2-MIB by 23 environmental isolates 129
4.24 Streptomyces and Nocardia isolates which demonstrated high geosmin 131 and/or 2-MIB producing potential. 4.25 Yields of geosmin and 2-MIB recovered from S. coelicolor A3(2) and S. 133 antibioticus cultures on growth media containing environmental samples 5.1 Experimental design for testing the production of geosmin and 2-MIB in 162 sediment under submerged and exposed conditions. 5.2 Anaerobic jar used to examine geosmin and 2-MIB production in sterilised 163 sediment inoculated with S. coelicolor A3(2) under anaerobic conditions. 5.3 Levels of geosmin and 2-MIB in sediment and overlying water 167
5.4 Relationship between geosmin and 2-MIB levels in the submerged 168 sediment and overlying water 5.5 Mean Streptomyces cell densities measured in each sediment treatment 170 under each experimental condition 6.1 Experimental design matrix to investigate the effects of changes in carbon, 189 nitrogen and phosphorous concentration on geosmin, 2-MIB and biomass production by Streptomyces spp. 6.2 Experimental design matrix to investigate the effects of changes in 190 temperature, pH and NaCl concentration on geosmin, 2-MIB and biomass production by Streptomyces spp. 6.3 Experimental design matrix to investigate the effects of changes in calcium, 190 magnesium and potassium concentration on geosmin, 2-MIB and biomass production by Streptomyces spp. 6.4 Experimental design matrix to investigate the effects of changes in iron, 191 copper, zinc and manganese concentration on geosmin, 2-MIB and biomass production by Streptomyces spp.
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6.5 Streptomyces different degrees of morphological differentiation 196
6.6 Contour plots showing the influence of nitrogen, phosphorous and carbon 199 concentration on the production of geosmin, 2-MIB and biomass by S. coelicolor A3(2) 6.7 Contour plots showing the influence of nitrogen, phosphorous and carbon 200 concentration on the production of geosmin, 2-MIB and biomass by S. antibioticus 6.8 MLR coefficients for geosmin, 2-MIB and biomass production by S. 201 coelicolor A3(2) (Experiment 1) 6.9 MLR coefficients for geosmin, 2-MIB and biomass production by S. 202 antibioticus (Experiment 1) 6.10 Contour plots showing the influence of NaCl concentration, pH and 205 temperature on the production of geosmin, 2-MIB and biomass by S. coelicolor A3(2) 6.11 Contour plots showing the influence of NaCl concentration, pH and 206 temperature on the production of geosmin, 2-MIB and biomass by S. antibioticus 6.12 MLR coefficients for geosmin, 2-MIB and biomass production by S. 207 coelicolor A3(2) (Experiment 2) 6.13 MLR coefficients for geosmin, 2-MIB and biomass production by S. 208 antibioticus (Experiment 2) 6.14 Contour plots showing the influence of potassium, calcium and magnesium 211 concentration on the production of geosmin, 2-MIB and biomass by S. coelicolor A3(2) 6.15 Contour plots showing the influence of potassium, calcium and magnesium 212 concentration on the production of geosmin, 2-MIB and biomass by S. antibioticus 6.16 MLR coefficients for geosmin, 2-MIB and biomass production by S. 213 coelicolor A3(2) (Experiment 3 ) 6.17 MLR coefficients for geosmin, 2-MIB and biomass production by S. 214 antibioticus (Experiment 3 ) 6.18 Contour plots showing the influence of micronutrient concentration on the 217 production of geosmin by S. coelicolor A3(2) 6.19 Contour plots showing the influence of micronutrient concentration on the 218 production of 2-MIB by S. coelicolor A3(2) 6.20 Contour plots showing the influence of micronutrient concentration on the 219 production of biomass by S. coelicolor A3(2)
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6.21 Contour plots showing the influence of micronutrient concentration on the 220 production of geosmin by S. antibioticus 6.22 Contour plots showing the influence of micronutrient concentration on the 221 production of 2-MIB by S. antibioticus 6.23 Contour plots showing the influence of micronutrient concentration on the 222 production of biomass by S. antibioticus 6.24 MLR coefficients for geosmin, 2-MIB and biomass production by S. 223 coelicolor A3(2) (Experiment 4) 6.25 MLR coefficients for geosmin, 2-MIB and biomass production by S. 224 antibioticus (Experiment 4) 6.26 Relationship between geosmin and 2-MIB production by S. coelicolor 226 A3(2) and differentiation value based on all experimental data 6.27 Relationship between geosmin and 2-MIB production by S. antibioticus 227 and differentiation value based on all experimental data 6.28 S. coelicolor A3(2) biomass under the different combinations of carbon, 229 nitrogen and phosphorous concentrations 6.29 S. antibioticus biomass under the different combinations of carbon, 229 nitrogen and phosphorous concentrations 6.30 S. coelicolor A3(2) biomass under different conditions of temperature, pH 230 and NaCl concentration 6.31 S. antibioticus biomass under different conditions of temperature, pH and 230 NaCl concentration 6.32 S. coelicolor A3(2) under different conditions of micronutrient 231 concentration 6.33 S. antibioticus under different conditions of micronutrient concentration 232
7.1 Co-culturing scheme to test the effect of Streptomyces spp. interaction with 262 other microorganisms on the production of geosmin and 2-MIB. 7.2 Protocol for testing the antimicrobial effects of geosmin and 2-MIB 264
7.3 Geosmin and 2-MIB production by S. coelicolor A3(2) when grown as a 266 pure culture and when co-cultured 7.4 Geosmin and 2-MIB production by S. antibioticus when grown as a pure 267 culture and when co-cultured
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7.5 Relationship between geosmin and 2-MIB production by Streptomyces spp. 270 and differentiation value 7.6 Microscopic photos of co-cultures biomass for S. coelicolor A3(2) 271
7.7 Co-cultures for studying the effect of interaction with other microbes on 272 the production of geosmin and 2-MIB by S. coelicolor A3(2) 7.8 Microscopic photos of co-culture biomass for S. antibioticus 273
7.9 Co-cultures for studying the effect of interaction with other 274 microorganisms on the production of geosmin and 2-MIB by S. antibioticus
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LIST OF TABLES
Table Brief description Page 2.1 Molecular structure, weight and formula of geosmin and 2-MIB 13 3.1 Chemical, biological and physical water quality parameters in the PLS 42 models 3.2 Overview of each PLS model for geosmin and 2-MIB in Grahamstown 45 Reservoir 3.3 Overview of each PLS model for geosmin in Chichester Reservoir 46
4.1 Sampling locations and samples collected from Grahamstown and 82 Chichester Reservoirs 4.2 Sequences of the primer pair for qPCR of Streptomyces-specific 16S rRNA 88
4.3 Chemical composition of two selective growth media used for the isolation 92 of Streptomyces-like colonies. 4.4 Inhibitors added to both selective growth media 93
4.5 Collection details of the 23 Streptomyces-like isolates selected for 96 identification and examination of geosmin and 2-MIB production ability 4.6 Samples collected from Grahamstown and Chichester Reservoirs for testing 105 Streptomyces growth and geosmin and 2-MIB production 4.7 Results from a BLASTn search of the 16S rRNA sequences of isolates 125
4.8 Recovery efficiency of geosmin and 2-MIB using the extraction procedures 128
5.1 DO levels measured above the sediment surface for sediment treatments 171 subjected to submerged conditions. 6.1 Stages of morphological differentiation and corresponding numerical values 196
6.2 Summary of the association of physico-chemical parameters and life cycle 234 stage of Streptomyces with geosmin and 2-MIB production
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ABBREVIATIONS 2-MIB 2-methylisoborneol AIA Actinomycete isolation agar ANOVA Analysis of variance ANZECC Australian and New Zealand Environment and Conservation Council ATCC American Type Culture Collection ATP Adenosine triphosphate BLAST Basic local alignment search tool bp Base pairs BSTFA N,O-Bis(trimethylsilyl)trifluoroacetamide CARD Catalysed reporter deposition CCR Carbon catabolite repression CFU Colony forming unit
CT Cycle threshold number DF Degrees of freedom DNA Deoxyribonucleic acid dNTP Deoxyribonucleotide triphosphate DO Dissolved oxygen EC Electrical conductivity FISH Fluorescent in situ hybridisation FPP Farnesyl diphosphate GC-MS Gas chromatograph-mass spectrometry GPP Geranyl diphosphate HWC Hunter Water Corporation IPP Isopentenly diphosphate LC 96 Light Cycler 96 LLE Liquid-liquid extraction LOO-CV Leave one out cross validation
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MEP/ DOXP 2-C-methyl-D-erythritol 4-phosphate/1-deoxy-D-xylulose 5-phosphate MLR Multiple linear regression MVA Mevalonic acid NA Nutrient agar NBRC National Biological Resource Center NCBI National Center for Biotechnology Information OTC Odour threshold concentration PAC Powdered activated carbon PCA Principal components analysis PLS Partial least squares PRESS Predicted residual sum of square qPCR Quantitative polymerase chain reaction RAS Recirculating aquaculture system RNA Ribonucleic acid rRNA Ribosomal ribonucleic acid SAM S-adenosylmethionine SCA Starch-casein agar SD Standard deviation SDS Sodium dodecyl phosphate SE Standard error SIM Selective ion monitoring SS Sum of squares T&O Taste and odour TE Tris Ethylenediaminetetraacetic acid
TMS Trimethylsilyl VOC Volatile organic compound
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ABSTRACT Providing adequate volumes of safe, clean drinking water to the world’s growing population is a continuous and increasing challenge for water utilities. While prime attention is placed on health aspects, consumers generally judge water quality by its aesthetic value. The presence of compounds which impart taste and odour (T&O) in drinking water supplies often leads to the misconception that the water is unsafe for consumption, triggering consumer complaints and high treatment costs for water utilities. Two biologically sourced compounds which respectively cause ‘earthy’ and ‘musty’ T&O in drinking water supplies worldwide are the secondary metabolites geosmin and 2-methylisoborneol (2-MIB).
The research presented in this thesis was initiated and supported by Hunter Water Corporation (NSW, Australia) in response to earthy-musty T&O problems periodically experienced in drinking water supplies (Grahamstown and Chichester Reservoirs). A preliminary analysis of historical water quality data in these reservoirs revealed a close association between abundance of the cyanobacterial genus Anabaena and elevated concentrations of geosmin, while 2-MIB could not be reliably linked to any routinely measured parameter. Although T&O events are notoriously ascribed to cyanobacteria, the filamentous bacterial genus Streptomyces, major producers of geosmin and 2-MIB in soil, have long been suspected to also play a potential role in imparting these metabolites into drinking water supplies. There has been a distinct paucity of knowledge regarding their ecology in freshwater environments and consequently, their significance as contributors to T&O events is not well-established. This thesis presents both field- and laboratory-based studies which were conducted in order to examine the potentiality of Streptomyces to contribute to earthy-musty T&O problems in drinking water reservoirs.
A temporal and spatial sampling program combined with the application of a molecular technique for Streptomyces quantification (qPCR targeting the 16S rRNA gene) established the widespread distribution and abundance of these bacteria within the water mass, bottom
XXI sediments and in marginal substrates of the reservoirs. The detection of significantly greater Streptomyces population densities in the water mass during prolonged wet conditions compared to extended dry conditions provided evidence to support the hypothesis that they are washed from surrounding marginal habitats into the reservoirs, consistent with them being ‘terrestrial’ bacteria. Contradicting widespread opinion in the literature that they are not ‘aquatic’ bacteria and survive only as dormant spores in water, vegetative cells (detected through a differential cell lysis protocol for DNA extraction) were found to comprise a considerable proportion of Streptomyces populations in the water mass. Together with the finding that sterilised reservoir water supported the growth and geosmin and 2-MIB production by Streptomyces spp. in the laboratory indicates the potentiality of these bacteria to be metabolically active in water and contribute to in situ production of T&O metabolites.
Substrates at the margins of the reservoirs including soil, sediment and plant debris represented the major habitat of Streptomyces and the hypothesis that exposure of such substrates following water level recession during dry conditions stimulates the growth and activity of these aerobic bacteria, was largely supported by both field data and a laboratory simulation of these conditions. Together these studies indicated the potential significance of marginal substrates as a major source of Streptomyces and their T&O metabolites, which can enter the adjacent water mass following rain events. Confirmation that all Streptomyces reservoir cultivars could produce geosmin and 2-MIB provided additional evidence to support their role as potentially significant contributors to T&O metabolites in drinking water supplies.
Multivariate laboratory studies examining the influence of physico-chemical factors on the production of geosmin and 2-MIB by Streptomyces spp. established that the production of these T&O metabolites was highly coordinated with the reproductive (sporulation) stage of the Streptomyces life cycle. Thus physico-chemical factors that trigger Streptomyces to cease vegetative growth and enter the reproductive developmental stage would conceivably allow
XXII elucidation of the conditions which also trigger significant production of their T&O metabolites. Such conditions included lowest concentrations of macronutrients (C, N and P) while higher concentrations of NaCl and copper were found to favour vegetative growth and thus inhibit T&O metabolite production. In consideration of these results, the levels of physico-chemical characteristics in the surface waters of Grahamstown and Chichester Reservoirs appear to be suitable for Streptomyces growth, differentiation and production of geosmin and 2-MIB.
While the biological function of geosmin and 2-MIB is not currently known, many other secondary metabolites produced by Streptomyces function as antimicrobial compounds, produced during times of adversity to antagonise competing microorganisms and coincide with their initiation of reproductive growth. Having established that geosmin and 2-MIB production was stimulated under conditions which also trigger reproductive growth (e.g. nutrient limitation), it was hypothesised that they too may function as antimicrobial compounds, however subsequent co-culturing and antimicrobial assays led to the rejection of this hypothesis. Based on literature evidence, several alternative propositions are outlined regarding the possible biological function of these compounds related to regulation of the formation, germination and dispersal of Streptomyces spores.
The findings of the studies presented in this thesis indicate the potential significance of Streptomyces as major contributors to the occurrence of geosmin and 2-MIB in drinking water supplies. Furthermore, understanding of the influence of environmental factors and Streptomyces life cycle stage on the biosynthesis of these compounds and their possible biological function has been extended.
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PUBLICATIONS Asquith, E.A., Evans, C.A., Geary, P.M., Dunstan, R.H. & Cole, B. (2013). The role of actinobacteria in taste and odour episodes involving geosmin and 2-methylisoborneol in aquatic environments. Journal of Water Supply: Research and Technology—AQUA, 62(7), 452-467.
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CHAPTER 1 - INTRODUCTION
The microbial world represents the largest unexplored reservoir of biodiversity on Earth. The vast array of microbial activities and their importance to the biosphere and to human economies, provide strong bases for understanding their diversity and in exploiting them for the benefit of human society (Ahmad et al., 2011). One ubiquitous group of microbes are
Streptomyces, a genus of soil inhabiting, filamentous bacteria which function as saprophytes
(Goodfellow & Williams, 1983; Williams et al., 1984). The diverse secondary metabolism of these microorganisms also renders them highly useful as a source of pharmaceutical products (Hopwood, 2007). Among the secondary metabolites they produce are the terpenoids geosmin and 2-methylisoborneol (2-MIB), responsible for the ‘earthy’ and
‘musty’ odour of soil respectively. These compounds present a challenge to the drinking water industry, as they are responsible for the majority of biologically induced taste and odour (T&O) problems in drinking water supplies worldwide (Juttner & Watson, 2007;
Srinivasan & Sorial, 2011).
Earthy-musty T&O episodes involving geosmin and 2-MIB are frequent and often unpredictable and although they do not pose a public health risk, can cause serious concerns regarding public perception of the aesthetic quality of the water supply. T&O problems therefore contribute to a palatability issue in drinking water systems rather than being a health concern for consumers (Juttner & Watson, 2007; Journey et al., 2013). As a consequence, water utilities must bear the costs of advanced treatment processes to remove these chemically stable compounds (Srinivasan & Sorial, 2011).
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The research presented in this thesis was initiated and supported by Hunter Water
Corporation (HWC) in response to a long history of sporadic earthy-musty T&O episodes experienced in two of their primary drinking water storages, Grahamstown and Chichester
Reservoirs (NSW, Australia). Prediction, control and management of severe T&O problems necessitates detection of their biological origins. The literature review presented in Chapter
2 emphasises that numerous organisms have been implicated in the production of T&O episodes involving geosmin and 2-MIB, particularly the photosynthetic cyanobacteria. Other microorganisms, notably members of the genus Streptomyces, have also been suggested as possible contributors. An analysis of historical water quality data in Chapter 3 reveals that the majority of (but not all) episodes of elevated geosmin concentration in Grahamstown and
Chichester Reservoirs have coincided with high abundance of the cyanobacterial genus
Anabaena, while 2-MIB concentrations have not been linked to cyanobacterial origin or any other biological source monitored in the reservoirs. HWC has considered that non- cyanobacterial related occurrences of geosmin and 2-MIB in these reservoirs may have originated from Streptomyces, which has prompted research into their potential involvement in T&O problems. Surveys of existing literature reveal a distinct lack of knowledge of the relative importance of Streptomyces in relation to T&O (Zaitlin & Watson, 2006; Juttner &
Watson, 2007) and the work presented in this thesis focuses on elucidating their potential contribution to this widespread aesthetic water quality issue. Both field-based and laboratory- based research were undertaken to achieve this goal. At the heart of the study are four basic research questions:
Are Streptomyces likely to be responsible for non-cyanobacterial related T&O
episodes involving geosmin and 2-MIB in drinking water reservoirs?
Are Streptomyces likely to actively produce geosmin and 2-MIB in aquatic habitats,
and if not, what is the most likely origin of Streptomyces-derived T&O metabolites
in reservoirs? 2
Which environmental factors and life cycle stage of Streptomyces are most relevant
to elevated geosmin and 2-MIB production by these bacteria?
Why do Streptomyces produce geosmin and 2-MIB?
Streptomyces, being the microorganisms from which geosmin and 2-MIB were first chemically characterised in the 1960s (Gerber & Lecheval, 1965; Gerber, 1969), have an ambiguous role as contributors to the occurrence of these compounds in freshwater systems.
As their aquatic ecology has largely been studied by cultivation on selective isolation media, there is a lack of reliable information on their abundance and activity in freshwater systems
(Cross, 1981; Zaitlin & Watson, 2006; Juttner & Watson, 2007). An objective of this research was to assess the occurrence (abundance and activity) of Streptomyces in a diverse range of freshwater habitats and to better understand their relative potential to contribute to T&O episodes in Grahamstown and Chichester Reservoirs. The results of a detailed spatial and temporal sampling program undertaken at both reservoirs to achieve this objective are presented in Chapter 4. A molecular technique, quantitative polymerase chain reaction
(qPCR) using 16S rRNA genus-specific primers, was developed to estimate the abundance of Streptomyces in samples collected. This study reports on the use of two cell lysis procedures for the extraction of DNA from environmental samples, which exploits the differential resistance of vegetative cells and spores of Streptomyces to cellular disruption, enabling determination of the relative abundance of vegetative cells (potentially active phenotype) as opposed to spores (dormant phenotype) in these samples.
In order to focus the investigation, several hypotheses concerning the presence and activity of Streptomyces in freshwater environments were developed. The growth and odour production by Streptomyces in the water mass of reservoirs has been largely disputed and
3 rather, the terrestrial wash-in of these bacteria in substrates at the margins of reservoirs entrained with their odorous secondary metabolites has been suggested as a significant means by which they may contribute to T&O (Johnston & Cross, 1976a; Cross, 1981; Wood et al.,
1983a; Jensen et al., 1994; Zaitlin et al., 2003a; Zaitlin et al., 2003b; Zaitlin & Watson, 2006;
Tung et al., 2006; Zuo et al., 2009a; Zuo et al., 2010; Lee et al., 2011). To support this proposition, it was hypothesised that their abundance in the water mass during periods of runoff (prolonged wet conditions) would greatly exceed that during extended dry conditions.
It was hypothesised that as Streptomyces are primarily terrestrial bacteria, their abundance and activity would be greater in substrates including sediment and soil at the margins of reservoirs compared to that in the deep bottom sediments. Being aerobic bacteria, it has been suggested that water level recession during dry conditions which exposes substrates such as sediment, soil and plant debris at the margins of reservoirs, stimulates growth and odour production by Streptomyces. Thus, dry conditions could represent a period of significant geosmin and 2-MIB production which can subsequently enter the adjacent water mass via run-off and rising water levels following rainfall (Wood et al., 1983a). This anecdotally occurred in Grahamstown Reservoir in the late 1980s, when a T&O event followed an extensive drought period. Thus it was hypothesised that marginal substrates would harbor larger and more active Streptomyces populations when exposed during dry conditions compared to submerged wet conditions. The objective of the study presented in Chapter 5 was to provide experimentally acquired evidence using a laboratory simulation of these conditions against which to evaluate this proposed mechanism of Streptomyces contribution to geosmin and 2-MIB occurrence in drinking water supplies.
Additional objectives of the field and laboratory work presented in Chapter 4 were to obtain, identify (by 16S rRNA sequence analysis) and examine the geosmin and 2-MIB production ability of Streptomyces isolates and to investigate the capacity of reservoir substrates (soil, 4 water, sediment and plant material) as the sole source of nutrition, to support the growth and
T&O metabolite production of Streptomyces spp. under laboratory conditions. Both of these studies were conducted to gather further evidence to assess the potential of Streptomyces to contribute to T&O episodes in the reservoirs.
Chapter 6 describes the use of a multivariate experimental design to examine the effects of changes in a number of physico-chemical factors representative of levels encountered in the environment, on geosmin and 2-MIB production by two Streptomyces spp. These included macronutrients (carbon, nitrogen, phosphorous, potassium, calcium and magnesium), temperature, pH, salinity and micronutrients (iron, copper, manganese and zinc). This laboratory-based study was undertaken with the objective to better understand and characterise important factors and interactions between factors which may either stimulate or inhibit geosmin and 2-MIB production by Streptomyces. Such understandings may have implications for their potential production of these compounds in the environment. Based on observations of enhanced geosmin and 2-MIB production in Streptomyces cultures exhibiting high degrees of morphological differentiation (aerial mycelial growth and sporulation) in several literature reports (Bentley & Meganathan, 1981; Dionigi et al., 1992; Scholler et al.,
2002; Tung et al., 2006), a key objective of this study was to gather quantitative evidence to assess the apparent association between T&O metabolite production and life cycle stage of the organisms.
Chapter 7 represents the final component of the experimental work in which the objective was to better understand the functional role of these secondary metabolites in the life of
Streptomyces and potentially challenge the notion that geosmin and 2-MIB are simply metabolic waste products, which is the widespread opinion held in the water industry
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(Watson, 2003). Here, the investigation was founded on the hypothesis that these secondary metabolites, like many others produced by Streptomyces, function as antimicrobial compounds (Hopwood, 2007). Other possible biological roles of these terpenoids are also discussed in this chapter.
Finally, Chapter 8 summarises the main findings and conclusions to be drawn from the research in this thesis. It includes recommendations for future work to further elucidate the contribution of Streptomyces and indeed, other organisms, to T&O issues involving geosmin and 2-MIB in drinking water supplies, and better understand the triggers for their production and possible biological functions.
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CHAPTER 2 - STREPTOMYCES AND THE OCCURRENCE OF GEOSMIN AND 2-METHYLISOBORNEOL IN AQUATIC ENVIRONMENTS
2.1 Introduction
Streptomyces, a prominent group of actinobacteria, have received significant research attention since the discovery of their rich and diverse secondary metabolism in the 1940s, making them valuable providers of clinically important biomolecules (Hopwood, 2007).
However, two secondary metabolites produced by many Streptomyces spp. and certain other actinobacteria – the volatile tertiary alcohols geosmin (derived from the Greek words ‘ge’ and ‘osme’, meaning earth and odour respectively) and 2-methylisoborneol (2-MIB) (Gerber,
1979) – have proven problematic to the drinking water industry. Better recognised for providing soil with its pleasant earthy-musty odour, these compounds are also responsible for biologically induced T&O problems in drinking water supplies worldwide which occur sporadically, causing great consumer dissatisfaction and high treatment costs for water utilities. Although often untraced to their biological origins, numerous organisms from prokaryotes to eukaryotes, have been implicated as the cause of this aesthetic water quality issue (Juttner & Watson, 2007). Brief consideration will be paid to these, but it is the potential role that Streptomyces and other actinobacteria play in generating unpleasant episodes of earthy-musty T&O that is the focus of this review. An overview of the taxonomy, physiology and ecology of these bacteria is presented followed by a detailed review of the current evidence documented in the literature concerning geosmin and 2-MIB production by
Streptomyces. A critical assessment of habitats within freshwater systems in which they may be metabolically active residents and thus potential sources of T&O is provided. To date, the aquatic ecology of these bacteria remains poorly understood and debate surrounds whether they exist solely as dormant spores of terrestrial origin or are capable of growing and biosynthesising these odorous compounds in aquatic environments. Current understandings 7 of the chemical ecology and biosynthetic pathways of geosmin and 2-MIB in addition to the conditions under which these secondary metabolites are produced by Streptomyces are reviewed.
2.2 Actinobacteria and the genus Streptomyces: What are they?
Actinobacteria comprise a ubiquitous class of Gram-positive heterotrophic bacteria, characterised by DNA rich in guanine and cytosine and display diverse morphologies from simple bacterial shapes of cocci through to forms with differentiated branched mycelium
(Vobis, 1997; Ventura et al., 2007). In the literature relating to T&O in drinking water, the term ‘actinomycetes’ has been widely used to refer to taxa of this class, particularly the large filamentous genus Streptomyces (often used as a pseudonym for this genus and other filamentous actinobacteria). This was somewhat appropriate since the order Actinomycetales, from which this term is derived, previously contained T&O producing, filamentous genera.
However, with the availability of new 16S rRNA gene sequence information, recent modifications in the phylogenetic relationships of the class actinobacteria have been made, with the reconstruction of higher (taxonomic) ranks (Zhi et al., 2009; Ludwig et al., 2012).
The order Actinomycetales now contains only one family (Actinomycetaceae) whereas the remaining families have been reclassified within new orders created by elevation of existing suborders to order status (e.g. Streptomyces now belongs to the new order Streptomycetales)
(Ludwig et al., 2012). Hence the common term ‘actinomycetes’ will be avoided in this review and throughout this thesis. Rather these bacteria will be referred to as actinobacteria where appropriate but predominately as Streptomyces, the genus of greatest relevance to T&O.
Streptomyces, the most representative genus, among other actinobacteria, exhibit a filamentous growth habit and produce spores, morphological traits which confused their 8 discoverers in the 1870s, growing like fungi but with tiny dimensions like bacteria. They are recognised by their tough, chalky or leathery and often pigmented colonies they develop on solid media (Figure 2.1). Their resilient spores disperse in the environment and germinate under favourable conditions to form a dense branching hydrophilic vegetative mycelium on solid substrates, which forages for nutrients and assembles them into cellular structures.
Upon depletion of nutrients, this biomass is utilised to build a vertically growing hydrophobic aerial (secondary) mycelium. As these specialised hyphae grow and mature, synchronised cell septations develop between nuclear areas, resulting in the formation spores (Figure 2.2)
(Goodfellow & Williams, 1983; Williams et al., 1984).
Figure 2.1 Sporulating colony growing on starch-casein agar (left) and a photomicrograph of a Streptomyces sp. detailing the presence of hyphae and spores (right).
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Figure 2.2 Development of mycelium in Streptomyces. Modified from Vobis (1997).
2.3 The ecological significance of actinobacteria
Actinobacteria, particularly the genus Streptomyces, are a successful group of bacteria occurring in a multiplicity of terrestrial, aquatic and man-made environments. Soil is their greatest natural reservoir where they constitute a significant proportion of the bacterial community (Goodfellow & Williams, 1983; Williams et al., 1984). Their impressive saprophytic abilities are perhaps of greatest ecological significance in soil, delivering an immense contribution to organic matter turnover. Hydrolytic and oxidative extracellular enzyme secretions combined with the penetrating abilities of their hyphae, make these filamentous bacteria highly adapted to extensively colonising complex organic substrates
(e.g. lignocellulose) which other microbes cannot exploit (McCarthy & Williams, 1992). In the rhizosphere they protect plant roots by antagonising phytopathogens and exhibit traits of plant growth promoting rhizobacteria (Franco-Correa et al., 2010). Streptomyces are of particular medical significance, being a major source of enzymes, vitamins and secondary
10 metabolites which have a broad spectrum of biological activities (e.g. antibacterial, antifungal and enzyme inhibitory) (Ventura et al., 2007).
As a collective group, actinobacteria have provided mankind and the environment with many beneficial microorganisms but also some of its greatest foes, namely pathogenic forms which cause some widespread and intensely studied animal and plant diseases (e.g. the genus
Corynebacterium and Mycobacterium) (Ventura et al., 2007). Actinobacteria are also problematic in causing immensely thick scumming on the surface of secondary sewage treatment aeration tanks and their overgrowth can create problems with sludge bulking, impairing sludge settling and reducing effluent quality (Kämpfer & Wagner, 2002). The production of secondary metabolites causing T&O problems in drinking water supplies is another significant challenge these bacteria present to the water industry. This chapter presents a review of the literature concerning the problem of biologically produced T&O involving geosmin and 2-methylisobornoel (2-MIB) in aquatic environments, with an emphasis on the role of Streptomyces and other taxa belonging to actinobacteria.
2.4 Geosmin and 2-MIB in drinking water supplies
Surface waters including reservoirs, natural lakes and rivers, are important sources of potable water throughout the world, in which water utilities often encounter sporadic episodes of earthy-musty T&O, rendering it unpalatable for human consumption. Despite numerous microorganisms being implicated as sources (e.g. cyanobacteria, actinobacteria and fungi), the majority of T&O outbreaks are neither anticipated nor traced to their biological origins
(Juttner & Watson, 2007). The status of the problem in Australia was captured in a 2005 survey involving 37 drinking water providers, representing five million consumers.
Approximately 78% of the suppliers reported problems regarding earthy-musty T&O and 11 only five providers had identified a definite link between potential causative microorganisms
(namely cyanobacteria) and T&O causing compounds (Hobson et al., 2010). The ambiguity of the cause of this serious aesthetic water quality problem is certainly not restricted to
Australia; it affects drinking water utilities worldwide (Juttner & Watson, 2007).
Although a number of microbial secondary metabolites give rise to unpleasant T&O in water, attention is mainly focused on the ‘earthy’ compound geosmin (trans-1, 10-dimethyl-trans-
9-decalol) and the ‘musty’ compound 2-MIB (2-methylisoborneol or 1,2,7,7-tetramethyl- exo-bicyclo-[2,2,1]-heptan-2-ol) (Table 2.1). While these semi-volatile terpenoids pose no risk to human health, T&O is perceived by the general public as a primary indicator of the safety and acceptability of drinking water and encourages them to switch to alternative supplies such as bottled water (Juttner & Watson, 2007; Srinivasan & Sorial, 2011). A major problem concerning these compounds in drinking water is that they can be detected by human olfactory sense at exceptionally low concentrations, the odour threshold concentrations
(OTCs) being as low as 4 ng L-1 and 9 ng L-1 for geosmin and 2-MIB respectively. Owing to their low concentration and chemical stability, their removal from drinking water has proven to be a difficult task and cannot be achieved using conventional water treatment such as flocculation, filtration and chlorination. Some alternative treatment technologies have been employed with variable success, including adsorption by powdered or granular activated carbon, advanced oxidation processes and biological/physical filtration technologies
(Srinivasan & Sorial, 2011; Ho et al., 2012). Despite it being well known that T&O events involving geosmin and 2-MIB are problematic in Australia, few documented reports have been made relative to other countries (Uwins, 2011). Considering the costs that these nuisance compounds present to the water industry, identifying their origins is critical.
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Table 2.1 Molecular structure, weight and formula of geosmin and 2-MIB. Modified from Juttner and Watson (2007). Compound name Chemical structure Molecular Molecular weight formula
Geosmin 182.3 C12H22O (trans-1, 10-dimethyl-trans-9- decalol)
2-methylisoborneol, 2-MIB 168.3 C11H20O (1,2,7,7-tetramethyl-exo-bicyclo- [2,2,1]-heptan-2-ol)
2.4.1 Non-actinobacterial producers of geosmin and 2-MIB
Predicting and preventing geosmin and 2-MIB accumulation in drinking water supplies is vital for long term delivery of high quality water to consumers. The literature demonstrates the multiplicity of microorganisms that can produce these compounds and the range of habitats that could foster their production. The phenomenon is highly complex and as such, a large number of T&O cases remain unsolved. Photoautotrophic cyanobacteria, being widely distributed in freshwater systems are thought to be the main producers of T&O. The regular monitoring of cyanobacterial seasonal dynamics in drinking water storages has provided ample opportunity to analyse relationships between these bacteria and T&O (Juttner
& Watson, 2007). Often, strong correlations between planktonic cyanobacteria and T&O have been documented (Izaguirre et al., 1999; Park et al., 2001; Zhang et al., 2010). Geosmin and 2-MIB occurrences have been well correlated with the abundance of planktonic
Anabaena (Bowmen et al., 1992; Jones & Korth, 1995; Izaguirre et al., 1999; Uwins et al.,
2007; Hobson et al., 2010). Some reports indicate that benthic cyanobacteria such as
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Aphanizomenon, Oscillatoria, Phormidium and Pseudanabaena are also associated with earthy-musty T&O (Vilalta et al., 2004; Izaguirre & Taylor, 2007).
The presence of geosmin and 2-MIB during periods of low cyanobacterial biomass indicates that these bacteria are not the only source of these compounds (Klausen et al., 2004;
Dzialowski et al., 2009). Indeed other microbes, notably actinobacteria, may also be responsible for their production. Myxobacteria, ameoba and liverworts have also been confirmed as geosmin producers (Hayes et al., 1991; Sporle et al., 1991; Yamamoto et al.,
1994; Schulz et al., 2004; Dickschat et al., 2005). In particular some genera of fungi (e.g.
Penicillium and Chaetomium) produce geosmin and as these eukaryotic organisms are readily detectable in aquatic environments, they may be an underestimated source of T&O which are yet to be systematically investigated (Kikuchi et al., 1981; Borjesson et al., 1993;
Larsen & Frisvad, 1995).
2.4.2 Geosmin and 2-MIB production by Streptomyces and other actinobacteria
Actinobacteria, particularly the genus Streptomyces, have gained attention throughout the drinking water industry as a major source of geosmin and 2-MIB since these secondary metabolites were first extracted and identified from Streptomyces cultures (Gerber &
Lecheval, 1965; Gerber, 1969). They were initially dealt with in the literature as a potential source of T&O at the end of the 1920s followed by a surge of studies from the 1950s ascribing almost all earthy odours in natural waters to filamentous actinobacteria (Silvey et al., 1950;
Silvey & Roach, 1953; Silvey, 1963; Higgins & Silvey, 1966; Silvey & Roach, 1975). Silvey and Roach (1975) stated that “there is a direct relationship between the actinomycete population that occurs in raw water and the T&O compounds that are produced by these organisms” (p. 270). These reports stimulated microbiologists to count ‘aquatic’
14 actinobacteria in water supplies and correlate them with the concentration of odours. Such monitoring efforts regularly failed to produce clear relationships. Cross (1981) provided a simple explanation for this: spores, being the principal propagules of Streptomyces and other filamentous actinobacteria found in water, are not metabolically active and therefore cannot produce T&O metabolites. Rather these compounds arise subsequent to secondary mycelial development. As spores and vegetative forms are indistinguishable using traditional culture techniques, it remains uncertain if Streptomyces or other actinobacteria can be metabolically active in freshwater environments and contribute to earthy-musty T&O events.
2.4.2.1 Conditions of production
Being secondary metabolites, geosmin and 2-MIB are produced by Streptomyces during secondary mycelial growth coinciding with sporulation. This has been demonstrated by the inhibition of geosmin production by Streptomyces mutants incapable of aerial mycelium development as well as by normal isolates growing on media not conductive to sporulation
(Bentley & Meganathan, 1981; Dionigi et al., 1992). Numerous studies have reported greater production of geosmin and 2-MIB by Streptomyces cultures exhibiting a higher degree of morphological differentiation (Scholler et al., 2002; Tung et al., 2006).
As the secondary mycelial stage of growth is obligately aerobic, Streptomyces require the presence of oxygen for geosmin and 2-MIB production. Dionigi and Ingram (1994) and
Schrader and Blevins (1999) reported that increased geosmin production occurred in
Streptomyces cultures exposed to higher concentrations of atmospheric oxygen, although contradictory observations were reported by Schrader and Blevins (1999). Carbon dioxide concentration also appears to have an effect, with an elevated concentration (10% CO2 atmosphere) observed to decrease geosmin production by S. albidoflavus (Sunesson et al.,
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1997), although Schrader and Blevins (1999) observed more geosmin production in cultures of S. halstedii containing 10% carbon dioxide atmospheres compared to 5% and ambient carbon dioxide concentration. Despite being neutrophiles, these bacteria have been detected in both moderately acidic (pH 5) and alkaline (pH 9) aquatic environments (Jiang & Xu,
1996). Blevins et al. (1995) showed that S. halsetdii grew optimally in a neutral pH range (6-
7) but interestingly, the highest geosmin production occurred at pH 9 and in the extensive range of 6 to 11. Similar observations were reported by Yagi et al. (1987) and Weete et al.
(1977).
The prevalence of earthy-musty T&O during warmer seasons is well documented (Tung et al., 2006; Zaitlin & Watson, 2006; Juttner & Watson, 2007). Undoubtedly, temperature is an important parameter affecting the metabolic activity of Streptomyces, which are predominately mesophilic, exhibiting optimum growth between 25°C to 30°C (Goodfellow
& Williams, 1983). Wood et al. (1985) determined that the minimum temperature for geosmin production by S. albidoflavus in nutrient amended reservoir water was 15°C and that all documented cases of earthy odour problems occurred when the water temperature exceeded this. However, more recently, Zuo et al. (2010) found that some sediment isolates of Streptomyces could grow slowly and produce relatively low concentrations of geosmin at
4°C and 10°C. Generally, Streptomyces cultures increase their production yields with increasing incubation temperature (≤40°C), with optimum production in the range of 25°C to
30°C (Weete et al., 1977; Yagi et al., 1987; Dionigi & Ingram, 1994; Blevins et al., 1995;
Tung et al., 2006).
Although low nutrient concentration was found by Wood et al. (1985) to be major constraint on the growth and geosmin production by a Streptomyces isolate in the water mass of 16 oligotrophic reservoirs, nutrient poor conditions have also supported the production of T&O compounds by these bacteria (Zaitlin et al., 2003a; Zaitlin et al., 2003b). Schrader and
Blevins (2001) found that easily assimilated carbon sources (e.g. glucose and maltose) promoted biomass production whilst providing carbon sources such as mannitol and glycerol, promoted maximal geosmin production by S. halstedii. Similarly, Saadoun (2005) found that glycerol promoted 2-MIB production compared to glucose which was more favourable for biomass development in cultures of S. violaceusniger. Carbon sources which readily serve as growth substrates often repress the need for secondary metabolism and therefore T&O production by Streptomyces, being normally triggered under nutrient limiting conditions.
Similarly, repression of cyanobacterial growth appears to promote geosmin biosynthesis due to the increased availability of precursor compounds (e.g. pyruvate and acetyl-CoA) (Blevins et al., 1995; Saadoun et al., 2001).
Blevins et al. (1995) observed that limiting concentrations of nitrate and ammonia (16.5 µg/L and 13.1 µg/L respectively) induced geosmin production and suppressed growth by S. halstedii. Uwins (2011) and Weete et al. (1977) also observed more geosmin production in nitrogen reduced liquid cultures of Streptomyces spp. Increasing phosphorus concentration
(up to 36.2 µM) was found to increase geosmin and biomass production by S. halstedii, with no geosmin detected at the lowest concentrations tested (0.7-7.3 µM) (Schrader & Blevins,
2001). Uwins (2011) similarly reported increasing geosmin production by S. coelicolor
A3(2) with increasing phosphorous concentration. The effect of other macronutrients and micronutrients on geosmin production by S. halstedii has also been investigated, with low concentrations of K, Fe, Co, Ca, Cu, Mn and Zn stimulating geosmin synthesis while higher concentrations inhibited or repressed production. Significant geosmin production was still observed in the absence of some of these trace elements (Fe, Co, Ca and Mn) (Schrader &
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Blevins, 2001). Salinity has also been observed to affect geosmin production, with Rezanka and Votrubam (1998) reporting that an increase in sodium chloride concentration (0% to
12%) in S. avermitilis cultures decreased geosmin biosynthesis.
An assessment of the apparent association between the stage of the Streptomyces life cycle
(i.e. the extent of morphological differentiation) and T&O production by Streptomyces spp. reported in the literature forms a major research objective of Chapter 6. Using a multivariate approach, this Chapter also comprises an examination of the influence of selected physico- chemical factors on the growth phase and T&O metabolite production by Streptomyces spp.
The premise for this study being that knowledge of the factors that trigger Streptomyces to cease vegetative growth and enter the reproductive developmental stage would conceivably allow elucidation of the conditions which also trigger significant production of their T&O metabolites.
2.4.2.2 Pathways of biosynthesis
Despite the identification of geosmin and 2-MIB in the late 1960s, their puzzling biosynthesis remained unsolved for several decades. By feeding radioactively labelled precursors to cultures of S. antibioticus, Bentley and Meganathan (1981) demonstrated that these compounds were terpenes, geosmin a degraded sesquiterpene and 2-MIB a methylated monoterpene. Two distinct pathways of terpenoid biosynthesis exist: the 2-C-methyl-D- erythritol 4-phosphate/1-deoxy-D-xylulose 5-phosphate pathway (MEP/ DOXP) or the mevalonic acid (MVA) pathway (Figure 2.3). Both pathways produce isopentenly diphosphate (IPP), the 5-carbon building block of terpene synthesis which forms geranyl diphosphate (GPP), the universal 10-carbon precursor of monoterpenes (e.g. 2-MIB) and farnesyl diphosphate (FPP), the 15-carbon precursor of sesquiterpenes (e.g. geosmin) (Lange
18 et al., 2000). The MEP pathway is believed to be the major biosynthetic pathway to produce these precursors in Streptomyces (Spiteller et al., 2002; Singh et al., 2009). Of recent years, a significant focus has been the elucidation of the genes and enzymes responsible for the final steps by which geosmin and 2-MIB are produced from these precursors.
Figure 2.3 Simplified biosynthetic scheme for the formation of geosmin and 2-MIB in Streptomyces. Modified from Juttner and Watson (2007).
Geosmin synthase has been identified as the enzyme responsible for the two-step production of geosmin. This bifunctional sesquiterpene synthase, first isolated from S. coelicolor A3(2), consists of 726 amino acids, encoded by the 2181-bp SCO6073 (cyc2) gene (Cane & Watt,
2003; Jiang et al., 2007). The enzyme’s two active sites. The N-terminal domain catalyses the cyclisation of FPP into the sesquiterpene alcohol (1(10)E,SE)-germacradien-11-ol
(germacradienol) and germacrene D (a bicyclic hydrocarbon). The highly homologous C- terminal active site diffusively rebinds the released germacradienol product and catalyses
19 proton initiated cyclisation-fragmentation of this molecule via the intermediate 8,10-dimethy
L-1-octalin to give geosmin. Both sites have an absolute dependence on the cofactor Mg2+, with each possessing two strictly conserved motifs for binding this cation (Cane et al., 2006;
Jiang et al., 2007). Homologous genes encoding geosmin synthases have been also identified in the genomes of S. avermitilis (SAV2163 or geoA, 725 aa) (Cane et al., 2006) and S. peucetius (Spterp13, 2,199bp, 732 aa) (Ghimire et al., 2008; Singh et al., 2009). A geosmin synthase encoding gene has also been identified from the cyanobacterium Nostoc punctiforme (Giglio et al., 2008). Citron et al. (2012) screened bacterial genomes for amino acid sequences with high similarity to the S. coelicolor A3(2) geosmin synthase gene and identified putative geosmin synthases in the genomes of 49 strains of bacteria, including all actinobacteria.
The pathway of 2-MIB biosynthesis in Nannocystis exedens was unravelled in feeding experiments by Dickschat et al. (2007) and the same enzymatic mechanism has been confirmed in Streptomyces and cyanobacteria (Wang & Cane, 2008; Giglio et al., 2011;
Wang et al., 2011). The pathway proceeds through the S-adenosylmethionine (SAM)- dependent methylation of GPP to (E)-2-methylgeranyl diphosphate (2-methyl-GPP) catalysed by a GPP-2-methyltransferase (2-methyl-GPP synthase) and its subsequent cyclisation to 2-MIB, catalysed by a C11 homo-monoterpene synthase (2-MIB synthase). In
S. coelicolor A3(2), these enzymes are encoded by two adjacent genes including SCO7701 and SCO7700 together with an upstream regulatory cyclic nucleotide binding protein (cnb) gene in a three-gene operon (Wang & Cane, 2008). These genes share highly conserved motifs for Mg2+ binding and recently, the X-ray crystal structures of both enzymes have been identified (Koksal et al., 2012a, 2012b). Komatsu et al. (2008) using bioinformatic analysis identified homologous genes constituting the 2-MIB synthase operon in the genome of other
Streptomyces capable of producing this compound. Homologous genes for the same 20 biosynthetic machinery have been also identified in cyanobacteria, suggesting that 2-MIB biosynthesis and the associated genes in these bacteria have a common origin (Komatsu et al., 2008; Giglio et al., 2011; Wang et al., 2011). Citron et al. (2012) conducted a BLAST search against the amino acid sequence of 2-MIB synthase from S. coelicolor (SCO7701), revealing the presence of homologues in 23 actinobacteria.
2.4.2.3 Purpose of biosynthesis
Considering literature assertions that the production of geosmin and 2-MIB coincides with
Streptomyces morphological differentiation and sporulation suggests that the possible biological function of these metabolites is related to the reproductive phase of the life cycle of these bacteria (Bentley & Meganathan, 1981; Dionigi et al., 1992; Scholler et al., 2002;
Tung et al., 2006). Similar to many other secondary metabolites, they may serve a defence strategy, to antagonise competing microorganisms in times of adversity (e.g. nutrient limitation) when their reproductive growth is initiated to ensure the survivability of the next generation of germinating spores (Hopwood, 2007). While the low-level toxicity of geosmin and 2-MIB to higher-order organisms is apparent from numerous studies (Nakajima et al.,
1996; Gagne et al., 1999; Mochida, 2009; Burgos et al., 2014), some authors have suggested they may exhibit antimicrobial activity. In high concentration they were found to inhibit the growth of Salmonella typhimurium and their antifungal effects have been reported (Dionigi et al., 1993; Wang et al., 2013a). Zaitlin and Watson (2006) maintain that the high concentrations used in these studies which greatly exceed those typically encountered in freshwater environments (<500 ng L-1) may be encountered by organisms in close proximity to source microorganisms or at the micro-scale level in sediments, soil and biofilms.
Although the water industry has tended to treat geosmin and 2-MIB as metabolic waste 21 products (Watson, 2003), it seems inconceivable that they could play no adaptive biological role given the complexity and energetic costs of their biosynthesis and the ubiquity of these compounds in nature. Chapter 7 attempts to further elucidate the possible biological function of these terpenoids as antimicrobial compounds as well as providing alternative adaptive roles in the life of Streptomyces.
2.5 A terrestrial origin of Streptomyces and T&O compounds?
As soil is a rich reservoir of Streptomyces, records of their occurrence in water samples must be considered critically with regards to their origin and in particular, whether or not they represent indigenous aquatic inhabitants or terrestrial wash-in forms (Cross, 1981). Periods of high runoff can introduce the soils of drainage basins, entrained with Streptomyces and their associated odorous secondary metabolites into surface waters causing T&O episodes
(Raschke et al., 1975; Niemi et al., 1982). The source of annual spring earthy-musty odour in the Saskatchewan River (Canada) was traced to high numbers of Streptomyces which, along with their odorous metabolites, were thought to be introduced into these surface waters during snowmelt and subsequent runoff (Jensen et al., 1994). In this case, the majority of recovered isolates (58%) were found to be producers of geosmin and/or 2-MIB. Furthermore, in the Elbow River Basin and Lake Ontario (Canada), Streptomyces abundance and geosmin and 2-MIB levels were associated with parameters indicative of terrestrial runoff (E. coli, turbidity and suspended sediment) (Zaitlin et al., 2003a; Zaitlin et al., 2003b). An assessment of this potentially significant means by which Streptomyces may contribute to T&O episodes in drinking water reservoir is presented in Chapter 4. Another focus of Chapter 4 concerned acquiring evidence to assess the possibility that these bacteria are able to actively grow in the water mass or submerged sediments of aquatic environments and are therefore capable of in
22 situ production of T&O metabolites. The following sections draw on literature evidence to date to support this proposition.
2.6 Streptomyces and T&O production in aquatic environments
The capacity of Streptomyces, among other actinobacteria to be metabolically active in aquatic environments has not yet been elucidated and many researchers argue that those isolated from such environments, are not truly aquatic, but rather an artefact of runoff and spore survival (Johnston & Cross, 1976a; Cross, 1981). Indeed Cross (1981) referred to aquatic Streptomyces as ‘mythical’ organisms. Wood et al. (1983a) argued that whilst it is plausible to consider the terrestrial wash-in hypothesis of Streptomyces, there seems no reason to assume that they cannot grow in aquatic environments, given suitable environmental conditions for their growth and activity. These bacteria readily grow and produce earthy-musty odour compounds in liquid media and there are numerous reports of
Streptomyces differentiation and sporulation occurring under such conditions (van Keulen et al., 2003; Manteca et al., 2008). Furthermore, gas vesicle protein (gvp) gene clusters with close homology to those found in cyanobacteria have been detected in the genome of
Streptomyces, implying that they may also possess gas-filled organelles that function as flotation devices. This suggests that these bacteria, primarily regarded as soil saprophytes, may also be physiologically adapted to the water environment (van Keulen et al., 2005b).
Actinobacteria have been isolated from samples taken many kilometres from land and at great oceanic depths and research using molecular techniques is providing more evidence supporting that they may be autochthonous and metabolically active inhabitants of marine communities (Jensen et al., 1991; Jensen et al., 2005; Bredholt et al., 2008). Moran et al.
(1995) extracted Streptomyces rRNA from coastal salt marsh sediment at concentrations 23 representing 2-5% of the sediment RNA. As dormant spores cannot produce rRNA, hybridisation analyses using Streptomyces-specific probes indicated they were actively growing. Isolates of the genus Salinispora were found to be present as vegetative forms in marine sediments at great depths (1,100m) by using a DNA extraction technique which excludes spores (Mincer et al., 2005).
What evidence do we currently have for their growth within freshwater environments?
Actinobacterial genera such as Streptomyces, Micromonospora, Actinoplanes, Rhodococcus and Thermoactinomyces have been frequently isolated from freshwater habitats including the waters, sediments and vegetation of lakes, reservoirs and rivers. However, the extent to which they represent a physiologically active and ecologically significant component of freshwater microbial communities remains unresolved. Such knowledge is critical to understand their capacity to contribute to in situ T&O production (Wood et al., 1983a; Zaitlin & Watson,
2006). Streptomyces, the main genus known to produce odorous metabolites, were described by Cross (1981) as “terrigenous species which occasionally find themselves in water” (p.
407). However, the isolation of several phages specific to Streptomyces from the surface of lake mud provided early evidence that they may be active in such environments (Willoughby,
1974). Micromonospora are generally more abundant in lake environments than
Streptomyces, where they have been regarded as a truly indigenous group of microbial inhabitants (Willoughby, 1969; Johnston & Cross, 1976a; Jiang & Xu, 1996). Supporting this, Johnston and Cross (1976a) found significant amounts of Micromonospora mycelium in lake sediments using homogenisation and sonication experiments, whereas Streptomyces were found to exist predominately as spores. They have been regarded as non-producers of
T&O in the literature, but recently one species of the genus (M. olivasterospora) was reported by Citron et al. (2012) to possess the gene encoding 2-MIB synthase and was confirmed to be a 2-MIB producer. 24
Species of the genus Actinoplanes have been isolated directly from river and lake water and have been found associated with allochthonous decomposing vegetation around lake edges and shallow streams (Willoughby, 1971; Willoughby et al., 1972; Johnston & Cross, 1976a;
Makkar & Cross, 1982). Makkar and Cross (1982) demonstrated that they have motile zoospores and Willoughby et al. (1972) found a phage specific to this genus from stream and lake water, which both serve as evidence for aquatic activity. There are currently no reports that this genus can produce T&O. Nocardia have been isolated from streams, rivers and lakes in low numbers, and have been assumed to originate from inactive spores of terrestrial origin (Willoughby, 1969; Johnston & Cross, 1976a). Three Nocardia isolates have been shown to produce geosmin in the laboratory (Gerber, 1979; Schrader & Summerfelt, 2010).
A strain of Microbispora rosea was shown by Gerber and Lecheval (1965) to produce geosmin in much higher concentrations (5.8 µg/mL) than the greatest yield by a Streptomyces
(1.75 µg/mL), but this genus has only been detected in very low numbers in lake sediments
(Johnston & Cross, 1976a). Gerber (1969) showed that a strain of Actinomadura could produce 2-MIB in culture, but the genus is not regarded as aquatic. Similarly, species of other genera including an Actinomyces from lake sediment (Kikuchi et al., 1973) and more recently strains of Actinosynnema, Kitasatospora, Rothia and Saccharopolyspora (Citron et al., 2012) have demonstrated geosmin and/or 2-MIB production in culture but have not been identified in T&O source tracking studies. It may be that conventional isolation and enumeration procedures have overlooked the occurrence of these non-Streptomyces and their potentiality to contribute to T&O events (Wood et al., 1983a).
Mere isolation of Streptomyces and other actinobacteria from a water source affected by earthy-musty T&O does not provide evidence that they are the causal organisms (Zaitlin &
Watson, 2006). The advent of culture-independent molecular techniques has made it possible to differentiate between actively growing vegetative forms and dormant spores in aquatic 25 environments. Fluorescence in situ hybridisation (FISH) targeting ribosomal RNA has been used to indicate the abundances of active filamentous actinobacteria (Sekar et al., 2003;
Klausen et al., 2005; Nielsen et al., 2006). The increased understanding of the functional genes involved in geosmin and 2-MIB biosynthesis may provide a more relevant tool to detect and track the emergence of T&O producing actinobacteria, particularly as methods such as plate counting and FISH cannot distinguish between producers and non-producers of these compounds. Auffret et al. (2011) reported on the use of primers targeting Streptomyces geosmin and 2-MIB synthase genes which combined with techniques such quantitative PCR
(qPCR), may be an effective means to measure the level of T&O metabolite producers prior to the occurrence of a T&O episode. More recently Du et al. (2013) developed a qPCR protocol using Streptomyces-specific primers targeting the geosmin synthase gene (geoA) to quantify and monitor the presence of geosmin producers in the Chinese-liquor making processes. Similarly, qPCR has been applied to quantify geosmin producing Anabaena and other cyanobacterial species in freshwater systems (Su et al., 2013; Kutovaya & Watson,
2014; Tsao et al., 2014b; Suurnakki et al., 2015). In these reports, geosmin concentrations were correlated with the bacterial populations detected by qPCR targeting the geosmin synthase gene, demonstrating the application of such protocols for monitoring potential T&O development.
Any attempt to source track and control T&O necessitates the identification of the causal organisms, their growth sites and conditions which trigger their production of these odorous secondary metabolites (Wood et al., 1983a). A review of current knowledge regarding
Streptomyces and other actinobacteria within freshwater environments and their association with T&O is presented in the following sections.
26
2.6.1 The water mass
The water mass of lakes, rivers and reservoirs has often been regarded as an unsuitable environment for the growth of actinobacteria with counts considered too low to contribute to
T&O production (Sugiura & Nakano, 2000; Juttner & Watson, 2007). However a source of nutrient enrichment may allow them to become metabolically active and important producers of T&O in the water mass (Bays et al., 1970). Wood et al. (1985) after demonstrating that oligotrophic reservoir water supported sparse growth of S. albidoflavus, showed that growth and geosmin production increased (3,900 ng/L) with the supplementation of carbon, nitrogen and phosphorus. Such nutrients could be obtained from suspended sediment material, plant debris, algae or from soil leachates. Some authors have detected high abundances of actinobacteria in the water mass during periods of low cyanobacterial biomass and have identified coincidences with T&O (Henatsch & Juttner, 1990; Lanciotti et al., 2003).
Molecular techniques are providing convincing evidence to support that actinobacteria are active and abundant microbial inhabitants in the water column and may contribute more to
T&O problems than previously expected. 16S rRNA sequencing of water samples from the cold and oligotrophic Lake Baikal (Russia) showed that up to 30% of the microbial population were actinobacteria (Denisova et al., 1999). A RNA hybridisation protocol
(FISH-CARD) also demonstrated high abundances of actinobacteria (32-55%) relative to the total bacterial planktonic communities in lakes of different trophic status (Sekar et al., 2003).
Using this technique, Nielsen et al. (2006) detected higher activity of actinobacteria (33-
49%) compared to other bacteria (27-39%) in the surface and bottom water of the North Pine
River Dam (Australia). Furthermore, high numbers of filamentous actinobacteria (1.3 x 108 cells/L) were detected, which suggests that Streptomyces may be metabolically active in water. In the same reservoir, rRNA analyses revealed that actinobacteria comprised 18-24% of all bacteria during a summer period of detectable geosmin and 2-MIB, with 5-10% of the 27 population estimated to be filamentous actinobacteria (Klausen et al., 2004). The low T&O producing cyanobacterial biomass (<1%) during this period suggested that actinobacteria were major T&O producers.
Recently, qPCR with 23S rRNA genus-specific primers was used to determine the abundance of Streptomyces in rivers and reservoirs in Southeast Queensland, Australia. Streptomyces were detected in all locations (average of 225 cells/L in surface waters and up to 45,650 cells/L at 8.5 m depth) aside from the deepest bottom waters (35-40m depth) which the authors attributed to anoxic conditions. However, based on the densities obtained and cell- specific production rates of geosmin and 2-MIB by Streptomyces determined by Klausen et al. (2005), the authors concluded that Streptomyces may not to be a major source of these odorous compounds (Lylloff et al., 2012).
2.6.2 Vegetation
Decaying vegetation and macrophyte communities along the margins of lakes, reservoirs and rivers have been considered to be a potential site of T&O production by actinobacteria
(Raschke et al., 1975; Cross, 1981; Makkar & Cross, 1982; Goodfellow & Williams, 1983).
This seems feasible given that their role as decomposers of complex plant polymers is firmly established. High counts of Streptomyces, Actinoplanes and Micromonospora have been obtained from plant debris adjacent to lakes and rivers (Raschke et al., 1975; Persson &
Sivonen, 1979; Makkar & Cross, 1982). Wood et al. (1985) found that significant concentrations of geosmin were produced when S. albidoflavus was grown on sterilised plant debris (180 ng/kg to 6,700 ng/kg) collected from the margins of English reservoirs. The wet- dry regime that this marginal habitat experiences may provide an ideal situation for their synthesis of T&O metabolites in aerobic conditions during dry periods, followed by their
28 subsequent wash-in by the action of rain, waves and rising water levels (Goodfellow &
Williams, 1983; Wood et al., 1983a).
Many rivers, natural lakes and reservoirs have large areas of shallow water which support extensive communities of submerged, emergent and floating macrophytes (Silvey & Roach,
1975). Besides their beneficial ecosystem services such as supplying oxygen, macrophytes provide an ideal habitat for actinobacteria, where they can obtain nutrients and grow and sporulate due to aerobic conditions as water levels fluctuate (Silvey & Roach, 1975; Cross,
1981). Actinobacteria, predominately Streptomyces, have been found to be particularly associated with dying macrophytes, with numbers increasing as vegetation begins to decompose (Silvey & Roach, 1975; Wohl & McArthur, 1998). Large populations have been observed growing on aquatic emergent vegetation such as Typha and Lemnain various reservoirs (Silvey et al., 1950; Bartholomew, 1958). Silvey and Roach (1975) refer to observations in a reservoir in East Texas, which experienced intense T&O and supported high populations of Chara and Najas species, with actinobacteria readily isolated from these macrophytes. Once the macrophyte populations were controlled at low densities, T&O problems disappeared. Zaitlin et al. (2003b) also detected actinobacteria associated with macrophytes (Valisnerai and Myriophyllum spp.) and overlying water samples (10-340 cfu/mL) collected in Lake Ontario (Canada), with some Streptomyces isolates capable of producing geosmin and 2-MIB.
2.6.3 Sediments and muds
Benthic sediments and muds in freshwater environments have been recognised as a possible habitat for actinobacterial growth and T&O production for some time (Adams, 1929;
Thaysen, 1936; Issatchenko & Egorova, 1944; Bays et al., 1970; Willoughby et al., 1972).
29
Sediments are an important reservoir of nutrients potentially available to actinobacteria, with their abundance often correlated with sediment nutrient status (Johnston & Cross, 1976b;
Sugiura et al., 1987; Jiang & Xu, 1996). Under anoxic conditions in sediments, they are unlikely to be metabolically active and produce T&O. Wood et al. (1985) found that the exposure of reservoir sediment during maintenance work led to earthy odour development, previously being undetectable in unexposed sediment. Furthermore, sterilised sediment was found to produce geosmin (460 ng/kg) when inoculated with S. albidoflavus. Shallow lakes with high summer temperatures and a supply of degradable organic matter would indeed provide ideal conditions for actinobacterial growth, particularly those with fluctuating levels, which provides aeration and therefore enables sporulation and secondary metabolism to occur (Cross, 1981). The notion that exposure of sediment stimulates the growth and T&O metabolite production by Streptomyces is investigated in the studies of Chapters 4 and 5.
Silvey and Roach (1975) suggest that the replenishment of oxygen to sediments during lake overturn may activate dormant actinobacteria, leading to T&O production. The authors claim that this phenomena has been observed in many water supplies in the USA but such a potential mechanism of T&O production is yet to be systematically investigated (Zaitlin &
Watson, 2006). Interestingly, Guttman and Rijn (2008) showed that although aerobic conditions supported higher geosmin and 2-MIB production, low concentrations of these compounds were also observed in anoxic cultures of Streptomyces spp. which were thought to be using nitrate as an electron acceptor.
T&O source tracking studies using RNA hybridisation techniques have found that actinobacteria may be metabolically active residents of sediment communities. Klausen et al. (2004) found a coincidence between high geosmin and 2-MIB concentrations and high abundance of actinobacteria (20-50 x 106 cells/g) in the upper 2cm of surface sediment in the
North Pine Dam (Australia). Nielsen et al. (2006) also detected a 35% to 80% higher 30 proportion of active actinobacterial cells in the surface sediment of the same reservoir and both authors suggest that the aeration systems deployed in the dam combined with organic matter availability provides an ideal habitat for actinobacteria and therefore T&O production. In hypereutrophic Lake Kasumigaura (Japan), T&O episodes have been associated with actinobacteria growing in benthic sediments. Sugiura et al. (1987) found that counts correlated with increasing musty (2-MIB) odour and organic carbon content and
Sugiura et al. (1994) attributed the actinobacterial odour production to their utilisation of sedimented algae and cyanobacteria as carbon substrates. In the same lake, Sugiura and
Nakano (2000) found that sedimented actinobacteria (up to 16,000 cfu/g) correlated with geosmin concentration with 40 isolates confirmed as producers. Several other studies have isolated geosmin and/or 2-MIB producing actinobacteria from sediment and have regarded them as the cause of earthy-musty odour (Zaitlin et al., 2003b; Tung et al., 2006; Zuo et al.,
2009a; Zuo et al., 2010; Lee et al., 2011). In order to determine their contribution to T&O, it is critical to understand whether they are metabolically active in this habitat and elucidate the environmental factors which stimulate their production of these secondary metabolites.
2.6.4 Other aquatic habitats
Drinking water distribution systems may be oligotrophic habitats, however oxygenated water and areas of slow flow where sediments and organic debris accumulate, may provide safe havens for biofilm establishment and optimal conditions for chronic T&O production by actinobacteria (Geldreich, 2002; Lanciotti et al., 2003). Counts of up to 4.4 x 105 cfu/L have been found in old soft deposits, three orders of magnitude greater than in water leaving the waterworks (Zacheus et al., 2001) and Silvey and Roach (1975) reported high numbers
(180,000-135,000 cfu/mL) in distribution systems during warm temperatures alongside elevated T&O. Streptomyces spores, being relatively resistant to chlorination readily breakthrough to the distribution system which may be an important site of T&O production 31
(Jensen et al., 1994). Uwins (2011) however, found no evidence to support the supposition that geosmin and 2-MIB were being produced by biofilm-attached or free actinobacteria in the reticulation system of the Gold Coast, Australia.
Evidence suggests that Streptomyces may live in association with filter feeding mussels and produce T&O. In the North American Great Lakes, T&O problems have coincided with the occurrence of heavy exotic Zebra Mussel (Dreissena) infestations. Geosmin and 2-MIB have been detected in concentrated Dreissena faeces/pseudofaceces and interstitial material which also supported the growth of Streptomyces (Lange & Wittmeyer, 1997). Similarly, Zaitlin et al. (2003b) detected geosmin and several isolates of Streptomyces in mussel and overlying water samples in Lake Ontario. The organic material provided by mussel filter feeding and dissolved oxygen from siphon activity may provide ideal conditions for Streptomyces to thrive in mussel beds and produce T&O compounds.
Actinobacteria have been isolated from periphyton and this complex mixture of microorganisms and detritus growing attached to submerged surfaces, has been identified as a source of geosmin and 2-MIB by several authors (Zaitlin et al., 2003b; Watson & Ridal,
2004; Ridal et al., 2007). Its major constituents (algae and cyanobacteria) can act as a growth substrate for actinobacteria which may support geosmin and 2-MIB production (Schrader &
Blevins, 1999). T&O problems have been ascribed to Streptomyces growing in the decaying filaments of algal blooms in surface waters and lyophilised cyanobacteria have been shown to support odour production by benthic Streptomyces isolates (Silvey & Roach, 1953;
Sugiura et al., 1994). However as cyanobacteria can also produce these compounds, their cell-bound odorous metabolites may be the major sources of T&O upon decomposition
(Juttner & Watson, 2007).
32
In addition to drinking water supplies, geosmin and 2-MIB are problematic in aquaculture where they bioaccumulate in the lipid rich tissues of fish creating great economic losses
(Guttman & van Rijn, 2008). Several studies have pointed to actinobacteria as the likely offenders of this tainting including Nocardia and Streptomyces species isolated from the water and sludge of aerobic sections in recirculating aquaculture systems (RAS) (Guttman & van Rijn, 2008; Schrader & Summerfelt, 2010). Pan et al. (2009) isolated species of
Streptomyces, Nocardia, Micromonospora and Actinomadura (counts up to 1.5 x 106 cfu/mL) from the odour affected brackish waters of intensive cultivation fishponds in Tianjin (China), with one Streptomyces isolate capable of producing large amounts of 2-MIB. Given the significant positive correlation between actinobacterial biomass and 2-MIB concentration in the ponds, they were regarded as responsible for the T&O. Recently, Auffret et al. (2013) monitored the abundance of the Streptomyces geosmin synthase gene (geoA) using qPCR in two RAS units. Although geoA was detected and coincided with higher geosmin concentrations in fish flesh, Streptomyces 16SrRNA gene sequences where not identified.
Sequences associated with other geosmin producers including Sorangium and Nannocystis
(Myxococcales) were detected, and geoA qPCR fragments generated were more related to the amino acids sequences from these taxa. Thus Myxobacteria may be an underestimated source of off-flavour compounds in aquaculture and indeed, drinking water supplies.
2.7 Conclusions
Evidence for the widespread distribution, abundance and activity of Streptomyces and other actinobacteria in natural and man-made aquatic environments has been appraised. Cultivars of these bacteria isolated from freshwater habitats including sediment, vegetation, the water mass or more specialised substrates (e.g. mussels or periphyton) readily demonstrate geosmin and 2-MIB producing abilities in vitro and may indeed be potent sources of earthy-musty
33
T&O. To elucidate the contribution of these bacteria to this aesthetic water quality problem, more research is required to verify their abundance and capability to be metabolically active in such habitats. Culture-dependent techniques alone cannot achieve this, being constrained by both the unculturability of many Streptomyces and the inability to discriminate between active and dormant forms. The development and application of nucleic-acid molecular methods provides a research avenue to address this knowledge gap.
The research presented in this thesis focuses on an investigation of the activity, abundance, diversity and geosmin and 2-MIB producing ability of Streptomyces in drinking water reservoir habitats using both a molecular-based protocol and culture-dependent assays to elucidate the potentiality of Streptomyces to contribute to T&O episodes. Furthermore, the experimental work attempts to build on previous studies which have investigated the influence of environmental factors on production of geosmin and 2-MIB by Streptomyces, the relationship of this production with life cycle stage and the biological purpose of production, in order to contribute to the body of knowledge concerning T&O production by these filamentous bacteria. Prior to the chapters pertaining to these research avenues, an analysis of historical water quality data establishes the ambiguity of the mechanisms that have triggered past T&O episodes in drinking water reservoirs (Grahamstown and Chichester
Reservoirs). This analysis presented in Chapter 3 provides a strong rationale for further investigation of the potential role of Streptomyces in T&O episodes.
34
CHAPTER 3 - EXPLORATORY MULTIVARIATE MODELLING OF PAST GEOSMIN AND 2-METHYLISOBORNEOL EPISODES IN GRAHAMSTOWN AND CHICHESTER DRINKING WATER RESERVOIRS
3.1 Introduction
The organoleptic quality of drinking water is an important issue for suppliers of potable water worldwide. It has long been recognised that consumers associate T&O in drinking water with its safety and acceptability for consumption (Srinivasan & Sorial, 2011). As emphasised in
Chapter 2, geosmin and 2-MIB are responsible for the majority of biologically caused T&O problems worldwide, with numerous organisms ranging from prokaryotes (e.g. cyanobacteria and actinobacteria) to eukaryotes (e.g. fungi) being implicated in the production of earthy- musty odours in drinking water supplies. Despite the substantial body of information that has accumulated, knowledge about the cause of T&O involving geosmin and 2-MIB remains poorly understood throughout the drinking water industry (Juttner & Watson, 2007).
Although earthy-musty T&O events are problematic and occur frequently in Australian reservoirs, there are few documented reports concerning these events in Australia. A major focus of geosmin and 2-MIB research in Australia has been on treatment and removal of these compounds (Newcombe & Cook, 2002; Ho et al., 2007).
Hunter Water Corporation (HWC) among other water utilities in Australia, often receive consumer complaints concerning earthy-musty T&O in drinking water. Two primary drinking water storages managed and operated by HWC are Grahamstown and Chichester
Reservoirs located in the Hunter Region of New South Wales, Australia. Both are prone to
T&O problems, having experienced numerous episodes of elevated concentrations of geosmin and 2-MIB above threshold levels of detection by humans (>10 ng/L). The dynamics
35 of the concentration of these compounds recorded weekly in Grahamstown (1998-2012) and
Chichester (2002-2012) are presented in Figures 3.1 and 3.2 respectively. HWC has been monitoring concentrations of geosmin and 2-MIB in the raw water intake of these freshwater storages for over a decade and it is apparent that the main compound accounting for T&O problems in both reservoirs is geosmin, although several cases of elevated concentration of
2-MIB have been recorded in Grahamstown Reservoir. Past T&O episodes have been largely unanticipated by HWC and there is a heavy reliance on the use of powdered activated carbon
(PAC) dosing to remove these chemically stable compounds through adsorption, which is an expensive management option.
300 1 4 6 Geosmin 250 2-MIB
200
150
2 100 7 Concentration Concentration (ng/L) 50 1 2 3 3 5
0
12/2005 10/2008 02/1998 07/1998 12/1998 05/1999 10/1999 03/2000 08/2000 01/2001 06/2001 10/2001 03/2002 08/2002 01/2003 06/2003 11/2003 04/2004 09/2004 02/2005 07/2005 05/2006 10/2006 03/2007 07/2007 12/2007 05/2008 03/2009 08/2009 01/2010 06/2010 11/2010 04/2011 09/2011 02/2012 Date Figure 3.1 Geosmin and 2-MIB concentrations in Grahamstown Reservoir from 1998 to 2012. Measurements have been taken from the raw water intake at the George Schroder pump station. The isolated episodes of elevated concentrations of each compound subjected to statistical analysis in this chapter are indicated with numbers.
36
180 Geosmin 160 4 2-MIB 140
120
100
80 1 60
Concentration Concentration (ng/L) 40 2 3 20
0
12/2007 04/2009 09/2002 01/2003 04/2003 07/2003 10/2003 02/2004 05/2004 08/2004 12/2004 03/2005 06/2005 09/2005 01/2006 04/2006 07/2006 11/2006 02/2007 05/2007 08/2007 03/2008 06/2008 10/2008 01/2009 07/2009 11/2009 02/2010 05/2010 09/2010 12/2010 03/2011 07/2011 10/2011 01/2012 04/2012 Date Figure 3.2 Geosmin and 2-MIB concentrations in Chichester Reservoir from 2002 to 2012. Measurements have been taken from the raw water intake. The isolated episodes of elevated geosmin concentration subjected to statistical analysis in this chapter are indicated with numbers.
In addition to geosmin and 2-MIB, a large number of other chemical, physical and biological parameters have been routinely measured in both reservoirs by HWC as part of a water quality monitoring scheme. The extensive data sets containing records of these water quality parameters in both reservoirs were analysed in this study. The aim of the analysis was to develop exploratory models to examine variables related to elevated concentrations of geosmin and 2-MIB in the reservoirs. This preliminary study was considered important in order to gain insight into the processes associated with the development of past T&O episodes in the reservoirs prior to commencing the research concerning the potential role of
Streptomyces in contributing to this aesthetic water quality issue.
Recently, several authors have reported the use of statistical models in an attempt to determine the underlying relationship between various water quality parameters and earthy-
37 musty secondary metabolites in fresh water systems. Various linear (multiple linear regression and principal components analysis) and non-linear (artificial neural networks) regression models have been employed for the purpose of predicting geosmin and 2-MIB based on chemical, physical and biological water quality parameters (Sugiura et al., 2004;
Dzialowski et al., 2009; Parinet et al., 2010; Qi et al., 2012). Such studies have revealed the large number of different factors associated with T&O events in diverse freshwater systems, indicating their complexity and variability.
It is often difficult to analyse large and complex environmental databases for meaningful interpretation, requiring reduction methods to simplify the data structure before useful and interpretable information can be extracted to explain the temporal variations of the variables.
Partial least squares (PLS) is a multiple regression method that focuses on maximising the variance of the dependent variables explained by the independent variables. That is, finding a relationship between independent (predictor) and dependent (response) data. It projects input-output data into a latent space, extracting principal components with an orthogonal structure, while capturing most of the variance in the original data. PLS differs from multiple linear regression (MLR), being a powerful multivariate modelling approach for its ability to analyse data with strongly collinear, noisy and numerous variables and missing values in the predictor (X) and response (Y) matrices. It is thus an ideal statistical tool for data mining
(Wold et al., 2010).
In this chapter, PLS analyses were carried out as an exploratory analysis tool with the aim to identify various routinely measured biological, physical or chemical water quality parameters related to geosmin and 2-MIB using historical data sets for Grahamstown and Chichester drinking water reservoirs. Although this statistical technique is predominately used for predicting dependent variables through modelling input-output relationships in the data, for 38 the purpose of this study, the tool was employed for exploratory rather than predictive purposes. PLS analyses were conducted to model geosmin and 2-MIB using the entire data sets in addition to data corresponding to selected isolated geosmin and 2-MIB episodes as indicated in Figures 3.1 and 3.2. PLS models can increase in predictive power when dealing with a smaller data set (i.e. isolated events) compared to models created for an entire large data set. It was considered that water quality parameters that revealed consistent relationships with geosmin or 2-MIB based on comparisons of PLS models developed for isolated events and for the entire data set of these compounds, would be associated with and may have a significant influence on the development of T&O problems. The PLS procedure is described in detail by Geladi and Kowalksi (1986). An overview of the mechanics of PLS analysis can be found in Appendix A.
3.2 Methods
3.2.1 Description of the drinking water reservoirs
The location of HWC’s drinking water supplies which have suffered T&O outbreaks involving geosmin and 2-MIB are shown on the map in Figure 3.3. Grahamstown Reservoir, located approximately 20 km north of Newcastle, NSW (32°44´S, 151°49´E), is an off-river storage. It is the primary raw water source for drinking water production in the Lower Hunter
Region, supplying 40% of its total volume. It has an average depth of 9 m, maximum depth of 11 m, a total capacity of 182,305 ML and surface area of 28 km2. Half of its water is pumped from the Williams River by the Balickera Canal and pumping station and the remainder from its own small catchment. The catchment is primarily located in the northern and eastern shores of the reservoir and receives a mixture of runoff from creeks draining through forested and small to medium sized farm allotments as well as directly from urban settlements located to the east and indirectly though Campvale Swamps. The land use
39 breakdown of the catchment is as follows: 39% rural, rural residential or urban, 39% HWC freehold and 22% state forest. Water stored in Grahamstown Reservoir is accessed for supply to customers at George Schroder pump station at the southern end, where it is distributed for treatment (Cole & Williams, 2011).
Figure 3.3 Map of study sites showing the location of Grahamstown and Chichester Reservoirs. Insert shows the state of New South Wales, Australia (Cole & Williams, 2011). 40
Chichester Reservoir, located 80 km north of Newcastle (32°13´S, 151°41´E) at the top of the Williams River Catchment, is also one of HWC’s important freshwater storage facilities, contributing about 35% of the Lower Hunter Region’s potable water supply. After some onsite treatment, water is transported via a gravity pipe to Dungog for treatment, after which half gravitates further to Newcastle for blending with water sourced from Grahamstown
Water Treatment Plant. In contrast to Grahamstown, it is a deeper reservoir (maximum depth
37 m), with a smaller surface area (1.8 km2), storage capacity (18,356 ML) and catchment size (199 km2). In addition, its catchment differs, largely being within the Barrington Tops
National Park which is declared a wilderness area. As a result, it is one of the most pristine catchments in Australia, largely unaffected by human activity. The land use breakdown is as follows: 76% forest (national park), 17% rural and 7% HWC freehold. The reservoir is fed by the Wangat River to the north with a catchment entirely vegetated and pristine and the
Chichester River to the north-west, which is partially cleared for agricultural and rural residential land use. There is high runoff from the area due to the abundant rainfall and the large catchment area. Hence the reservoir is filled quickly following medium to heavy rain
(Cole & Williams, 2011).
3.2.2 Water quality data sets
The data used in the development of the PLS models in this study were the routine monitoring information collected for water quality assessment by HWC. These data sets have been generated by continuous monitoring of water quality in Grahamstown (1998-2012) and
Chichester (2002-2012) Reservoirs. Measurements of geosmin and 2-MIB, in addition to most other chemical, physical and biological parameters have been recorded predominately on a weekly basis. Those parameters recorded more frequently were converted to weekly averages and rainfall was recorded as the weekly sum. The parameters included in the analyses are listed in Table 3.1. The routine monitoring programs have included
41 phytoplankton identification and enumeration to genus level. Not all genera were included in the analysis, and for the non-cyanobacterial parameters, total counts at higher taxonomic levels (e.g. class) were used. For cyanobacteria, genera which are frequently or at least periodically abundant were included in the analysis.
Table 3.1 Chemical, biological and physical water quality parameters in the historical water quality data sets used as explanatory variables in the PLS models. Water quality parameter Abbreviation Unit Water temperature Water temp °C Air temperature Air temp °C pH pH -
Alkalinity Alk CaCO3 mg/L Dissolved oxygen DO mg/L Rainfall (sum weekly) Rain mm Water level Level m AHD Secchi disk depth SDD m Electrical conductivity EC µS/cm Suspended solids SS mg/L Turbidity Turb NTU Thermotolerant coliforms FC cfu/100mL Heterotrophic bacteria HB cfu/100mL Total organic carbon TOC mg/L Total nitrogen TN mg N/L Dissolved inorganic nitrogen DIN mg N/L + Free ammonia NH4 mg N/L Total Kjeldahl nitrogen TKN mg N/L - Nitrate NO3 mg N/L
Nitrogen oxide NOx mg N/L Total phosphorus TP mg P/L Soluble reactive phosphorus SRP mg P/L Metals Fe, Mg, Mn, Ca mg/L Silica Si mg/L
42
Water quality parameter Abbreviation Unit 2- Sulfate SO4 mg/L Chlorophyll α Chl-α µg/L Pheophyton Pheo µg/L Total Cyanobacteria Total Cyano Cells/mL Potentially toxic PT Cyano Cells/mL Cyanobacteria Non-toxic Cyanobacteria NT Cyano Cells/mL Cyanobacterial genera Anabaena, Microcystis, Aphanocapsa, Cells/mL Aphanothece, Coelosphaerium, Cyanogranis, Cyanothece, Cyanocatena, Cyanodictyon, Cyanonephron, Planktolyngbya. Zooplankton Zooplankton Counts/mL Bacillariophyta (diatoms) Bacillarioph. Counts/mL Chlorophyta (green algae) Chlorophy. Cells/mL Chrysophyta (golden brown Chrysoph Cells/mL algae) Cryptomonads Cryptophy. Cells/mL Dinoflagellates (Dinophyta) Dinoph. Cells/mL Euglenoids (Euglenophyceae) Euglenoph. Cells/mL
3.2.3 Statistical analysis
PLS modelling was performed using SIMCA-P+ version 12.0 (Umetrics AB). In order to transform the data into a form suitable for analysis, all variables were pre-processed using the SIMCA-P+ default settings including mean centering and autoscaling to unit variance
(UV). The former centres variables by subtracting their averages whereas autoscaling to UV gives all of the variables axes of equal length and thus equal importance (i.e. puts variables on the same scale).
The variance in Y (geosmin or 2-MIB) that is explained by the model was expressed as R2, that is, how well the model fits the data. The variance in Y according to cross-validation was 43 expressed as Q2, indicating how well the model predicts new data. Cross-validation, which determines the robustness of the models, was evaluated by using leave one out cross validation (LOO-CV). The SIMCA-P+ default for LOO-CV is the removal of every 1/7th of the data during development of the model, with the model being built on 6/7th of the data.
The left out data are predicted from the model and compared with the actual values to calculate the sum of square error. This is then called the predicted residual sum of square
(PRESS). The lower this value is, the better the predictability of the model. In SIMCA-P+,
PRESS is converted into Q2 to resemble the scale of R2. Good predictions will have low
PRESS and therefore high Q2. The optimum number of latent variables in the PLS models constructed was selected on the basis of the percent variance captured by the models. The autofit facility of SIMCA-P+ extracts as many components as considered significant for the developed model. Variables that did not vary or contained too many missing values were excluded in the model development as recommended in the program.
3.3 Results
3.3.1 Overview of PLS models
PLS analysis was conducted to explore if variations in any routinely measured chemical, physical and biological parameters in Grahamstown and Chichester drinking water reservoirs were strongly related to variations in geosmin and 2-MIB levels. An overview of the PLS models developed, based on the entire data sets and data corresponding to selected isolated
T&O events are presented in Tables 3.2 and 3.3 for Grahamstown and Chichester Reservoir respectively. R2 values, which indicate how well the model fits the data, are shown for each principal component in addition to the predictive ability of the model (Q2) according to cross- validation. For biological PLS models, R2 and Q2 values of >0.5 and >0.4 respectively are regarded as acceptable for a good model (Umetrics, 2012). It can be seen that the first
44 principal component of the majority of the models explains a sufficient amount of the variation (>50%) in geosmin or 2-MIB and hence loading plots were only analysed for the first model components. The exceptions were the PLS models developed for Grahamstown
2-MIB and Chichester geosmin concentration based on the entire data sets, and geosmin event 1 in both reservoirs which captured lower variation.
Table 3.2 Overview of the fit and number of principal components of each PLS model for geosmin and 2-MIB in Grahamstown Reservoir. The percent variation of the dependent variable (geosmin or 2-MIB) explained by the model (R2) and the percent variation of the dependent variable predicted by the model according to cross validation (Q2) are shown.
Model name
Component 1 Component 2 Component 3 Cumulative
Component 1 Component 2 Component 3 Cumulative
2 2 2 2
2 2 2 2
R R R R Q Q Q Q Geosmin Event 1 0.478 - - 0.478 0.453 - - 0.453 Geosmin Event 2 0.560 0.182 - 0.738 0.524 0.275 - 0.655 Geosmin Event 3 0.691 0.093 - 0.784 0.606 0.195 - 0.682 Geosmin Event 4 0.775 0.068 - 0.843 0.693 0.115 - 0.723 Geosmin Event 5 0.605 - - 0.605 0.503 - - 0.503 Geosmin Event 6 0.564 0.144 0.104 0.812 0.449 0.138 0.093 0.569 Geosmin Event 7 0.510 - - 0.510 0.421 - - 0.421 All geosmin data 0.557 0.122 0.045 0.724 0.537 0.252 0.115 0.693 2-MIB Event 1 0.709 0.177 0.028 0.913 0.696 0.592 0.164 0.897 2-MIB Event 2 0.715 0.104 - 0.820 0.643 0.217 - 0.720 2-MIB Event 3 0.518 0.137 - 0.654 0.450 0.088 - 0.498 2-MIB data 0.136 0.042 - 0.178 0.134 0.041 - 0.170
45
Table 3.3 Overview of the fit and number of principal components of each PLS model for geosmin in Chichester Reservoir. The percent variation of the dependent variable explained by the model (R2) and the percent variation of the dependent variable predicted by the model according to cross validation (Q2) are shown.
Model name
nent 2
Component 1 Compo Component 3 Cumulative
Component 1 Component 2 Component 3 Cumulative
2 2 2 2
2 2 2 2
R R R R Q Q Q Q Geosmin Event 1 0.409 - - 0.409 0.257 - - 0.257 Geosmin Event 2 0.735 - - 0.735 0.678 - - 0.678 Geosmin Event 3 0.813 0.088 0.051 0.952 0.660 0.368 0.140 0.815 Geosmin Event 4 0.523 - - 0.523 0.414 - - 0.414 All geosmin data 0.385 0.092 0.040 0.517 0.363 0.140 0.060 0.485
PLS analysis results in model coefficients for the variables, referred to as weights or loadings.
The weights for the X-variables (W) indicate the importance of these variables, that is, how much in a relative sense, they participate in the modelling of Y (C). The loading plots presented in the following sections with W*C[1] as the y-axes, therefore show the correlation structure and specify the direction of associations between the various chemical, physical and biological parameters and either geosmin or 2-MIB as the response variable. Loadings do not exactly correspond to correlation coefficients, but position the variables in the same way with regards to factors. Variables situated around zero have little significant contribution in explaining geosmin or 2-MIB and variables with approximately equal loadings may be collinear (Singh et al., 2007). The primary purpose of PLS modelling of geosmin and 2-MIB events in both reservoirs was to identify the independent variables (water quality parameters) which show consistent relationships with these compounds. Loading plots were analysed to graphically interpret the models to achieve this objective.
46
3.3.2 PLS models for geosmin in Grahamstown Reservoir
In Grahamstown Reservoir, the cyanobacterial genus Anabaena was overall the most strongly associated variable with geosmin, revealing the highest positive loading in the model based on the entire data set (Figure 3.4). Other phytoplankton parameters also showed positive but comparatively weaker loadings. The PLS models developed for the selected seven isolated geosmin events (Figures 3.5 to 3.11) generally revealed the positive relationship between geosmin and Anabaena, although the strength of the association was variable, being greatest for the models corresponding to geosmin episodes of greatest concentration (events 4 and 6). The time series graph in Figure 3.12 clearly shows the close relationship between the dynamics of geosmin concentration and Anabaena abundance in
Grahamstown Reservoir. The PLS model based on the entire data set revealed that geosmin had the strongest inverse relationship with DO. The negative relationship with DO was also apparent for geosmin events 2, 4, 6 and 7 and is shown in the times series plot in Figure 3.13.
Water temperature was also positively associated with geosmin in these models and the entire data set model, but this variable had a negative loading or no association with geosmin in the other models. Aside from Anabaena and DO, the PLS models for geosmin events generally revealed different patterns of associations of variables with the compound. Nutrient parameters did not reveal any consistent relationship with geosmin.
47
Figure 3.4 Loading column plot for the first principal component in the PLS model developed for geosmin based on the entire data set for Grahamstown Reservoir.
Figure 3.5 Loading column plot for the first principal component in the PLS model developed for the first geosmin event in Grahamstown Reservoir.
48
Figure 3.6 Loading column plot for the first principal component in the PLS model developed for the second geosmin event in Grahamstown Reservoir.
Figure 3.7 Loading column plot for the first principal component in the PLS model developed for the third geosmin event in Grahamstown Reservoir.
49
Figure 3.8 Loading column plot for the first principal component in the PLS model developed for the fourth geosmin event in Grahamstown Reservoir.
Figure 3.9 Loading column plot for the first principal component in the PLS model developed for the fifth geosmin event in Grahamstown Reservoir.
50
Figure 3.10 Loading column plot for the first principal component in the PLS model developed for the sixth geosmin event in Grahamstown Reservoir.
Figure 3.11 Loading column plot for the first principal component in the PLS model developed for the seventh geosmin event in Grahamstown Reservoir.
51
5000 250 Geosmin (ng/L) 4500 2-MIB (ng/L) 4000 200 Anabaena (cells/L) 3500
150 3000 2500
100 2000 Cells/mL 1500
Concentration (ng/L) Concentration 50 1000 500
0 0
08-2002 02-2007 02-1998 08-1998 02-1999 08-1999 02-2000 08-2000 02-2001 08-2001 02-2002 02-2003 08-2003 02-2004 08-2004 02-2005 08-2005 02-2006 08-2006 08-2007 02-2008 08-2008 02-2009 08-2009 02-2010 08-2010 02-2011 08-2011 02-2012 Date Figure 3.12 Grahamstown Reservoir geosmin and 2-MIB concentration and Anabaena abundance. Geosmin peaks marked with an arrow correspond to those coinciding with low Anabaena abundance.
2-MIB 300 14 Geosmin DO 250 12
10 200 8 150 6 100
4 Concentration (ng/L) Concentration 50 2 (mg/L) oxygen Dissolved
0 0
02/1998 09/1998 03/1999 10/1999 04/2000 11/2000 06/2001 12/2001 07/2002 01/2003 08/2003 02/2004 09/2004 04/2005 10/2005 05/2006 11/2006 06/2007 12/2007 07/2008 01/2009 08/2009 03/2010 09/2010 04/2011 10/2011 Date Figure 3.13 Grahamstown Reservoir geosmin, 2-MIB and dissolved oxygen concentration.
52
3.3.3 PLS models for 2-MIB in Grahamstown Reservoir
Loading plots for the PLS models developed to explore water quality parameters related to
2-MIB in Grahamstown Reservoir based on the entire data set and the three major isolated events are shown in Figures 3.14 to 3.17. A parameter with which 2-MIB levels showed a strong and consistent association was water level, which was negatively related to 2-MIB.
The times series graph in Figure 3.18 reveals how periods of elevated 2-MIB have coincided with lower water levels. Similar to its relationship with geosmin levels, DO also appeared to be negatively associated with 2-MIB levels based on the entire data set, and in the models developed for events 1 and 3, but did not appear to be an associative factor in relation to event 2. Silica also had a negative loading on the PLS models developed for all isolated 2-
MIB events, although this parameter revealed a positive loading on the model built with the entire data set. Water temperature was positively associated with 2-MIB, with the exception of event number 2. Electrical conductivity showed a strong positive relationship with 2-MIB based on the entire data set and weaker positive associations in events 1 and 2, but was negatively related to 2-MIB in event 3. None of the biological parameters examined showed consistent associations with 2-MIB levels. The cyanobacterial genera Microcystis and
Planktlyngbya revealed the strongest positive associations with 2-MIB in models developed for event 1 and 2 respectively. However, the time series plots of 2-MIB with these genera
(Figure 3.19), which are generally in low abundance during the 2-MIB peaks, do not show a consistent or strong relationship. High levels of another, more abundant cyanobacterial genus, Aphanocapsa, coincided with the second 2-MIB peak, having a positive loading on the second 2-MIB event PLS model.
53
Figure 3.14 Loading column plot for the first principal component in the PLS model developed for 2-MIB based on the entire data set for Grahamstown Reservoir.
Figure 3.15 Loading column plot for the first principal component in the PLS model developed for the first 2-MIB event in Grahamstown Reservoir.
54
Figure 3.16 Loading column plot for the first principal component in the PLS model developed for the second 2-MIB event in Grahamstown Reservoir.
Figure 3.17 Loading column plot for the first principal component in the PLS model developed for the third 2-MIB event in Grahamstown Reservoir.
55
300 2-MIB 14 Geosmin Level 250 13
200 12
150 11
100 10
Level (m AHD) (m Level Concentration Concentration (ng/L) 50 9
0 8
02/1998 09/1998 03/1999 10/1999 04/2000 11/2000 06/2001 12/2001 07/2002 01/2003 08/2003 02/2004 09/2004 04/2005 10/2005 05/2006 11/2006 06/2007 12/2007 07/2008 01/2009 08/2009 03/2010 09/2010 04/2011 10/2011 Date Figure 3.18 Grahamstown Reservoir geosmin and 2-MIB concentration and water level.
3.3.4 PLS models for geosmin in Chichester Reservoir
As observed in the PLS models developed for geosmin in Grahamstown Reservoir,
Anabaena was the only biological parameter that consistently showed a positive relationship with geosmin in Chichester Reservoir for the model built using the entire data set (Figure
3.20) in addition to the four isolated geosmin events analysed (Figures 3.21 to 3.24).
Anabaena abundance and geosmin concentration through time are shown in Figure 3.25 and reveal the close relationship between the dynamics of this genus and compound in Chichester
Reservoir. The strength of positive association of geosmin levels with Anabaena abundance varied between models, being strongest in the model built with the entire data set, followed by the isolated event of longest duration and with the highest geosmin concentration (event
4). Silica revealed the overall greatest negative loading, but was not a variable associated with geosmin in event 1. Water level was also negatively related to geosmin based on the entire data set and events 2 and 4, but this variable was not significant for other geosmin events in the reservoir. Turbidity was found to consistently have a negative loading on all
56
300 8,000 Geosmin 2-MIB Microcystis 250 7,000 6,000 200 5,000 150 4,000 3,000
100 Cells/mL 2,000 50 1,000
Concentration (ng/L) Concentration 0 0
02/1998 02/2002 08/1998 02/1999 08/1999 02/2000 08/2000 02/2001 08/2001 08/2002 02/2003 08/2003 02/2004 08/2004 02/2005 08/2005 02/2006 08/2006 02/2007 08/2007 02/2008 08/2008 02/2009 08/2009 02/2010 08/2010 02/2011 08/2011 02/2012
Date
300 1,000,000 Geosmin 2-MIB Planktolyngbya 250 100,000
200 10,000
150 1,000
100 100 Cells/mL
50 10 Concentration (ng/L) Concentration
0 1
12/2010 02/1998 09/1998 04/1999 11/1999 06/2000 01/2001 08/2001 03/2002 10/2002 05/2003 12/2003 07/2004 02/2005 09/2005 04/2006 11/2006 06/2007 01/2008 08/2008 03/2009 10/2009 05/2010 07/2011 02/2012
Date
300 350,000 Geosmin 2-MIB Aphanocapsa 250 300,000 250,000 200 200,000 150 150,000
100 Cells/mL 100,000
50 50,000 Concentration (ng/L) Concentration
0 0
02/1998 09/1998 04/1999 11/1999 06/2000 01/2001 08/2001 03/2002 10/2002 05/2003 12/2003 07/2004 02/2005 09/2005 04/2006 11/2006 06/2007 01/2008 08/2008 03/2009 10/2009 05/2010 12/2010 07/2011 02/2012
Date
Figure 3.19 Grahamstown Reservoir geosmin, 2-MIB and abundance of the cyanobacterial genera Microcystis, Planktolyngbya and Aphanocapsa. The red circle indicates the 2-MIB event that coincided with high abundance of Aphanocapsa. 57 geosmin models. Figure 3.26 shows time series plots of geosmin and these three aforementioned parameters that revealed an association with this compound. Other significant associations of variables with 2-MIB levels were not consistent between the isolated events analysed.
Figure 3.20 Loading column plot for the first principal component in the PLS model developed for geosmin based on the entire data set for Chichester Reservoir.
Figure 3.21 Loading column plot for the first principal component in the PLS model developed for the first geosmin event in Chichester Reservoir. 58
Figure 3.22 Loading column plot for the first principal component in the PLS model developed for the second geosmin event in Chichester Reservoir.
Figure 3.23 Loading column plot for the first principal component in the PLS model developed for the third geosmin event in Chichester Reservoir.
59
Figure 3.24 Loading column plot for the first principal component in the PLS model developed for the fourth geosmin event in Chichester Reservoir.
180 8,000 Geosmin 160 7,000 2-MIB 140 Anabaena 6,000 120 5,000 100 4,000 80
3,000 Cells/mL 60 2,000 Concentration (ng/L) Concentration 40
20 1,000
0 0
06/2011 09/2002 02/2003 07/2003 12/2003 05/2004 10/2004 03/2005 08/2005 01/2006 06/2006 11/2006 04/2007 09/2007 02/2008 07/2008 12/2008 05/2009 10/2009 03/2010 08/2010 01/2011 11/2011 04/2012
Date
Figure 3.25 Chichester Reservoir geosmin and 2-MIB concentration and Anabaena abundance. Arrows indicate geosmin peaks that did not coincide with high abundance of Anabaena.
60
Geosmin 180 25 180 Geosmin 60 2-MIB 160 160 2-MIB Si 50 140 20 140 Turb 120 120 40 15 100 100 30 80 80 10
60 60 20 Si (mg/L) Si
40 40 Turbidity (NTU)
5 10 Concentration(ng/L)
20 Concentration(ng/L) 20
0 0 0 0
05/2004 09/2002 04/2003 10/2003 12/2004 06/2005 01/2006 07/2006 02/2007 08/2007 03/2008 10/2008 04/2009 11/2009 05/2010 12/2010 07/2011 01/2012
01/2012 09/2002 04/2003 10/2003 05/2004 12/2004 06/2005 01/2006 07/2006 02/2007 08/2007 03/2008 10/2008 04/2009 11/2009 05/2010 12/2010 07/2011 Date Date
Geosmin 180 2-MIB 158 160 Level 157 140 156 120 155 100 154 80 153 60
40 152 Level (m AHD) Concentration(ng/L) 20 151
0 150
11/2009 09/2002 04/2003 10/2003 05/2004 12/2004 06/2005 01/2006 07/2006 02/2007 08/2007 03/2008 10/2008 04/2009 05/2010 12/2010 07/2011 01/2012 Date Figure 3.26 Time series plots of geosmin and 2-MIB in Chichester Reservoir with silica concentration (top left), turbidity (top right) and water level (bottom).
61
3.4 Discussion
PLS regression models were developed in this study using historical water quality data for the purpose of attempting to identify routinely monitored parameters that may be related to the occurrence of geosmin and 2-MIB episodes in drinking water reservoirs. Through interpretation of the loading plots, many variables showed little or an inconsistent relationship with the T&O causing compounds when comparing the models developed for the isolated events and the entire data set. Those few variables that did consistently show an association with these compounds were considered potentially important in the development of T&O events and are further discussed.
Many studies have suggested that trophic status of reservoirs, particularly nutrients and chlorophyll ɑ levels are a good indicator of T&O causing compounds, with eutrophic or hypereutrophic reservoirs considered most vulnerable to T&O events (Rosen et al., 1992;
Downing et al., 2001; Parinet et al., 2013). Geosmin concentration levels in three Swiss lakes were found to align with nutrient status, increasing from oligotrophic to eutrophic status
(Peter et al., 2009). Grahamstown and Chichester Reservoirs are generally in the mesotrophic range and overall, when considering the PLS models developed using the entire data set and those for isolated events, both compounds had little association with chlorophyll ɑ or nutrient parameters. Dzialowski et al. (2009) developed a series of predictive models to relate reservoir geosmin concentrations to water quality variables and found that reservoir trophic state alone could not be used to predict T&O episodes and even observed that a reservoir with the lowest nutrient and chlorophyll ɑ level had the highest geosmin concentration. A similar trend was also observed by Watson et al. (2007) where geosmin peaks in western
Lake Ontario (Canada) increased significantly over an 8 year period without any change in the ambient nutrient or chlorophyll ɑ level. Although nutrient parameters showed little influence on the models, total phosphorous was found to have a moderate negative loading 62 on all but one of the 2-MIB models. Phosphorous has been observed in several studies to be inversely associated with geosmin or 2-MIB production, possibly due to its depletion by the growth of aquatic organisms such as cyanobacteria in phytoplankton and benthic environments, and the subsequent production of the secondary metabolites by some of these organisms (Park et al., 2001; Dzialowski et al., 2009).
In addition to eutrophic conditions, T&O events have been largely associated with warm temperatures, occurring predominately in the summer months (Tung et al., 2006). Peter et al.
(2009) observed that concentration peaks of geosmin and 2-MIB occurred in the summer concurrent with high phytoplankton biomass. Figures 3.27 and 3.28 show the seasonal mean presence of both compounds in Grahamstown and Chichester Reservoirs respectively. It is evident that, particularly for Grahamstown, cases of elevated geosmin and 2-MIB concentration have occurred largely in summer, but have also occurred in spring, autumn and winter. In Chichester, both compounds have occurred in all seasons to a comparatively similar extent. While some PLS models revealed water temperature was positively related to
T&O compounds (e.g. 2-MIB in Grahamstown), others showed a negative association or no significant relationship. This contradicts the widespread view that warmer water temperatures, which favour cyanobacterial growth, are more prone to T&O events. Similarly,
Dzialowski et al. (2009) observed that T&O events in drinking water reservoirs in Kansas
(USA) were not confined to summer months, with elevated geosmin concentrations observed in several reservoirs during winter. Similar findings have been observed in other reports
(Sugiura et al., 2004; Parinet et al., 2010). The fact that the reservoirs are prone to T&O problems all year round may be attributed to the borderline oceanic/humid subtropical climate of most of central and northern NSW, with warm, often humid summers and mild winters which permit the growth of phytoplankton all year round. Cyanobacteria, some of
63 which are T&O producers, generally dominate phytoplankton communities in both reservoirs based on cell counts (Cole & Williams, 2011).
25 Geosmin 2-MIB 20 15 10 5
Concentration Concentration (ng/L) 0 Spring Summer Autumn Winter Season Figure 3.27 Seasonal presence of geosmin and 2-MIB in Grahamstown Reservoir from 1998 till 2012. Mean values are presented with the associated standard error bars.
12 Geosmin 2-MIB 10 8 6 4 2 Concentration Concentration (ng/L) 0 Spring Summer Autumn Winter Season Figure 3.28 Seasonal presence of geosmin and 2-MIB in Chichester Reservoir from 2002 till 2012. Mean values are presented with the associated standard error bars.
The model results indicated quite convincingly that the periodically abundant cyanobacterial genus Anabaena was likely responsible for the occurrence of geosmin in both reservoirs.
Anabaena numbers were high during the rise and peak of most geosmin episodes and correspondingly decreased with the decline of the events as indicated in the time series plots
(Figures 3.12 and 3.25). Furthermore, the PLS models developed for geosmin based on the
64 entire data sets in addition to the episodes of particularly high geosmin concentration, exceeding 100 ng/L in Chichester (event 4) and 200 ng/L in Grahamstown (events 4 and 6), revealed the strongest loadings by Anabaena in the models. More than 40 species of cyanobacteria from many genera have been associated with geosmin and 2-MIB which are summarised by Juttner and Watson (2007) and Krishnana et al. (2008). Anabaena, a well- known bloom-forming cyanobacterial genus has been reported to be responsible for 46% of geosmin related T&O events (Su et al., 2013). In Australia, geosmin and 2-MIB occurrence have been well correlated with planktonic Anabaena and Phormidium (Jones & Korth, 1995;
Izaguirre et al., 1999; Uwins et al., 2007; Hobson et al., 2010). Cees et al. (1974) observed correlations between the presence of geosmin in water and the occurrence of Anabaena in one instance, and Aphanizomenon, Microcystis, and Oscillatoria in another. Park et al.
(2001) found a high correlation between geosmin and Anabaena spiroides density in the
Daechung Reservoir, Korea, with a density above 10,000 cells/mL indicating the presence of geosmin. In the Hinze Dam (Gold Coast, Australia) geosmin correlated significantly with
Anabaena numbers as well as water temperature. The authors attributed the rise in cell numbers to rainfall events followed by a pulse of nutrients into the dam at the onset of summer where the water temperature exceeded 22°C, thus promoting growth of Anabaena cells (Uwins et al., 2007). In the Xionghe Reservoir (China), geosmin was found to be highly correlated with planktonic Anabaena circinalis and 2-MIB with Pseudoanabaena sp. (Zhang et al., 2010).
Dissolved oxygen concentration was found to consistently negatively associate with geosmin in most Grahamstown Reservoir PLS models, in addition to 2-MIB, which may be related to depletion of oxygen during die-off of cyanobacterial blooms, during which, odour metabolites are generally released from within the cells by their breakdown (Juttner &
Watson, 2007). This inverse relationship was not observed however, for geosmin events in 65
Chichester Reservoir. The negative relationship between geosmin and turbidity observed for
Chichester Reservoir may be attributed to the fact that low turbidity levels would allow sunlight to readily penetrate the water column, therefore stimulating the growth of photosynthetic geosmin producing Anabaena. In this reservoir, silica revealed consistent negative loadings on geosmin PLS models, probably as this micronutrient is an important factor controlling phytoplankton succession. Diatoms are dependent on silica for growth and when concentrations become exhausted, the growth of cyanobacteria is favoured, thus leading to odour production (Tsujimura & Okubo, 2003). The inverse relationship was also apparent between 2-MIB and silica in Grahamstown Reservoir for the models developed for the three isolated events. However, unlike the strong association between geosmin and
Anabaena, no cyanobacterial or indeed any other biological parameter could be consistently related to 2-MIB. The lack of correlation between geosmin and 2-MIB, and the absence of their concurrence supports the notion that these T&O secondary metabolites are derived from separate sources.
Geosmin has been the main nuisance compound responsible for T&O in both reservoirs, although several episodes of elevated 2-MIB concentration have occurred in Grahamstown
Reservoir, up to a concentration of 258 ng/L. Therefore elucidation of the origins of this compound is important to anticipate future outbreaks. While the PLS models for 2-MIB events 1 and 2 found that the cyanobacterial genera Microcystis and Planktolyngbya respectively positively associated with this musty secondary metabolite, examination of time series plots (Figure 3.19) revealed the fairly low abundance of each genus that coincided with elevated 2-MIB concentrations. Examination of all cyanobacterial parameters monitored by
HWC revealed a peak in high abundance of the more numerous genus Aphanocapsa which occurred at the onset of the second 2-MIB event (Figure 3.19). However, previous work on strains of this genus isolated from Grahamstown Reservoir demonstrated that they were 66 incapable of producing 2-MIB or geosmin (Cole & Williams, 2011). Given the fact that 2-
MIB events occurred in the absence of high abundance of this genus, and high abundance occurred without producing 2-MIB episodes suggests these bacteria are not the source of 2-
MIB.
There are a multiplicity of microorganisms that can produce geosmin and 2-MIB and a multiplicity of habitats that can foster development of these compounds which have been implicated in T&O episodes (Juttner & Watson, 2007; Parinet et al., 2010). While it may seem that cyanobacteria are not responsible for imparting 2-MIB into the reservoirs, it is important to recognise that this analysis was conducted using surface water quality data.
Many of the known cyanobacterial producers are non-planktonic and the potential importance of these cyanobacteria, such as those in benthic and littoral zones can be easily overlooked (Butakova, 2013). Grahamstown Reservoir was included in a series of studies by
Hobson et al. (2010) on the sources of T&O not associated with phytoplankton, focusing on benthic, epiphytic and sediment associated sources. The work identified epiphytic cyanobacteria attached to the abundant introduced plant Torpedo Grass (Panicum repens) fringing the margins of the reservoir including Phormidium, Pseudanabaena and Anabaena spp. Additionally, benthic Phormidium spp. were isolated from one sampling site. Such non- planktonic cyanobacterial populations have been suggested to be a possible source of 2-MIB in the reservoir (Cole & Williams, 2011). Benthic cyanobacteria such as Phormidium have been linked to T&O problems in Australia (Burch & Baker, 2000) and other locations
(Jahnichen et al., 2011; Sun et al., 2013). Although the severe episodes of geosmin were all tied to the dynamics of Anabaena in both reservoirs, there were some instances of elevated geosmin concentrations which did not align with high abundance of this cyanobacterial genus. This implies that other geosmin producing organisms may be responsible for imparting relatively lower concentrations of this compound in the reservoirs. 67
Geosmin and 2-MIB are also thought to be sourced from terrestrial microbial activity, such as organisms growing in substrates in the shallows margins of freshwater systems or substrates adjacent to water bodies. As indicated in Chapter 2, one prime suspect group that inhabit such environments and may contribute to T&O is Streptomyces. The study by Hobson et al. (2010) also identified Streptomyces in the sediment of marginal sampling locations, but given that isolates were only found capable of producing geosmin, it was suggested they were not the unidentified source of 2-MIB, but may contribute to geosmin production. The research presented in the proceeding chapters focuses on the potential role of these bacteria in contributing to geosmin and 2-MIB caused T&O episodes in drinking water reservoirs.
While these bacteria have been recognised as contributors to T&O in drinking water supplies for many years, as emphasised in Chapter 2, knowledge of their distribution, abundance and activity in freshwater systems and thus their involvement in T&O is limited and requires further research (Zaitlin & Watson, 2006). It has previously been argued that Streptomyces do not actively grow in the water mass, being an artefact of terrestrial wash-in and spore survival, and thus do not directly contribute to T&O (Cross, 1981).
Water level was a parameter that was consistently inversely related to 2-MIB and the time series plot in Figure 3.18 shows that 2-MIB peaks have coincided with lowered water level.
This was also apparent for geosmin events in Chichester Reservoir (Figure 3.26). It has been suggested by several authors (Wood et al., 1983a; Bailey, 1988; Wnorowski, 1992) that exposure of sediments and debris at the margins of freshwater systems due to water level recession stimulates the growth and odour production of aerobic Streptomyces inhabiting such substrates. Subsequent wash-in of these substrates entrained with Streptomyces and their odorous secondary metabolites can then contribute to T&O. Indeed, during a drought period in the late 1980s in Grahamstown Reservoir, water levels were lowered, exposing shallow margins. This was followed by refilling and submerging of the shoreline, coinciding with 68 complaints of an earthy-musty T&O in drinking water (Cole & Williams, 2011). Thus periods of decreasing water level may be indicative of periods of increased risk of T&O problems developing in water supplies. This possible means by which Streptomyces at the margins of reservoirs have been proposed to contribute to T&O episodes in reservoirs is systematically explored in the field and laboratory studies presented in Chapters 4 and 5.
It is important to acknowledge that because geosmin and 2-MIB break down relatively slowly in the water column, the dynamics of these compounds may become disconnected from instantaneous measurements of various water quality parameters (Dzialowski et al., 2009).
Although in this study parameters that revealed consistent relationships with geosmin and 2-
MIB in the models developed were considered potentially important in T&O episodes, it should be noted that isolated events may indeed be caused by different mechanisms. In a review based on combined research, Watson et al. (2007) emphasised that different mechanisms are responsible for T&O in Lake Ontario and the Saint Lawrence River in North
America. In the North West of the lake, brief late summer geosmin peaks have been traced to planktonic cyanobacteria in the illuminated offshore surface layers, whereas in the East
Basin and Saint Lawrence River, both 2-MIB and geosmin are produced annually over a long period (Autumn), largely being derived from the substrate and macrophyte biofilms in shoreline areas (both cyanobacteria and actinobacteria). The multiplicity of organisms that produce geosmin and 2-MIB and the multiplicity of habitats that foster their production means that source tracking T&O events is a complex task (Juttner & Watson, 2007).
3.5 Conclusions
The analysis of large and complex historical water quality data sets in this study using the
PLS regression methodology has enabled identification of some routinely measured variables
69 which are associated with and may be responsible for triggering the occurrence of geosmin and 2-MIB in Grahamstown and Chichester drinking water reservoirs. Geosmin in both reservoirs appears to originate from bursts of the periodically abundant cyanobacterial genus
Anabaena. However, observation of some cases of elevated geosmin concentration coinciding with low abundance of this genus indicates that other geosmin producing organisms may also be contributing to the occurrence of this compound. The occurrence of
2-MIB in Grahamstown Reservoir could not be attributed to any biological parameter.
Although non-planktonic cyanobacteria may be responsible for imparting this musty compound to the water mass, it has long been suspected that Streptomyces may play a role in contributing to its presence in Grahamstown and Chichester drinking water supplies. As a consequence, the potential role of Streptomyces in contributing to this aesthetic water quality issue forms the focus of research presented in proceeding chapters of this thesis.
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CHAPTER 4 - STREPTOMYCES IN DRINKING WATER RESERVOIRS: DISTRIBUTION, ACTIVITY AND POTENTIAL CONTRIBUTION TO TASTE AND ODOUR EPISODES INVOLVING GEOSMIN AND 2-METHYLISOBORNEOL
4.1 Introduction
Geosmin and 2-MIB are biosynthesised by a range of organisms, from prokaryotes such as cyanobacteria and actinobacteria to eukaryotes such as fungi (Juttner & Watson, 2007; Zaitlin
& Watson, 2006). Cyanobacteria are considered to be major sources of these secondary metabolites in aquatic environments, including planktonic and benthic populations (Juttner
& Watson, 2007). In the historical water quality data analysis of Grahamstown and
Chichester drinking water reservoirs presented in Chapter 3, a strong correlation was identified between geosmin concentration and Anabaena spp. abundance in the surface waters. However, not all individual geosmin events coincided with elevated cell densities of
Anabaena spp., suggesting that these organisms were almost certainly not the only source of this compound in the reservoirs. Furthermore, the biological source of 2-MIB remained unclear. Consequently actinobacteria, and in particular the genus Streptomyces, has been considered as a possible cause of these unexplained T&O events. As emphasised by Wood,
Williams and White (1983a), the successful prediction and control of geosmin and 2-MIB episodes originating from microbes requires identification of the producers, locations of their growth sites in reservoirs and environmental factors influencing their biosynthesis of these secondary metabolites. This study endeavoured to focus on the first two of these three recommendations, with the objective of assessing Streptomyces, the most representative genus more frequently implicated in T&O events, as a potential causative microorganism and investigating their abundance and activity in a variety of habitats in drinking water reservoirs.
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The third recommendation to better understand T&O episodes involving Streptomyces is the focus of experimental work presented in Chapters 5 and 6.
In a review of existing literature, Zaitlin and Watson (2006) indicated that the evidence linking Streptomyces to T&O episodes is tenuous, largely because their total abundances and distribution across drinking water reservoirs and the degree to which they represent a metabolically active component of aquatic microbial communities has been difficult to establish. Numerous investigations have attempted to enumerate Streptomyces using culture- dependent methods and establish correlations with concentrations of geosmin and 2-MIB
(Morris, 1962; Weete et al., 1977; Jensen et al., 1994; Persson, 1995; Sugiura & Nakano,
2000; Uwins, 2011). The general lack of correlation identified in the literature may indicate that Streptomyces are not the primary causative microorganisms; however, it does not exclude these bacteria from contributing to the presence of geosmin and 2-MIB in drinking water reservoirs. Many researchers have considered that the causative organisms for earthy- musty odours should be found within the water column and have not investigated beyond water samples (Aoyama, 1990; Jensen et al., 1994). With this approach, researchers could overlook Streptomyces found in environments such as sediments, soils and plant material at the margins of reservoirs or bottom sediments which could represent potential sources of geosmin and 2-MIB.
Streptomyces, being saprophytic bacteria, are not only known to be abundant and widely distributed in soils (Goodfellow & Williams, 1983; Williams et al., 1984), it has been suggested that they are strictly terrestrial bacteria, with Cross (1981) for example, describing them as “terrigenous species which occasionally find themselves in water” (p. 407). In fact, the existence of ‘aquatic’ Streptomyces has been largely disputed and detections of these
72 bacteria in water are often considered an artefact of wash-in of surrounding terrestrial substrates and spore survival (Johnston & Cross, 1976a; Cross, 1981). Evidence for their growth in aquatic environments is undoubtedly lacking. It has been suggested that their primary mechanism of contributing to T&O events would be through their growth and odour production in the marginal soils, sediments and plant material adjacent to reservoirs. As
Streptomyces are obligate aerobes, their growth and production of earthy-musty secondary metabolites requires exposure to adequate levels of oxygen. Substrates such as soil, sediment and plant debris at the margins of reservoirs periodically become exposed to air, when water levels recede during dry conditions. Exposure as opposed to submersion of marginal substrates would stimulate the growth and activity of Streptomyces and thus their production of odorous metabolites (Wood et al., 1983a). Marginal substrates could then represent a significant source of Streptomyces and it is therefore highly plausible to consider that following rainfall events, Streptomyces, entrained with their odorous metabolites, would be washed into reservoirs. This potential means of contributing to T&O episodes has been supported by several studies (Raschke et al., 1975; Jensen et al., 1994; Lanciotti et al., 2003;
Zaitlin et al., 2003a; Zaitlin et al., 2003b).
These propositions regarding the distribution and population dynamics of Streptomyces in drinking water reservoirs demand systematic investigation in order to better understand their potential to contribute to T&O episodes. In addressing this, several specific research questions were identified on the basis of literature assertions that Streptomyces are strictly aerobic, predominately terrestrial and that their wash-in, accompanied by their odorous metabolites from marginal environments, represents their primary mechanism of contributing to the occurrence of T&O compounds in surface waters (Lanciotti et al., 2003; Zaitlin et al.,
2003a). These were:
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Are Streptomyces widely distributed throughout the water body of drinking water
reservoirs and are they potentially active in this environment?
How does Streptomyces abundance and potential activity in the water body compare
with that in physical substrates1 at the margins of reservoirs?
Which marginal substrates harbour the greatest abundance and potentially active
populations of Streptomyces?
Does Streptomyces abundance and activity in the reservoir water body, and in/on
marginal substrates, vary significantly between prolonged periods of rainfall with
relatively high reservoir volumes, and extended dry conditions with decreased
reservoir volumes?
In order to address these questions, an appropriate spatial and temporal sampling program was implemented at Grahamstown and Chichester drinking water reservoirs, including the collection of a variety of samples (water, soil, sediment and plant material) from different locations (marginal and offshore). Two sampling events were scheduled at each reservoir, one during wet weather conditions when both reservoirs were at maximum storage capacity and again during an extended dry period, which allowed the water levels to recede considerably, exposing the marginal sampling locations. Both reservoirs were considered ideal for carrying out this research as they are spatially diverse aquatic systems with a number of potentially suitable habitats to harbour Streptomyces populations and both have suffered
T&O outbreaks involving geosmin and 2-MIB, with the source of 2-MIB in particular remaining largely unidentified. Additionally, these reservoirs were considered ideal for comparing wet and dry conditions as they are vulnerable to drought, filling quickly but
1 In this context the term substrate refers to physical matrices (such as plant debris, soil and sediment) as distinct from the more conventional chemical use (reference to carbon source) found among the microbiological literature. 74 emptying relatively fast as well. Water levels in these reservoirs drop faster than those in the water storages of most other major Australian urban centres during prolonged dry conditions because of their relatively shallow water profile and high evaporation rates (Cole & Williams,
2011).
The sampling program and experimental protocols were designed around the following hypotheses:
1. During wet conditions involving run-off from rain events, densities of Streptomyces
in water samples would exceed those detected during prolonged dry conditions due
to their ‘wash-in’ from marginal environments.
2. As Streptomyces are predominately terrestrial bacteria, their abundance and activity
in soil and sediment collected at the margins of reservoirs would greatly exceed those
detected in bottom sediment samples collected in offshore locations.
3. Considering that Streptomyces are aerobic bacteria, their abundance and activity in
substrates located at the margins of reservoirs would be greater during dry conditions
when these environments become exposed to air following water level recessions
compared to wet conditions when these substrates are submerged.
The detection of Streptomyces in the environment has to date primarily been determined using culture-based techniques such as growth on selective agar media. This approach is problematic as it excludes those Streptomyces spp. which are not readily cultivated on traditional media and provides no information on potential activity. That is, discrimination between colonies that originate from either dormant spores or metabolically active mycelial cells cannot be achieved. Thus, there has been a lack of reliable evidence on Streptomyces abundance in aquatic environments in addition to the degree to which they exist as
75 metabolically active components of aquatic microbial communities. A number of quantitative molecular methods have of recent years been applied for estimating the abundance and activity of Streptomyces in terrestrial and aquatic environments. Several studies have employed the fluorescence in situ hybridisation (FISH) technique targeting rRNA molecules to quantify densities and determine activity levels of filamentous actinobacteria (assumed to represent predominately Streptomyces) in the waters of aquaculture ponds and reservoirs (Klausen et al., 2005; Nielsen et al., 2006). However, these studies were limited by the non-specificity of the probes for Streptomyces, which is not the only filamentous genus of this class.
The polymerase chain reaction (PCR) is a well-established, reliable, robust, sensitive and fast molecular approach for microbial detection. It exploits the semi-conservative replication of
DNA to enable exponential amplification of a target sequence. By using fluorescently labelled probes or dyes, the progression of DNA amplification during PCR can be measured, which indicates the concentration of the target sequence in a sample. This technique, known as quantitative PCR (qPCR), coupled with nucleic acid isolation, is a powerful tool for quantifying bacterial abundance in complex environmental samples (Smith & Osborn, 2008;
Hirsch et al., 2010). Recently, Lylloff et al. (2012) used a TaqMan qPCR protocol targeting the 23S rRNA gene to measure Streptomyces abundance in a number of drinking water reservoirs. The abundance of Streptomyces in soil has also been monitored using a qPCR protocol targeting the 16S rRNA gene (Schlatter et al., 2010). This study reports on the use of a qPCR protocol with genus-specific primers targeting the 16S rRNA gene to investigate the presence and abundance of Streptomyces in various habitats within drinking water reservoirs. The use of primers targeting 16S rRNA genes can allow for low detection limits due to the high copy number of ribosomal sequences and detection of specific taxa in
76 complex microbial communities due to the highly variable regions of the gene (Hirsch et al.,
2010).
The molecular detection and quantification of Streptomyces in environmental samples using qPCR requires the isolation and preparation of nucleic acid. The first critical step in the process of extracting DNA from a sample involves microbial cell disruption to release its genomic material. Bacterial cells are often disrupted by mechanical, enzymatic or chemical means or a combination of such treatments. Metabolically dormant spores of Streptomyces are much more resistant than the corresponding growing (vegetative) cells to relatively gentle enzymatic, chemical and heat treatments commonly employed for lysing cells (Elliot &
Flardh, 2012). Many factors contribute to the resistance of spores to adverse environmental conditions including thicker cell walls (>50 nm) compared to hyphal cell walls (10-12 nm), higher proportion of the disaccharide trehalose (10% dw) contributing to macromolecule and membrane protection, condensed and protected nucleoids, low metabolic activity and some degree of cytoplasmic dehydration, making them primed for survival and dispersal in the environment (Elliot & Flardh, 2012). More aggressive approaches such as mechanical lysis are required for disrupting the thicker cell walls of spores for the extraction of nucleic acids.
This study reports on the use of two procedures of cell lysis adapted from previous methods for the extraction of environmental DNA which exploits the differential resistance of
Streptomyces vegetative cells and spores to rupturing (Daza et al., 1989; Cresswell et al.,
1991; Filippova et al., 2005; Mincer et al., 2005). This was proposed to enable the differentiation between Streptomyces occurring as vegetative cells and as spores in environmental DNA subjected to qPCR. Considering there is little experimental evidence about the activity of Streptomyces in aquatic environments, distinguishing between growing and dormant forms in total population estimates is important. Without knowledge of their
77 ability to be metabolically active and therefore capacity for in situ geosmin and 2-MIB production, their contribution to T&O remains yet to be fully established.
This is the first report to employ qPCR using genus-specific primers targeting 16S rRNA to measure Streptomyces abundance in environmental samples collected from drinking water reservoirs and to distinguish between vegetative cells and spores to provide evidence for their potential ability to be metabolically active in aquatic environments. These molecular approaches were considered appropriate for testing the stated hypotheses. This study also focused on culture-dependent enumeration of Streptomyces for comparison with qPCR determined abundance in addition to identifying separate Streptomyces species found in the collected samples based on 16S rRNA sequences and to examine their geosmin and 2-MIB producing ability. Although Streptomyces are readily detected in environmental samples, this does not necessarily imply that the source substrate can support their growth and production of earthy-musty secondary metabolites. Experimental work presented in this chapter represents an attempt to investigate the ability of environmental substrates collected from drinking water reservoirs to support the growth and production of geosmin and 2-MIB by
Streptomyces spp. It was considered that such information may indicate their ability to contribute to in situ production of T&O causing secondary metabolites and identify significant sources, especially in relation to Grahamstown and Chichester Reservoirs. Two additional research questions related to the aforementioned components of the study include:
What proportion of Streptomyces species found in reservoir habitats exhibit capacity
for production of geosmin and 2-MIB?
Are conditions in reservoir waters and habitat substrates likely to support production
of geosmin and 2-MIB by Streptomyces spp.?
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4.2 Materials and methods
4.2.1 Sampling sites and collection
Grahamstown and Chichester drinking water reservoirs, operated by HWC were selected for the examination of abundance, activity and T&O producing potential of Streptomyces. Both freshwater storage facilities supply drinking water to the Lower Hunter Region.
Characteristics of the reservoirs have been described in depth in Chapter 3 (section 3.2.1). In brief, Grahamstown Reservoir, the primary drinking water supply for the region, is shallower
(average depth of 9 m) with a larger surface area (28 km2) and storage capacity (182,305
ML). In contrast, Chichester Reservoir is deeper (maximum depth of 37 m), with a smaller surface area (1.8 km2) and storage capacity (18,356 ML). The majority of Chichester
Reservoir’s 199 km2 catchment area is comprised of forest, whereas Grahamstown
Reservoir’s catchment (115 km2) is largely impacted by rural or urban land use (Cole &
Williams, 2011).
In order to address the hypotheses concerning the differences in Streptomyces abundance and activity between wet and dry conditions, each reservoir was sampled under both settings. The first sampling event occurred when the reservoirs were at 100% storage capacity after a prolonged period of rain (February 12th and April 23rd 2013 for Grahamstown and Chichester
Reservoir respectively). The second sampling events took place after an extensive dry period, when Grahamstown Reservoir had dropped to 88% capacity (9th November 2013) and
Chichester Reservoir to 83% capacity (1st October 2013). In both reservoirs, the lowering of the water level during the extended dry conditions exposed the soil and sediments at the margins of the reservoirs.
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In this study, two types of sampling site were of interest, these being marginal and open water locations as shown in Figure 4.1. Marginal sampling sites were those at the edge of the reservoirs where the effects of water level fluctuations could be investigated. The open water sampling locations were offshore sites, generally located in the mid sections of the reservoirs.
The maps presented in Figures 4.2 and 4.3 show the locations of the selected marginal and open water sampling sites for Grahamstown and Chichester drinking water reservoirs respectively. Marginal sampling locations were selected on the basis of potential sources for substances that may serve as substrates for growth and odour production by Streptomyces.
The sampling protocol was designed to examine a wide variety of substrates which have been identified in the literature as possible growth sites of Streptomyces and where there is potential for T&O production to originate. Table 4.1 and Figure 4.1 provide details of the samples collected.
Figure 4.1 Schematic of sampling protocol for the marginal and open water sampling sites. Samples were collected from each reservoir during both wet weather conditions (A) and prolonged dry conditions (B).
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Figure 4.2 Grahamstown Reservoir indicating the three marginal and three open water sampling locations (source: Google Earth).
Figure 4.3 Chichester Reservoir showing the two marginal and two open water sampling locations (source: Google Earth). 81
Table 4.1 Sampling locations and samples collected from Grahamstown and Chichester drinking water reservoirs. Reservoir Sampling site Samples collected R2 Marginal Soil, sediment, Nuphar sp. (Lily), water
R6 Marginal Soil, sediment, water, Panicum repens (Torpedo Grass).
Western (W) Soil, sediment, water, Eleocharis sphacelata (Tall Marginal Spikerush).
R2 Open Water Surface water, bottom water, bottom sediment (7-8 m).
Grahamstown R6 Open Water Surface water, bottom water, bottom sediment (10 m).
R12 Open Water Surface water, bottom water, bottom sediment (10 m).
South Marginal Soil, sediment, plant debris, water, E. sphacelata (Tall Spikerush). North Marginal Soil, sediment, plant debris, water, E. sphacelata (Tall Spikerush).
South Open Surface water, bottom water, bottom sediment (6-7m). Water
Chichester North Open Surface water, bottom water, bottom sediment (30-32 m). Water
At marginal locations (examples shown in Figures 4.4 and 4.5), surficial (top 5 cm) soil and sediment samples were collected and placed in 500 mL screw-capped sterile cylindrical polycarbonate sample jars (Bacto Laboratories Pty Ltd.). Where present, plant debris and macrophytes were also collected. Marginal water samples were collected within a 1 m distance from the exposed bank. At the offshore sampling sites, surface water samples were collected from a depth of 0.5 m and bottom water samples were obtained with a Van Dorn sampling device. Water samples were collected in duplicate and stored in 600 mL bacteriological bottles (Bacto Laboratories Pty Ltd.). A grab sampler was used to collect bottom sediments at each of the open water locations.
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Figure 4.4 A comparison of the south marginal sampling site in Chichester Reservoir during prolonged wet (left) and extended dry (right) conditions. The exposed marginal sediments and macrophytes (E. sphacelata) are evident during dry conditions.
Figure 4.5 A comparison of the western marginal sampling site in Grahamstown Reservoir during prolonged wet (left) and extended dry (right) conditions.
Samples were transported back to the laboratory within 4 hours of collection, stored at 4°C and processed within 48 hours. Subsamples were used for DNA extractions and dilution for inoculation of selective growth media. As the abundance of Streptomyces was expressed on a per dry weight basis of solid sample, subsamples were used to determine the moisture content gravimetrically by freeze drying to constant weight.
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4.2.2 Extraction of DNA from environmental samples
Prior to DNA extraction, microorganisms in the water samples were concentrated by filtration (250 mL sample volume) onto 0.45 µm (47 mm diameter) sterile polycarbonate membranes (Millipore). The filters were aseptically cut into small pieces prior to commencing DNA extraction. Plant material samples were also aseptically cut into small pieces (<1 cm2) for DNA extraction. Environmental samples such as soil, sediment, plant material and water containing particulate matter can be problematic for molecular analysis owing to the presence of organic compounds such as humic acids. Specialised kits for the extraction of DNA from soil are available for this reason. The PowerSoil® DNA isolation kit
(MO BIO Laboratories, Inc.) was selected for use as it is designed for isolating DNA of a high level of purity from environmental samples containing a high humic acid content including common soil types and more difficult soil types such as compost, sediment and manure. The effectiveness of this commercial kit in isolating high yields and pure bacterial and eukaryotic DNA from a variety of soil types and from Streptomyces specifically, has been previously demonstrated in several reports (Cotarlet et al., 2010; Schlatter et al., 2010;
Mahmoudi et al., 2011).
Cellular disruption is one of the most critical steps in DNA extraction and in this study, two lysis methods were used to obtain environmental DNA and differentiate between the occurrence of Streptomyces as vegetative cells and resilient dormant spores in a given sample. For the extraction of DNA from both spores and vegetative forms of Streptomyces, the complete PowerSoil® DNA isolation kit protocol was used in which cells were vigorously lysed through mechanical, heat and chemical methods. In duplicate, 250 mg of each solid sample or filter pieces containing the filtered water samples (250 mL) were added to bead beating tubes with a solution containing sodium dodecyl sulphate (SDS) and other disrupting agents for complete cells lysis. Following brief vortexing, the bead beating tubes were 84 incubated at 70°C for 1 hour, a modification to the manufacture’s protocol to target lysis of
Streptomyces spores, which have cell walls that are particularly difficult to disrupt. Further cell lysis was then carried out via mechanical shaking on a vortex at maximum speed for 20 minutes. Bead beating tubes were then centrifuged (10,000 x g for 1 min) and the supernatant containing crude DNA was transferred to a clean 2 mL tube. Removal of PCR inhibitors in the supernatant was then carried out in two steps using patented solutions containing reagents to precipitate non-DNA organic and inorganic material (e.g. humic substances, cell debris and proteins). A highly concentrated saline solution was mixed with the resulting supernatant to allow total genomic DNA to be captured on a silica membrane in a spin column format.
Following an ethanol based washing process to clean DNA bound to the silica filter membrane, DNA was eluted using 100 µL of sterile elution buffer.
A lysis method to extract environmental DNA from vegetative cells using conditions determined not to lyse Streptomyces spores was adapted from previous methods (Daza et al.,
1989; Cresswell et al., 1991; Filippova et al., 2005; Mincer et al., 2005). Cresswell et al.
(1991) could not detect plasmid DNA in suspensions of Streptomyces violaceolatus spores or sterilised soil inoculated with a spore suspension that were extracted using lysis conditions of SDS (1.5% w/vol) and heat (70°C 1 hour) as opposed to an intensive mechanical cell disruption method (bead-beating protocol). Both cell lysis methods liberated similar amounts of plasmid DNA from a mycelium inoculum. Similarly, Mincer, Fenical and Jensen (2005) effectively differentiated between spores and vegetative cells of the filamentous actinobacterial genus Salinospora by using a mechanical (bead-beating) and an enzymatic- heat-chemical (lysozyme 1 mg/mL, 37°C 1 hour, SDS 1% w/vol, 65°C 1 hour) lysis protocol respectively. Numerous studies have reported the complete resistance of Streptomyces spores to treatment with lysozyme in contrast to the complete loss of viability for mycelial cells
(Sohler et al., 1958; Koepsel & Ensign, 1984; Glazebrook et al., 1990; Filippova et al., 2005). 85
Additionally, the high thermoresistance of Streptomyces spores compared to vegetative cells has been widely reported in the literature (Ebner & Frea, 1970; Daza et al., 1989; Filippova et al., 2005).
The combination of conditions reported in the literature as being ineffective in lysing spores including enzymatic treatment using lysozyme (hydrolyses 1,4-beta glycosidic linkages in peptidoglycan of bacterial cells walls), chemical treatment using SDS (to disrupt membrane phospholipids), and heat treatment were applied. In duplicate, 250 mg of solid substrate samples or filter pieces containing filtered water samples (250 mL) were placed in 2 mL polyethylene tubes and suspended in TE buffer (450 µL) containing lysozyme (Sigma-
Aldrich) at a concentration of 1 mg/mL, followed by incubation at 37°C for 1 hour. SDS
(Sigma-Aldrich) was added at a concentration of 1.5% (w/v) and samples were vortexed briefly and further incubated for 1 hour at 55°C. Tubes were then centrifuged (10,000 x g, 1 min) and the supernatant containing crude DNA was transferred to clean 2 mL polyethylene tubes for purification following the PowerSoil® DNA Isolation kit purification protocol.
To ensure that the enzymatic-chemical-heat cell lysis method was ineffective at rupturing
Streptomyces spores, the following experiment was performed. Suspensions of Streptomyces spores in TE buffer were made by harvesting spores using an inoculating loop from heavily sporulating cultures (>2 weeks old) of S. coelicolor A3(2) (ATCC BAA-471) and a
Streptomyces sediment isolate STRO (tentatively identified as S. antibioticus) grown on starch-casein agar (SCA). Similarly, vegetative cell suspensions were made by harvesting mycelium from non-sporulating cultures of each Streptomyces sp. grown on SCA (3-4 days old). Equal aliquots (200 µL) of the spore and vegetative cell suspensions were then processed by both lysis methods followed by DNA purification using the commercial kit.
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The concentration and purity of DNA extracted from the spore and vegetative cell solutions by both methods was measured using a NanoDrop ND-1000 Spectrophotometer (Fisher
Scientific), using the sterile DNA elution buffer solution as a blank. DNA concentration was expressed in ng/µL and purity was assessed by the A260/A280 ratio. The absorbance maxima for nucleic acids and proteins are 260 and 280 nm respectively. A ratio of approximately 1.6-
1.8 is generally accepted as pure for DNA, with lower values indicating protein contamination (Thermo Fisher Scientific., 2013).
4.2.3 Quantitative polymerase chain reaction (qPCR) protocol
A molecular technique widely applied in microbial ecology to quantify the abundance of taxonomic and functional gene markers within the environment is qPCR. Here, DNA extracted from samples is subjected to PCR with fluorescent detection technologies including fluorescent dyes such as SYBR Green I or fluorescent probes (TaqMan) used to measure the accumulation of amplicons in ‘real time’ during each cycle of the PCR amplification. The
16S rRNA gene is one of the most exploited taxonomic gene markers in environmental microbial qPCR. This gene is highly conserved due to its importance in cell functionality and its high copy number in cells enables detection in samples containing low target cell concentrations. The qPCR protocol described below involves 16S rRNA targeting primers and the SYBR Green I fluorescent dye. This fluorescent dye preferentially intercalates into the DNA helix between adjacent base pairs. In solution, the unbound dye exhibits very little fluorescence but is greatly enhanced (1000-fold) when bound to double stranded DNA.
Therefore, during qPCR, the increase in SYBR Green I fluorescence (wavelength 530 nm) is directly proportional to the amount of double-stranded DNA (i.e. amplicon) being generated
(Smith & Osborn, 2008; Rastogi & Sani, 2011).
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For detection and quantification of Streptomyces in DNA extracted from environmental samples, genus-specific primers targeting the 16S rRNA gene were employed, generating an amplicon of 99 bp. These sequences were designed by Suutari et al. (2002) based on the alignment of Streptomyces 16S rRNA sequences and selection of sequences homologous to the genus. Taxonomically broader primers designed by Schafer et al. (2010) targeting actinobacteria-specific 16S rRNA sequences were also applied for all DNA extracted from samples generating a product approximately 274 bp. Details of this primer pair and data from their use in this study are presented in Appendix D. The primers used in this study (made and supplied by Sigma-Aldrich) for qPCR and their target DNA segments are shown in Table
4.2.
Table 4.2 Sequences of the primer pair used in the qPCR protocol for detection and quantification of Streptomyces-specific 16S rRNA. The amplified fragment size (base pairs, bp) is indicated. Primer ID Primer sequences (5’ → 3’) Amplicon Target gene Reference size 139F ACAAGCCCTGGAAACGGGGT ≈99 Streptomyces Suutari et 16S rRNA al. (2002) 237R1 GATAGGCCGCGGGCTCAT
1The reverse compliment of a 16S rRNA sequence reported by Suutari et al. (2002) for detection of Streptomyces spp. in order to generate a suitably small amplicon for qPCR. qPCR was carried out using a hot start PCR reaction master mix (FastStart Essential DNA
Green Master 2x conc, Roche Diagnostics Australia Pty Ltd) containing FastStartTaq DNA polymerase, reaction buffer, dNTPs, SYBR Green I dye and MgCl2. Each qPCR reaction was performed with 5 µL of the master mix, 0.2 µL dimethyl sulfoxide (Sigma-Aldrich), 0.1 µL of each forward and reverse primer from 25 µM working solutions (0.25 µM), 2.5 µL template DNA and 2.1 µL of PCR grade water to bring the reaction volume to 10 µL. The amplification conditions for Streptomyces-specific 16S rRNA targeting primers were as
88 follows: pre-incubation at 95°C for 600s followed by 45 cycles at 95°C for 15s
(denaturation), 64°C for 15s (annealing) and 72°C for 15s (extension).
All qPCR reactions were performed in triplicate for each DNA sample. As duplicate DNA extractions were performed for each environmental sample, this gave six determinations per sample for measuring Streptomyces abundance. Negative controls containing PCR grade water rather than template DNA were included for each qPCR experiment in addition to positive controls containing S. coelicolor A3(2) DNA. Reactions were performed in 96-well
PCR plates using the LightCycler® 96 Real-Time PCR System (Roche Diagnostics Australia
Pty Ltd). A post-PCR dissociation (melting) curve analysis was performed at the end of qPCR runs to determine the melting temperature (Tm) of the PCR product produced for the primer pair and thus to confirm that the fluorescent signal was generated from the target amplified product in samples. The melting program consisted of incubation at 95°C for 10s, followed by 65°C for 60s with continuous heating at 0.2°C /s to 97°C. Finally, the PCR products were cooled down to 37°C at 2.2°C/s. The Tm value of the single peak was identified to be 84°C
(Appendix C). Only samples displaying a single peak aligning with this temperature in the melting curve analysis were considered positive results. It is critical to avoid false positive results when using qPCR protocols to determine abundance values from different environmental matrices for comparative purposes. The SYBR Green I quantification approach enables measurement of all DNA products in qPCR. This may introduce errors if unspecific PCR products and artifacts (e.g. primer-dimers) are measured together with the specific product of interest (Smith & Osborn, 2008).
The specificity of the qPCR assay for detecting the target gene was verified by conventional
PCR using DNA isolated from S. coelicolor A3(2) (ATCC-471) and S. antibioticus (STR0).
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Amplifications were performed using the same reaction conditions specified above using a
MJ Research PTC-220 Dyad® Thermal Cycler (Bio-Rad Laboratories Inc.). The amplification products were separated by gel electrophoresis for 60 min at 80V in 2.5% agarose in TAE buffer (tris-borate and ethylenediaminetetraacetic acid) stained with ethidium bromide (0.5 mg/mL) (Sigma-Aldrich). The gel was visualised by exposure to UV light to verify the length of the amplicons. The HyperLadderTM100bp DNA ladder (Bioline
Australia) served as a base pair length indicator. Despite the specificity of the primers pair being already established in a previous report (Suutari et al., 2002), they were also validated using the NCBI Primer-BLAST tool, selecting the GenBank database covering the genomes of all organisms to identify potential target organisms and to verify the expected amplicon length (Ye et al., 2012).
DNA isolated from cultures of S. coelicolor A3(2) and S. antibioticus (STRO) were used as standards for the qPCR assay. Both species were grown on SCA for the production of spores.
After the formation of heavily sporulating colonies, the spores were gently harvested using a sterile plastic loop and suspended in 1 mL of sterile Milli-Q water. DNA was extracted from
200 µL of the spore suspensions as described previously and ten-fold serial dilutions (100 to
10-6) in PCR grade water of the purified extracts were analysed using the qPCR assay to determine cycle threshold values (CT) which were related to spore numbers. qPCR determines the amount of a target sequence present in a sample and is detected by the accumulation of a fluorescent signal (SYBR Green I). The CT value is the number of PCR cycles required for the fluorescent signal to cross the threshold level of detection (i.e. exceeds background fluorescence). CT values are inversely proportional to the concentration of the target sequence. Thus, the lower the CT value, the greater the amount of target nucleic acid in a sample and vice versa. Standard curves for the primer pair were determined by the correlation between CT values and the density of Streptomyces spp. spores. Thus raw data 90 from the LightCycler were converted into spores (cells). The amplification efficiency (E) of the primer pair was also calculated using the formula E = -1 + 10 (1/slope), in which the slope is calculated by the regression analysis of the obtained CT values versus the log number of cells in the serial dilution.
The density of the Streptomyces spores in the suspensions prepared for DNA extraction was determined by diluting and plating the suspensions on SCA followed by incubation for 7 days at 28°C and enumeration of colony forming units (CFU), with the assumption that each spore produced a single colony. Densities of spores were related to the CT values obtained through the qPCR reactions of diluted spore DNA and were used to determine calibration curves. As two standard curves were generated for the qPCR assay (two Streptomyces spp.), the measured abundance of the target sequences in environmental DNA determined by the CT value was presented as an average of values calculated from both regression equations.
Densities of cells were expressed on a per dry weight basis for solid samples (cells/g) and in cells/L for water samples. The qPCR data on Streptomyces abundance were log-transformed.
4.2.4 Enumeration and isolation of Streptomyces
Enumeration of Streptomyces-like colonies from samples was carried out to estimate the abundance of the culturable populations and compare these to qPCR-determined
Streptomyces abundance data. Additionally, this was undertaken for the purpose of obtaining isolates for identification and examination of the geosmin and 2-MIB producing ability of
Streptomyces sourced from a wide range of samples. Cultivation of Streptomyces was achieved using the dilution plate technique and two selective growth media including starch- casein agar (SCA) (United States Biological) and actinomycete isolation agar (AIA) (BD
Difco Inc). These media have been frequently reported as selective for the growth of
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Streptomyces and their composition is detailed in Table 4.3 (Kuster & Williams, 1964;
Williams & Davies, 1965; Mackay, 1977).
Table 4.3 Chemical composition of two selective growth media used for the isolation of Streptomyces-like colonies. Actinomycete isolation agar (AIA) (g/L) Starch-casein agar (SCA) (g/L) Agar 15.0 Agar 15.0 Glycerol 5.0 Soluble starch 10.0 Sodium caseinate 2.0 Casein 0.3 Asparagine 0.1 Potassium phosphate 2.0 dibasic Sodium propionate 4.0 Potassium nitrate 2.0 Potassium phosphate dibasic 0.5 Sodium chloride 2.0 Magnesium sulphate (heptahydrate) 0.1 Magnesium sulphate 0.05 (heptahydrate) Ferrous sulphate (heptahydrate) 0.001 Calcium carbonate 0.02 Ferrious sulphate 0.01 (heptahydrate)
A major problem in culturing Streptomyces from environmental samples is that they are often outcompeted by fast growing and spreading bacteria and fungi. Supplementation of isolation agar with chemicals to suppress the growth of other microorganisms is therefore necessary.
Preliminary testing using soil samples (data not shown) revealed that a combination of the antifungal compounds cycloheximide and nystatin (50 µg/mL) and antibiotics polymyxin B sulphate (5 µg/mL) and sodium penicillin (1 µg/mL) were required in the growth media to reduce contamination by fungi and other bacteria and maximise the numbers of
Streptomyces-like colonies as recommended by Williams and Davies (1965). In addition, the antibacterial compound nalidixic acid was supplemented (20 µg/mL) in both media to suppress the growth of certain bacteria (e.g. Bacillus spp.) which were found to be resistant 92 to sodium penicillin and polymyxin B sulphate (Alferova et al., 1989; Takizawa et al., 1993;
Lee et al., 2011). Table 4.4 provides an overview of each of these inhibitor compounds. Filter sterilised (0.22 µm pore size) solutions of the antifungal and antibacterial agents were added to molten media following sterilisation by autoclaving at 121°C (20 min) to give the specified concentrations. For the insoluble nystatin, gamma irradiated salts were suspended in sterile water. All antifungal and antibacterial compounds were purchased from Sigma-Aldrich.
Volumes of approximately 20 mL of the prepared media were distributed to polystyrene Petri dishes (90 x 15mm, BD Falcon).
Table 4.4 Inhibitors added to both selective growth media to increase the numbers and proportion of Streptomyces. Compound Concentration Mode of action Reference Cycloheximide 50 µg/mL Blocks protein synthesis in Williams and eukaryotic cells by blocking Davies (1965). ribosome translocation. Nystatin 50 µg/mL Inactivates membranes Williams and containing sterols. Davies (1965). Polymyxin B 5 µg/mL Damages cytoplasmic Williams and sulphate membranes of gram-negative Davies (1965). bacteria by binding to membrane phospholipids. Penicillin G 1 µg/mL Inhibits gram-positive bacteria by Williams and preventing cell wall Davies (1965). (peptidoglycan) synthesis and murein assembly. Nalidixic acid 20 µg/mL Binds to DNA gyrase enzyme Alferova et al. (topoisomerase), and therefore (1989). inhibits proper DNA replication.
The various substrates collected from both reservoirs (soil, sediment, debris and aseptically cut 1cm pieces of plant material) were serially diluted 10-fold by placing 1g (ww) in 9 mL 93 of sterile Milli-Q water followed by vigorous vortexing for 30s to release cells from particulate matter. The appropriate dilutions (determined by preliminary testing) from the resulting suspensions were used for inoculating the selective growth media in duplicate. For water samples, 1 mL was deposited onto growth media.
Due to the slow growing nature of Streptomyces, inoculated plates were inverted and incubated for three weeks at 28°C, a common temperature selected for these predominately mesophilic bacteria (Goodfellow & Williams, 1983). Numbers of Streptomyces were expressed as colony forming units (CFUs) per dry weight or volume of sample.
Streptomyces-like colonies were recognised by their macroscopic features including colonies that adhered to the agar surface and appeared as lichenoid, leathery or butyrous which may be smooth if aerial mycelium has not developed or appear floccose, granular, powdery or velvety when aerial mycelium is present and often pigmented (Figure 4.6). Selected morphologically diverse colonies displaying these Streptomyces-like features from a range of environmental samples were repeatedly transferred onto SCA (devoid of antibiotics and antifungal compounds) until pure cultures were obtained for sequencing and testing for geosmin and 2-MIB production ability. All isolates were Gram-stained and examined by light microscopy to identify them as potentially belonging to the genus Streptomyces (filamentous and spore forming).
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Secondary mycelium Primary mycelium
Figure 4.6 Colonies of predominately Streptomyces spp. growing on starch-casein agar (SCA) from a diluted (10-4) marginal sediment sample from Chichester Reservoir showing the typical morphological characteristics of this genus. The powdery/chalky secondary mycelium (white, grey or brown colours) of many isolates can be seen, in addition to the smooth and more transparent vegetative mycelium of many colonies. The insert indicates the differences between the appearances of these two growth stages.
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4.2.5 Identification of Streptomyces-like isolates
A total of 23 isolates which displayed Streptomyces-like morphological features were sourced from a range of environmental samples (Table 4.5). These were then subjected to
DNA extraction and phylogenetic identification in addition to examination of their geosmin and 2-MIB producing ability.
Table 4.5 Collection details (source and date) of the 23 Streptomyces-like isolates selected for identification and examination of geosmin and 2-MIB production ability. Isolate Collection site Collection date ID STR0 Chichester marginal sediment (south) 01/10/13 STR3 Grahamstown bottom sediment (R2) 19/11/13 STR4 Grahamstown bottom sediment (R2) 19/11/13 STR6 Grahamstown marginal water (W) 19/11/13 STR7 Chichester soil (north) 01/10/13 STR9 Grahamstown marginal sediment (R2) 19/11/13 STR11 Chichester soil (south) 01/10/13 STR12 Chichester marginal sediment (south) 01/10/13 STR13 Grahamstown surface water (R6) 19/11/13 STR14 Grahamstown bottom water (R6) 19/11/13 STR15 Chichester marginal sediment (south) 01/10/13 STR16 Chichester marginal sediment (north) 01/10/13 STR17 Grahamstown marginal water (R6) 19/11/13 STR18 Grahamstown marginal sediment (R6) 19/11/13 STR19 Grahamstown submerged sediment (R2) 19/11/13 STR20 Chichester bottom sediment (south) 01/10/13 STR22 Chichester debris (south) 01/10/13 STR25 Grahamstown bottom sediment (R2) 19/11/13 STR26 Grahamstown surface water (R2) 19/11/13 STR27 Grahamstown surface water (R2) 19/11/13 STR28 Grahamstown soil (R2) 19/11/13 STR29 Grahamstown bottom sediment (R2) 19/11/13 STR30 Grahamstown marginal water (W) 19/11/13
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4.2.5.1 Extraction of DNA from Streptomyces-like isolates
Genomic DNA from colonies of these isolates grown on SCA was extracted and purified using the UltraClean® Microbial DNA Isolation Kit (MO BIO Laboratories, Inc.) according to the manufacturer’s protocol. The method involves the lysing of cells by a combination of chemical (detergent), heat and mechanical forces (bead-beating). An additional incubation step at 65°C for 10 minutes was carried out as recommended by the manufacturer prior to mechanical cell lysis to ensure breakage of the thick cell wall of these bacteria. The released DNA was bound to a silica spin filter followed by ethanol washing and elution of the DNA.
4.2.5.2 Amplification of Streptomyces-specific 16S rRNA sequences
All isolates were tested with the primers used in the qPCR protocol to assess their identity as belonging to the genus Streptomyces, in addition to providing further testing of the specificity of the primer pair. Conventional PCR was carried out in a MJ Research PTC-
220 Dyad® Thermal Cycler (Bio-Rad Laboratories Inc.) in polycarbonate tubes using the same chemical parameters and cycling conditions described for qPCR (section 4.2.3), followed by verification of the target sized amplicons through gel electrophoresis (60 min,
80V, 2.5% agarose stained with EtBr). Amplicons of the target size were visualised under
UV light. A negative (PCR grade water) and positive control (S. coelicolor A3(2)) were included in each gel.
4.2.5.3 Sequencing of Streptomyces-like isolates
Identification of the isolates was based on 16S rRNA sequence analysis. The 16S rRNA gene for each isolate was amplified by PCR using a pair of universal (degenerate) primers (POmod and PC3mod) which recognise conserved sequences of the 5′ and 3′ ends respectively of the
97
16S rRNA genes of all eubacteria. The sequences of the primers were as follows: forward primer (POmod) 5’AGAGTTTGATCMTGG3’ and reverse primer (PC3mod)
5’GGACTAHAGGGTATCTAAT3’ (Wilson et al., 1990) (made and supplied by Sigma-
Aldrich). The primers were diluted in PCR grade water to a concentration of 25 µM and stored at -20°C. PCR reactions were carried out in a final volume of 50 µL containing 25 µL of 2 x concentrated SYBR Green master mix (Roche Diagnostics Australia Pty Ltd), 1 µL of forward and reverse primers, 5 µL of template DNA and 18 µL of PCR grade water.
Amplification was achieved using a MJ Research PTC-220 Dyad® Thermal Cycler (Bio-Rad
Laboratories Inc.) using the following cycling protocol: initial denaturation at 94.5°C for 10 min, followed by 35 cycles at 94.5°C for 30 s, 46°C for 30s and 72°C for 30s with a final extension of 8 min at 72°C. The concentration and size of the PCR fragments were verified by agarose gel electrophoresis (2.5%) with EtBr staining (0.5 µg/mL) and visualisation under
UV light using a HyperLadderTM100bp DNA ladder (Bioline Australia) as the molecular weight marker. Following verification of the correct PCR fragment size (789 bp), DNA was purified using the QIAquick® PCR Purification Kit (QIAGEN Pty Ltd) according to the protocol described by the manufacturer. For bacterial identification, purified products of the
PCR reactions were mixed with the reverse primer (PC3mod) described above and were submitted to the Australian Genome Research Facility Ltd (Brisbane, Australia) for Sanger sequencing by DNA BDT (BigDye®Terminator) labelling (sequencing) reaction. The sequencing products were analysed by capillary separation with an Applied Biosystems 3730 xl DNA analyser.
4.2.5.4 Phylogenetic analysis
Sequences of 16S rRNA which were all approximately 749-861 bp were subjected to
BLASTn analysis to identify possible close matches available in the GenBank database on
98 the NCBI (National Centre for Biotechnology Information) website. The closest sequence matches of identified type strains were obtained and sequences were then aligned using
ClustalW (Larkin et al., 2007). Phylogenetic analysis was carried out on aligned sequences using MEGA6 (Tamura et al., 2013). A phylogenetic tree was constructed using the
Neighbour-Joining method (Saitou & Nei, 1987). The percentage of replicate trees in which the associated taxa clustered together in the boostrap test (2000 replicates) were shown above the branches. The tree was drawn to scale, with branch lengths in the same units as those of the evolutionary distances used to infer the phylogenetic tree. The evolutionary distances were computed using the Maximum Composite Likelihood method (Tamura et al., 2004) and are in the units of base substitutions per site. The analysis involved 38 nucleotide sequences.
4.2.6 Geosmin and 2-MIB production by Streptomyces-like isolates
In the second phase of this work, the 23 environmental isolates were tested for geosmin and
2-MIB production ability using a simple protocol that was developed for the rapid extraction of these compounds from solid cultures, followed by their quantification using gas chromatograph-mass spectrometry (GC-MS). Spores obtained from colonies of each isolate were harvested and suspended in 1 mL of sterile Mill-Q water. An aliquot (250 µL) of the spore suspensions of each isolate was deposited in duplicate on pre-weighed 45 mm diameter and 0.45 µm pore size polycarbonate membranes (Millipore) placed on the surface of SCA plates (60 x 15mm, BD Falcon). Plates were inverted and incubated at 28°C for 7 days and were subsequently examined for geosmin and 2-MIB production. To determine the biomass of each culture, the polycarbonate membranes were removed from the medium, freeze-dried to constant weight followed by subtraction of the predetermined membrane weight. Although testing Streptomyces isolates for T&O producing capacity has most often been conducted using liquid cultures, several other studies have similarly employed the use of solid cultures
99 grown over polycarbonate membranes in order to investigate geosmin production (Dionigi et al., 1992; Dionigi & Ingram, 1994; Dionigi et al., 1996; Scholler et al., 2002).
Hexane has been reported in the literature as a suitable solvent for the extraction of both geosmin and 2-MIB (Dionigi et al., 1992; Dionigi & Ingram, 1994; Jensen et al., 1994;
Dionigi et al., 1996; Ma et al., 2007). Gas chromatography (95%) grade n-hexane (1 mL)
(Sigma-Aldrich) was deposited onto each agar plate to entirely cover the surface. Each culture was extracted for 2 minutes by gently tilting the plates back and forth to distribute the solvent. The solvent was then transferred to a 2 mL eppendorf tube and centrifuged (2,152 x g, 2 minutes) to remove cell material. The solvent (200 µL) was placed in a GC-MS vial
(Agilent Technologies) with Biphenyl-d10 (Sigma-Adrich) dissolved in hexane as an internal standard at a concentration of 20 ng/mL in addition to 50 µL of N,O-
Bis(trimethylsilyl)trifluoroacetamide (BSTFA). The latter chemical compound is used to derivatise labile groups such as hydroxyl on other chemicals, with the more stable trimethylsilyl (TMS) group, which protects the labile group and allows the compound to be used for analytical purposes. TMS derivatives are usually more volatile than the corresponding hydroxyl compounds, and thus can be analysed with gas chromatography better than the parent compound (Schummer et al., 2009). The vials were sealed with a septum screw cap. The recovery efficiency of the extraction procedure was evaluated by spiking in triplicate the surface of polycarbonate membrane covered SCA plates with known quantities of both geosmin and 2-MIB from analytical standards (5 ng, 25 ng and 100 ng) and then performing the extraction procedure as described above. Recovery percentages were determined.
100
All analyses of geosmin and 2-MIB in culture extracts were carried out on an Hewlett
Packard 5973A gas chromatograph and mass spectrometer (GC-MS) instrument (Agilent
Technologies) equipped with a HP-5ms capillary column (J & W, Scientific Inc., 30 m x 0.25 mm ID x 0.25 µm film thickness). A volume of 1 µL was injected into the inlet from each sample vial. The inlet temperature was maintained at 300°C and operated in splitless mode.
Helium was used as the carrier gas at a flow rate of 1.3 mL /min in constant flow mode. The oven temperature program was as follows: initially held at 40°C for 2 min, and then heated at 10°C/min to a final temperature of 300°C. After eluting from the GC column, the samples were carried through a MS transfer line with the temperature set at 280°C and into the source of the mass spectrometer held at 230°C with the electron ionisation at 200eV. To detect the analytes and internal standard, the mass spectrometer was operated in selected ion monitoring
(SIM) mode. In SIM mode, five ions were monitored for each analyte including m/z 164,
162, 160, 136, 80 for biphenyl-d10, m/z 112, 111, 125, 126, 182 for geosmin and m/z 95, 108,
135, 150, 168 for 2-MIB. According to the mass spectra (Figure 4.7), the base peak ion (m/z) selected for the quantification of geosmin and 2-MIB was 112 and 95 respectively and 164 for the internal standard. Quantification was determined by integrating the base peak area.
The analytes were identified and quantified based on retention time (Figure 4.8) and matching of the mass spectra against that of pure standards prepared from commercial geosmin and 2-MIB solutions (100 µg/mL MeOH) purchased from Sigma-Aldrich.
Standards were diluted in hexane over the range 0.25 ng/mL to 25 ng/mL and were analysed to generate calibration curves of geosmin and 2-MIB to allow quantification of both compounds in samples. An acceptable linearity (R2>0.98) was obtained for the calibration curves of both compounds (Figure 4.9). Biphenyl-d10 salts were dissolved in hexane and used as an internal standard (20 ng/mL) for the relative quantification of both analytes as recommended by Watson (2000). Calibration curves were constructed based on the relative response of the analytes to the internal standard (Ac/Ais) versus the concentration of the 101 analyte to the concentration of the internal standard (Cc/Cis). Detection limits for both compounds were determined to be 0.25 ng/mL.
Figure 4.7 Mass spectra of the two analytes geosmin and 2-MIB and internal standard biphenyl-d10 under SIM mode. The peak ions selected for the detection and quantification of geosmin, 2-MIB and the internal standard were (m/z) 112, 95 and 164 respectively. 102
Figure 4.8 Gas chromatogram showing the retention time (minutes) of 2-MIB (6.59), biphenyl-d10 (7.73) and geosmin (7.92) using standard solutions of each compound.
0.3 2-MIB Geosmin 0.25 y = 0.270x y = 0.331x R² = 0.994 R² = 0.983 0.2
0.15 Ac:Ais 0.1
0.05
0 0 0.2 0.4 0.6 0.8 1 Cc:Cis Figure 4.9 Calibration curves of geosmin and 2-MIB analytical standards showing mean values and standard error bars (n=3). The y-axis represents the ratio of the analyte (Ac) and internal standard (Ais) peak area whereas the x-axis is a measure of the ratio of the analyte (Cc) and internal standard (Cis) concentration.
4.2.7 Production of geosmin and 2-MIB by Streptomyces on environmental substrates
A primary objective of this study was quantifying the abundance and activity of Streptomyces in a wide variety of drinking water reservoir habitats and establishing the identity and
103 geosmin and 2-MIB production potential of a number of isolates. In addition, this study sought to assess the potential of various substrates collected including reservoir water, soil, sediment, debris and macrophytes to support the growth and geosmin and 2-MIB production by Streptomyces spp. A Chichester sediment isolate identified as S. antibioticus (STR0) was selected for use in this investigation based on its ability to produce large and relatively equivalent levels of both compounds in addition to the geosmin and 2-MIB producing ATCC strain (BAA-471) S. coelicolor A3(2). This is genetically, the best known representative of the genus, with its 8,667,507 bp linear chromosome (predicted to contain 7,825 genes) completely sequenced. Furthermore, it is a model organism for antibiotic production (Bentley et al., 2002).
Two water samples were used for testing the ability of reservoir water to support
Streptomyces spp. growth and earthy-musty odour production (R6 marginal and R6 surface water). Both water samples were sterilised by autoclaving at 121°C (20 minutes). Conical flasks (250 mL) containing 100 mL of the sterilised reservoir water were inoculated with spore suspensions (250 µL) of each Streptomyces spp. and were incubated statically at 28°C for 7 days. The solid substrates selected for examining their ability to support Streptomyces growth and earthy-musty odour production are listed in Table 4.6. These substrates were dried by lyophilisation and broken down by grinding with a mortar and pestle for a duration of 5 minutes. Dried and ground substrates were placed in water (amended with 1.5% agar) at a concentration of 50 g/L and sterilised by autoclaving at 121°C (20 minutes). A volume of
10 mL of each environmental substrate media was distributed to polystyrene Petri dishes (60 x 15mm, BD Falcon). Sterile polycarbonate membranes (0.45 µm pore size, 47 mm diameter,
Millipore) were placed over the agar surface and were inoculated with the Streptomyces spp. spore suspensions (250 µL). Following drying of the inoculum, the plates were inverted and incubated at 28°C for 7 days. Each environmental medium was tested in triplicate using both 104
Streptomyces spp. Examples of the environmental solid media plates are shown in Figure
4.10.
Table 4.6 Samples collected from Grahamstown and Chichester reservoir for testing Streptomyces spp. growth and geosmin and 2-MIB production. Sample Type Sampling Location Water Grahamstown R6 marginal and surface water Soil Chichester north and south marginal; Grahamstown western marginal Plant debris Chichester north and south marginal Bottom sediment Chichester south open water; Grahamstown R6 open water Torpedo Grass Grahamstown R6 marginal Nuphar sp. (lily) Grahamstown R2 marginal
Figure 4.10 Examples of solid media prepared using environmental substrates as the sole source of nutrition for examining their ability to support Streptomyces spp. growth and geosmin and 2-MIB production.
After the period of incubation, liquid reservoir water cultures were filtered through pre- weighed polycarbonate membranes (0.45 µm, 47 mm diameter, Millipore) and the biomass 105 was determined after drying to constant weight by freeze-drying. The culture filtrate (50 mL) was examined for geosmin and 2-MIB by a liquid-liquid extraction (LLE) protocol adapted from previous methods (Ma et al., 2007). In 250 mL conical flasks, NaCl (25% w/v) and 1 mL of n-hexane were added to the filtrate, which was vigorously mixed using a magnetic stirring device for 30 minutes (sealed with a Teflon-lined glass stopper). The addition of
NaCl was added to improve the partitioning of T&O compounds into the solvent phase. NaCl increases the ionic strength of the media which leads to a decrease in the number of water molecules surrounding geosmin and 2-MIB molecules and thus lowering their solubilities
(Xie et al., 2007). The non-aqueous solvent phase residing on the top was then removed, centrifuged (2,152 x g, 2 minutes) and supplemented with internal standard and BSTFA for analysis by GC-MS as described previously. The efficiency of the LLE protocol was determined by assessing the recovery of each compound by spiking sterile water with known quantities of geosmin and 2-MIB (500 ng/L, 100 ng/L and 50 ng/L). Geosmin and 2-MIB production by solid cultures was examined by the simple extraction procedure described previously (section 4.2.6). Biomass of solid cultures was determined by freeze-drying the membranes and subtracting the pre-determined membrane weight. Yields of geosmin and 2-
MIB produced by the Streptomyces spp. was expressed as ng/mg dw.
4.2.8 Data analysis
For the log-transformed qPCR data, t-tests were used to assess differences between wet and dry conditions in relation to the abundance of Streptomyces 16S rRNA in each environmental sample. One way analysis of variance (ANOVA) tests were performed to examine significant differences in the abundance of Streptomyces detected between environmental samples obtained during the same sampling events. ANOVA was also used to assess differences in production of geosmin and 2-MIB by Streptomyces on environmental substrate media and t- tests were used to establish significant differences between geosmin and 2-MIB production 106 yields for each environmental isolate. For all analyses the probability threshold for significant difference was set at p<0.05. Statistical analyses were carried out using JMP version 7.0.
4.3 Results
The following results comprise data pertaining to both the distribution and activity of
Streptomyces in collected samples and the efficacy of the methods employed to obtain these data. These include an evaluation of the efficiency of the cell lysis procedures for DNA extraction to differentiate between vegetative cells and spores, and the reliability of the qPCR protocol for measurement of abundance. This is followed by the qPCR derived Streptomyces abundance data for the reservoir samples collected during the wet and dry sampling events, including an estimation of their levels of activity (percent of vegetative cells), and comparison of these results with those obtained by culture-dependent enumeration of
Streptomyces in these samples. The final analyses focus on the identity and geosmin and 2-
MIB producing ability of Streptomyces-like cultivars isolated from the reservoirs, and the capacity of various reservoir-associated substrates to support growth and T&O production by Streptomyces spp.
4.3.1 Efficiency of cell lysis methods to differentiate between Streptomyces vegetative cells and spores
Streptomyces are thought to predominately exist as dormant spores in aquatic environments, incapable of active vegetative growth (Johnston & Cross, 1976a; Cross, 1981). As vegetative growth and metabolic activity is a requirement for production of geosmin and 2-MIB, the ability to discriminate between dormant and active forms of Streptomyces in samples is important to assess their ability to grow and potentially contribute to in situ production of
T&O causing compounds in drinking water reservoirs. This formed a major objective of this 107 study and was achieved by using two methods of cell lysis for DNA extraction which exploited the differential resistance of vegetative cells and spores (refer to 4.2.2 above). The purpose was to enable the relative proportions of vegetative cells and dormant spores comprising the total Streptomyces population to be determined in the genus-specific qPCR analysis of the various environmental samples.
The effectiveness of the two lysis methods in differentiating between DNA derived from these cell types was evaluated by comparing the spectrophotometrically determined yields of
DNA extracted from Streptomyces spp. spore and vegetative cell suspensions. Figure 4.11 shows a relative comparison of DNA yields between the two lysis methods for spore and vegetative cell suspensions (exact DNA yields and purity ratios [A260/A280] are presented in
Appendix B). The more aggressive lysis method of the PowerSoil® DNA isolation kit
(mechanical-chemical-heat) resulted in higher DNA yields from the spore suspensions compared to the less efficient enzymatic-chemical-heat method. The DNA concentration measured from spores lysed using the commercial procedure was approximately 29 and 4.3- fold greater for S. antibioticus and S. coelicolor A3(2) respectively compared to the enzymatic-chemical-heat protocol. The low amounts of DNA detected when subjecting spores to the latter developed lysis protocol could be attributed to the presence of contaminating hyphal fragments in the suspensions (Figure 4.12), as obtaining spores from heavily sporulating solid cultures cannot completely exclude mycelial cells. The DNA yields obtained from the vegetative cell suspensions were similar when using both lysis techniques, however the commercial technique was slightly more efficient (1.43 and 1.32 times more
DNA yield for S. antibioticus and S. coelicolor A3(2) respectively). The presence of some spores in the vegetative cell suspensions, which would resist breakdown by the enzymatic- chemical-heat treatment, may account for these slight discrepancies in DNA yield (Figure
4.12). 108
Figure 4.11 Comparison of the relative DNA yield obtained between the two cell lysis methods for DNA extractions of spores (assigned value of 1) and vegetative cell suspensions of S. antibioticus (A) and S. coelicolor A3(2) (B).
Figure 4.12 Spore (left) and vegetative cell (right) suspensions prepared from cultures of S. antibioticus (top) and S. coelicolor A3(2) (bottom) for comparing the two methods of cell lysis for DNA extraction.
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Overall, based on the comparison of spectrophotometrically determined DNA yields, it was considered that the cell lysis methods could successfully discriminate between DNA extracted exclusively from Streptomyces vegetative cells and from the total population
(vegetative cells and spores).
4.3.2 Evaluation of the qPCR assay
The choice of primers for a qPCR-based protocol can influence the results from an analysis of bacterial abundance in environmental samples. Therefore, the specificity of primers must be tested prior to their application for investigations of bacterial populations in environmental samples. The specificity of the genus-specific 16S rRNA primers designed by Suutari et al.
(2002) was confirmed using the primer-BLAST tool (only Streptomyces spp. matches based on sequences available in the GenBank database) and experimentally by PCR amplification and agarose gel electrophoresis using template DNA from Streptomyces spp. as shown in
Figure 4.13.
Figure 4.13 Gel electrophoresis of the PCR product (99 bp) obtained by amplification of DNA isolated from two Streptomyces spp. using the primer pair targeting 16S rRNA. Lane 1 contains S. antibioticus (STR0) DNA, lane 2 contains S. coelicolor A3(2) DNA and lane 3 is a negative control amplified with the Streptomyces-specific primers respectively.
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Calibration curves (Figure 4.14) were produced for the primer pair by using 10-fold serially diluted DNA extracted from spores of S. coelicolor A3(2) and S. antibioticus cultures. The
CT values obtained through qPCR were linearly correlated with the log-transformed spore densities, with significant regression coefficients obtained (R2>0.99). The qPCR assay allowed for the detection of SYBR Green I fluorescence from double stranded DNA over a linear range spanning six orders of magnitude. The cell (spore) numbers detected in the highest dilution of S. antibioticus spore DNA was equivalent to 130 cells. For S. coelicolor
A3(2), the highest dilution tested was equivalent to 970 cells. For the Streptomyces-specific primers targeting 16S rRNA, amplification efficiency was 94.21% (S. antibioticus) and
92.03% (S. coelicolor A3(2)). The results of the melting curve analysis showed a sharp melting peak at 84°C for the Streptomyces 16S rRNA targeting primers (see Appendix B).
The regression equations of the standard curves generated from Streptomyces spore DNA were used for quantifying the abundance of these bacteria in environmental samples based on the sample CT values.
35 )
T Streptomyces isolate S. coelicolor A3(2) C 30 y = -3.469x + 39.850 y = -3.529x + 40.279 R² = 1.000 R² = 0.996 25
20
15 Cycle theshold value ( value theshold Cycle
10 1 2 3 4 5 6 7 8 9 Log10 number of Streptomyces spores (cells) Figure 4.14 Standard curves for the qPCR assay using Streptomyces 16S rRNA targeting primers. The linear relationship between the threshold cycle (CT) and the number of Streptomyces spores (cells) was obtained by serially diluting spore DNA extracts and performing qPCR. The standard error based on three replicates is indicated.
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4.3.3 Detection and quantification of Streptomyces in environmental samples
Based on the assessment of the specificity of the primers for Streptomyces and their PCR amplification performance, they were considered suitable for the amplification and quantification of target 16S rRNA sequences in environmental DNA. Amplification products of the expected size were obtained for all environmental samples based on the melting curve analyses, indicating the presence of Streptomyces in all samples tested. The developed molecular techniques for quantifying Streptomyces abundance and activity in environmental samples and the implementation of an appropriate temporal and spatial sampling program enabled the hypotheses concerning possible means by which Streptomyces may contribute to
T&O episodes in drinking water reservoir to be addressed. These concerned comparisons between wet weather and dry conditions and between diverse habitats within and at the margins of the drinking water reservoirs.
4.3.3.1 Streptomyces abundance and activity in water samples
Water samples were collected from a number of locations including at the margins of both reservoirs and from offshore locations (surface and bottom water) as described in section
4.2.1. Streptomyces were detected in all water samples, with differences in abundance representing several orders of magnitude observed between samples collected during the same sampling event and between sampling events for the same source. The Streptomyces population densities in each water sample collected during both wet and dry conditions are shown in Figures 4.15 and 4.16 for Chichester and Grahamstown Reservoir respectively. The maximum abundance detected in Grahamstown Reservoir was 846,079 cells/L (R12 bottom water), compared to the lowest Streptomyces population density of 1,829 cells/L (western marginal water). In Chichester Reservoir, the largest Streptomyces population was found to be 291,160 cells/L (south marginal water), compared to the lowest 2,675 cell/L (south bottom
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Figure 4.15 Mean cell densities of Streptomyces in Chichester Reservoir water samples based on qPCR targeting genus-specific 16S rRNA sequences. Standard error bars based on duplicate samples and triplicate qPCR results are shown (n=6). Asterisks indicate significant differences in abundance for the same sample between wet and dry sampling events based on t-tests (p<0.05). Results not connected by the same number or letter (for wet and dry sampling period respectively) indicate significant differences between samples based on the ANOVA tests (p<0.05) (top graph). The bottom stacked column graph shows the proportion of vegetative cells and spores comprising the total Streptomyces population density for all samples in each sampling event.
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Figure 4.16 Mean cell densities of Streptomyces in Grahamstown Reservoir water samples based on qPCR targeting genus-specific 16S rRNA sequences. Standard error bars based on duplicate samples and triplicate qPCR results are shown (n=6). Asterisks indicate significant differences in abundance for the same sample between wet and dry sampling events based on t-tests (p<0.05). Results not connected by the same number or letter (for wet and dry sampling period respectively) indicate significant differences between samples based on the ANOVA tests (p<0.05) (top graph). The bottom stacked column graph shows the proportion of vegetative cells and spores comprising the total Streptomyces population density for all samples in each sampling event.
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water). The marginal water samples generally contained significantly greater Streptomyces population densities compared to water collected from the offshore locations in Chichester
Reservoir. In contrast, at Grahamstown Reservoir the observed abundance of Streptomyces in open water samples (surface and bottom) was greater or similar to those detected in marginal water samples.
Common to both reservoirs was the significant change in Streptomyces abundance in water samples between sampling events. Specifically, significantly greater Streptomyces population densities were detected in all water samples collected during the wet sampling period following rainfall events compared to those collected from the same locations during the extended dry period. The overall mean Streptomyces cell densities detected in water samples during the wet sampling period were 178, 919 cells/L and 325, 480 cells/L compared to the dry period, which were 21,879 cells/L and 12,522 cells/L for Chichester and
Grahamstown Reservoir respectively, corresponding to 8.2 and 26 fold differences. The larger wet period abundances observed for all water sample locations at Chichester ranged from approximately 4-fold greater in the north marginal sample (270,921 cells/L [wet] versus
64,262 cells/L [dry]) to 75-fold greater in the north surface water sample (273,534 cells/L
[wet] versus 3,635 cells/L [dry]). At Grahamstown, the differences ranged from a minimum of 2.8-fold larger population densities in the R2 bottom water sample (121,118 cells/L [wet] versus 43,641 cells/L [dry]) up to 295-fold greater abundances in the R6 marginal water sample (673,561 cells/L [wet] versus 2,286 [dry]).
In the Chichester water samples, the proportion of vegetative cells was greater during wet conditions, with an overall mean of 82% and varying from 36% (north surface water) to
100% in some samples (south marginal and bottom water samples from both offshore
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locations) (Figure 4.15). During dry conditions, vegetative cells were still the dominant morphological form present in Streptomyces populations and the overall average of 60% was not significantly different to wet conditions, with the lowest and maximal proportion being
25% (north surface water) and 81% (south bottom water) respectively. These findings differed greatly to Grahamstown water samples. During wet conditions, the proportion of vegetative cells out of the total Streptomyces abundance was low, comprising no more than
10% (overall mean 2%) whereas the samples collected during dry conditions contained a much greater vegetative cell proportion, ranging from 13% (western marginal water) to 94%
(R6 bottom water), with an overall average proportion of 54%.
4.3.3.2 Streptomyces abundance and activity in solid environmental substrates
The abundance of Streptomyces was investigated in the soils and sediments at the margins of both reservoirs in addition to bottom sediment collected at the offshore sampling locations.
Streptomyces were successfully detected and quantified in all samples (Figures 4.17 and 4.18 for Chichester and Grahamstown Reservoir respectively). For bottom sediment samples, the observed abundance of Streptomyces was generally found to be lower compared to those measured in marginal soil and sediments, particularly during the dry sampling period. The overall mean Streptomyces abundance in Chichester bottom sediment was found to be 84,972 cells/g (range 48,769 cells/g to 130,737 cells/g). In comparison, marginal soil and sediment populations were up to 10 and 100-fold greater, with the overall mean population density being 1,890,018 cells/g (range 18,965 to 10,944,850 cells/g). Similarly, Streptomyces populations in Grahamstown marginal soils and sediments were greater compared to bottom sediment populations, with overall mean abundances of 341,885 cells/g (range 78,105 to
2,034,284 cells/g) and 44,148 cells/g (range 7,848 to 126,080 cells/g) respectively.
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Figure 4.17 Mean cell densities of Streptomyces in Chichester Reservoir solid substrate samples based on qPCR targeting genus- specific 16S rRNA sequences. Standard error bars based on duplicate samples and triplicate qPCR results are shown (n=6). Asterisks indicate significant differences in abundance for the same sample between wet and dry sampling events based on t-tests (p<0.05). Results not connected by the same number or letter (for wet and dry sampling period respectively) indicate significant differences between samples based on the ANOVA tests (p<0.05) (top graph). The bottom stacked column graph shows the proportion of vegetative cells and spores comprising the total Streptomyces population density for all samples in each sampling event. 117
.
Figure 4.18 Mean cell densities of Streptomyces in Grahamstown Reservoir solid substrate samples based on qPCR targeting genus-specific 16S rRNA sequences. Standard error bars based on duplicate samples and triplicate qPCR results are shown (n=6). Asterisks indicate significant differences in abundance for the same sample between wet and dry sampling events based on t-tests (p<0.05). Results not connected by the same number or letter (for wet and dry sampling period respectively) indicate significant differences between samples based on the ANOVA tests (p<0.05) (top graph). The bottom stacked column graph shows the proportion of vegetative cells and spores comprising the total Streptomyces population density for all samples in each sampling event.
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Population densities of Streptomyces in soils and sediments at the margins of the reservoirs were found to change between wet and dry conditions. In Chichester, all marginal soil and sediment samples collected during the dry sampling period contained significantly greater abundances than the corresponding wet period samples (Figure 4.17). The overall average abundance of 2,469,561 cells/g observed during the dry sampling period was profoundly greater than the density obtained during wet conditions (114,986 cells/g). In Grahamstown, the mean density of Streptomyces in these samples during dry conditions (441,191 cells/g) exceeded that obtained in wet conditions (241,851cells/g), although the difference was comparatively less and not statistically significant.
The Streptomyces population in the south soil sample from Chichester was 2.5 times greater during dry conditions (792,900 cells/g) compared to wet conditions (306,806 cells/g). A 100- fold increase in abundance was detected in northern soil, from 39,285 cells/g to 10,944,850 cells/g. South marginal sediment, which was exposed during the dry sampling period contained a Streptomyces density 18.5 times greater (2,439,088 cells/g) compared to wet conditions when the sediment was submerged (131,293 cells/g). Similarly, Streptomyces abundance in sediment from the north marginal site when exposed was 23.6 times greater
(446,960 cells/g) compared to submerged conditions (18,965 cells/g). In contrast, no significant changes in Streptomyces densities were detected in bottom Chichester samples between sampling events. Similar findings were also observed but to a lesser extent in
Grahamstown Reservoir. Streptomyces densities detected in R2 and R6 soil samples were significantly greater during dry conditions compared to wet conditions, being 4.2 and 2-fold greater respectively (2,034,284 cells/g [dry] versus 485,205 cells/g [wet] and 147,401 cells/g
[dry] versus 78,105 cells/g [wet] respectively). However, no significant changes in
Streptomyces densities were observed for western soil between wet and dry conditions
(118,255 cells/g [dry] and 86,027 cells/g [wet] respectively). This was also observed for the 119
R2 marginal sediment samples whereas for R6 and western marginal sediment, the measured
Streptomyces abundances were significantly greater during wet conditions. Bottom sediments in Grahamstown (R2 and R12) showed some significant variation between wet and dry conditions, but the disparities were smaller compared to marginal soils and sediment and no significant difference was observed between the overall mean bottom sediment
Streptomyces densities between dry and wet conditions (49,645 cells/g [dry] versus 38,651 cells/g [wet]).
The proportion of vegetative cells out of the total Streptomyces population remained relatively constant between wet and dry conditions in Chichester marginal soil and sediment samples (overall mean values of 40% and 43% respectively) but varied between samples, ranging from 13% (north sediment) to 84% (north soil) (Figure 4.17). Bottom sediment samples contained a lower proportion of cells that were vegetative (2% to 27%), which were slightly higher during the wet conditions (overall mean of 18%, range 9% to 27%) compared to dry conditions (overall mean of 3.5%, range 2% to 5%). In contrast, relative constancy in the proportion of vegetative cells was observed between wet and dry sampling events for bottom sediment samples in Grahamstown with overall mean values of 12% (range 5% to
19%) and 10% (range 3% to 17%) respectively (Figure 4.18). The overall average proportion of vegetative cells detected in marginal soil and sediment samples in Grahamstown during wet conditions (5%, range 1% to 27%) was found to be lower compared to dry conditions
(24%, range 9% to 100%).
Plant debris collected from the margins of Chichester Reservoir were found to support large
Streptomyces populations comparable to those detected in marginal soil and sediment, with an overall average abundance of 422,164 cells/g (Figure 4.17). Similar to sediment and soil
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samples, significantly greater numbers were detected in debris during dry conditions when these substrates were exposed compared to wet conditions. Densities of this genus were 5.5- fold (179,035 cell/g [wet] versus 991,878 cells/g [dry]) and 35-fold greater (14,534 cells/g
[wet] versus 503,208 cells/g [dry]) during dry conditions in south and north marginal debris samples respectively. Furthermore, the proportion of vegetative cells was greater during dry conditions, changing from 40% to 79% and from 27% to 76% for south and north marginal debris respectively.
A variety of macrophyte samples were also obtained from marginal sampling sites to examine
Streptomyces populations. In Chichester, macrophytes (E. sphacelata) supported comparatively lower densities of this genus (ranging from 10,033 cells/g to 14,525 cells/g) compared to other substrates, with no significant differences between samples or between sampling events for the same macrophyte samples (Figure 4.17). Vegetative cells comprised a higher overall mean percentage of the total Streptomyces population during wet conditions
(40%, range 13% to 66%) compared to dry conditions (5%, range 0% to 11%) on these substrates. The three macrophyte species collected along the margins of Grahamstown
Reservoir (Nuphar sp., E. sphacelata and P. repens) also contained several orders of magnitude lower Streptomyces population densities compared to most other solid substrate samples (Figure 4.18). P. repens supported significantly larger populations (10,077 cells/g to
17,478 cells/g) compared to Nuphar sp. and E. sphacelata samples (1,790 to 3,949 cells/g and 2,360 to 3,868 cells/g respectively). The overall mean density detected on macrophytes during dry conditions (7,209 cells/g) was not significantly different to that measured during wet conditions (5,964 cells/g). In contrast to Chichester, a higher percentage of vegetative cells were detected in populations on macrophytes in Grahamstown during the dry sampling period (overall mean 60%, range 46% to 100%) compared to wet conditions (overall mean
3%, range 0% to 7%). 121
4.3.4 Enumeration of Streptomyces on selective growth media and a comparison with qPCR-determined abundance
Streptomyces-like isolates were detected in the majority of samples using selective growth media. Raw data (in Appendix D) of the culturable counts expressed as colony forming units
(CFU) per gram or litre did not reveal any patterns similar to the qPCR data on Streptomyces abundance. Figures 4.19 and 4.20 show the percent of CFU of Streptomyces-like colonies obtained for each isolation media compared to the qPCR data on Streptomyces abundance for all samples in Chichester and Grahamstown Reservoir respectively. It can be seen that for Grahamstown Reservoir, CFU represented between 0% to 87% of the numbers determined by qPCR and for Chichester Reservoir, between 0% and 100%. The comparison of the CFU data with qPCR on Streptomyces abundance revealed the large discrepancies between the two methods of bacterial quantification. For some samples, highly consistent results were obtained between the two methods (e.g. north soil during the wet sampling period). However, the CFU data obtained for the majority of samples only represented a small percentage of the Streptomyces densities determined by qPCR. Despite the addition of fungal and bacterial inhibitors, many of the plates, particularly those containing the macrophyte samples, were overgrown by fungi and other bacteria, impeding the growth of Streptomyces colonies.
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100 SCA (Wet) SCA (Dry) AIA (Wet) AIA (Dry) 90 80 70 60 50 40 30 20 10
0 Percent CFU/qPCR CFU/qPCR Percent
Sample ID Figure 4.19 A comparison of Streptomyces-like colonies enumerated (CFU) on starch-casein agar (SCA) and actinomycete isolation agar (AIA) from all environmental samples obtained from Chichester Reservoir to the qPCR data on Streptomyces abundance. Comparisons are based on the percent ratio of CFU to Streptomyces densities determined through qPCR.
100 SCA (Wet) SCA (Dry) AIA (Wet) AIA (Dry) 90 80 70 60 50 40 30 20 10 Percent CFU/qPCR CFU/qPCR Percent 0
Sample ID
Figure 4.20 A comparison of Streptomyces-like colonies enumerated (CFU) on starch-casein agar (SCA) and actinomycete isolation agar (AIA) from all environmental samples obtained from Grahamstown Reservoir to the qPCR data on Streptomyces abundance. Comparisons are based on the percent ratio of CFU to Streptomyces densities determined through qPCR.
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4.3.5 Streptomyces-like isolates: identification and ability to produce geosmin and 2-MIB
An important component of this study was the identification of Streptomyces-like isolates obtained from a range of environmental samples on the basis of their 16S rRNA sequences in order to examine the diversity of this genus in the reservoirs. Furthermore, the ability of these isolates to produce geosmin and 2-MIB was examined in order to establish their potential to contribute to geosmin and 2-MIB production in the reservoir environments.
Difficulties in isolating pure Streptomyces colonies, due to overgrowth by other organisms despite the supplementation of selective isolation media with antifungal and antibacterial agents, meant that only a fraction of Streptomyces-like colonies obtained could be characterised.
A partial length 16S rRNA gene sequence of between 749 and 861 nucleotides was obtained for each isolate and subject to BLASTn search to identify close matches. Of the 23 isolates obtained from environmental samples that showed Streptomyces-like macroscopic and microscopic features, based on 16S rRNA sequencing and GenBank comparison, 22 were identified as belonging to the genus Streptomyces, whilst one (STR18) was found to belong to the actinobacterial genus Nocardia. The tentative identification of each isolate based on
16S rRNA gene sequences is listed in Table 4.7. Based on BLASTn results, all isolates showed 99% sequence similarity with previously reported Streptomyces strains (and
Nocardia). The cut-off point 97.5% 16S rRNA sequence similarity has been suggested for species definition (Stackebrandt & Goebel, 1994). The type strain sequences of closely related organisms were chosen and were used as reference sequences in the phylogenetic tree
(Figure 4.21), showing the relationships of environmental isolates to reference strains of the genus. S. coelicolor A3(2) was included in the phylogenetic analysis and sequences from the genus Nocardia served as the outgroup.
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Table 4.7 Results from a BLASTn search of the 16S rRNA sequences of the environmental isolates revealing the tentative identification of each. Isolate Length of Closest hit in Genbank Similarity ID nucleotides (accession number) (%) (bp) STR0 821 Streptomyces antibioticus NBRC 13271 (AB184340) 99 STR3 749 Streptomyces sioyanensis IHBB5562 (KF475886) 99 STR4 762 Streptomyces nigrescens DSM40276T (HG794417) 99 STR6 835 Streptomyces parvus 3151 (EF063462) 99 STR7 764 Streptomyces tubercidicus 14241 (EF371435) 99 STR9 754 Streptomyces badius CB00830 (HF935087) 99 STR11 751 Streptomyces chattanoogensis CGMCC 4.1415 99 (JN566019) STR12 764 Streptomyces tubercidicus 14241 (EF371435) 99 STR13 746 Streptomyces tubercidicus 1492 (EF371435) 99 STR14 760 Streptomyces bungoensis NRRL B24305 (AY999905) 99 STR15 760 Streptomyces chattanoogensis OSI-44 (FN 99 STR16 754 Streptomyces olivochromogenes 99 IHBB12025(KF475820) STR17 750 Streptomyces sioyaensis IHBB5562 (KF475886) 99 STR18 861 Nocardia anaemia IFM0323 (NR_041010) 99 STR19 836 Streptomyces canus BYB (JQ700298) 99 STR20 759 Streptomyces chattanoogensis GP4 (JF827350) 99 STR22 762 Streptomyces chattanoogensis GP4 (JF827350) 99 STR25 770 Streptomyces nigrescens DSM 40276T (HG794417) 99 STR26 836 Streptomyces gramineus JR-43 (NR_109017) 99 STR27 755 Streptomyces chattanoogensis GP4 (JF827350) 99 STR28 761 Streptomyces padanus MTKK-103 (AY941253) 99 STR29 762 Streptomyces parvus 3151 (EF063462) 99 STR30 749 Streptomyces badius CB00830 (HF935087) 99
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Figure 4.21 Phylogenetic relationship between 23 actinobacteria (22 Streptomyces, 1 Nocardia) environmental isolates and related sequences based on 16S rRNA gene sequences (neighbour-joining method). Bootstrap values are from 2000 replicates, with values >50% shown. The bar represents 1% sequence divergence.
DNA extracted from all isolates was also amplified using the primer pair used in the qPCR protocol. All isolates confirmed as Streptomyces based on the sequencing results produced the correct sized PCR product (99 bp), whereas no amplicon was generated using template
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DNA extracted the Nocardia isolate, confirming the specificity of the primer pair for detecting Streptomyces (Figure 4.22).
Figure 4.22 Agarose gel electrophoresis of the 99 bp PCR product amplified with the Streptomyces-specific 16S rRNA targeting primers using template DNA extracted from the 23 environmental isolates. Both negative and positive (S. coelicolor A3(2), SC) controls are shown in the furthest right hand lanes. The DNA ladder in the first lanes represents 100 bp intervals (100-1000 bp).
In order to test for geosmin and 2-MIB production ability of the environmental isolates, a simple procedure for the extraction of these secondary metabolites from cultures grown on solid media was developed. The recovery efficiency of this method was assessed by measuring known concentrations of compounds on spiked agar plates and are shown in Table
4.8. Additionally, the recovery efficiency of the LLE protocol developed for extraction from liquid media is presented. This extraction procedure was applied to quantify production of
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these earthy-musty secondary metabolites in sterilised reservoir water (section 4.2.7). Both extraction protocols demonstrated high recovery efficiencies over the concentration ranges tested, for geosmin (81.5-94.6%) and 2-MIB (72.2-93.7%) in the case of the solid culture extraction procedure and up to 96% for both compounds for the LLE method. Geosmin and
2-MIB were identified by GC-MS in the culture extracts of all 22 Streptomyces and the single
Nocardia environmental isolates grown on SCA for 7 days at 28°C. Production yields are shown in Figure 4.23.
Table 4.8 Recovery efficiency of geosmin and 2-MIB from solid and liquid media using the hexane extraction procedures. Mean percent recoveries and the standard deviation are shown (n=3). Extraction Procedure Recovery (%) 2-MIB Geosmin Extraction from solid media Quantity of compound spiked on agar (ng) 100 93.72 ± 6.11 94.62 ± 3.85 25 88.35 ± 6.41 86.38 ± 5.99 5 72.12 ± 8.19 81.49 ± 16.93 Extraction from liquid media (LLE) Quantity of compound in water (ng/L) 500 93.48 ± 14.45 96.64 ± 13.80 100 96.61 ± 7.95 95.73 ± 12.67 50 85.73 ± 5.61 88.85 ± 14.46
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* 160 2-MIB Geosmin 140
120
100 *
80 * * 60 *
Concentration (ng/mg dw) (ng/mg Concentration 40 *
20 * * 0
Isolate ID Figure 4.23 Production yields of geosmin and 2-MIB by 23 environmental isolates on SCA at 28°C for 7 days. Mean and standard error values are shown (n=2). The asterisks indicate that the production of one compound is significantly (p<0.05) greater than the other for each isolate.
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While all isolates were confirmed as producers of geosmin and 2-MIB, production yields were highly variable between these. The maximum yields of geosmin (33.87 ng/mg) were recovered from cultures of the isolate STR16 (S. olivochromogenes IHBB12025) followed by STR28 (S. padanus MTKK 103, 29.68 ng/mg) and STRO (S. antibiotics AB184340, 25.21 ng/mg). Higher yields of 2-MIB were observed with maximal production detected in the culture extracts of STR28 (S. padanus MTKK103, 131.19 ng/mg) followed by STR4 (S. nigrescens DSM40276T, 94.18 ng/mg), STR7 (S. tubercidicus 14241, 64.91 ng/mg), STR27
(S. chattanoogensis GP4, 42.89 ng/mg) and STR29 (S. parvus 3151, 35.57 ng/mg). STR28, a soil isolate from Grahamstown western marginal sampling site, demonstrated the greatest overall potential for contributing to the occurrence of T&O. Photos of the colony morphology and cell structure determined by light microscopy of several of the isolates which demonstrated particularly high production potential of geosmin and/or 2-MIB are shown in
Figure 4.24. Although all isolates demonstrated geosmin and 2-MIB producing potential, many produced significantly greater amounts of one compound compared to the other. For example the Grahamstown surface water isolate STR27 (S. chattanoogensis GP4) produced
20.6 times more 2-MIB than geosmin while a Chichester marginal sediment isolate STR16
(S. olivochromogenes IHBB12025) produced 7.8 more geosmin than 2-MIB. The overall average production of 2-MIB (16.86 ng/mg) exceeded that of geosmin (6.94 ng/mg), with five isolates producing significantly greater amounts of 2-MIB than geosmin and only two producing significantly more geosmin. A number of the isolates were found to match the same 16S rRNA sequence type strains based on the BLAST analysis. These isolates, given the same tentative identity, were obtained from different locations within the same reservoirs.
Despite being identified as the same strains, many of these isolates varied significantly with respect to odour production.
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Figure 4.24 Streptomyces isolates and the Nocardia isolate which demonstrated high geosmin and/or 2-MIB producing potential. Light microscope images (1000x) are shown above and colony morphology directly below for each isolate.
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4.3.6 Geosmin and 2-MIB production by Streptomyces on environmental substrates
Despite the demonstration of geosmin and 2-MIB production ability by the Streptomyces isolates when grown on starch-casein agar (SCA), it is important to recognise that the nutrient composition provided in laboratory growth media often greatly differs to the available nutrients encountered in the natural environment. Additionally, while it was shown that
Streptomyces do indeed occur in the reservoir habitats examined, whether or not these substrates can support their production of geosmin and 2-MIB is an important consideration.
Thus, the ability of selected substrates and water samples to support the growth of two
Streptomyces spp. (S. coelicolor A3(2) and the STRO isolate tentatively identified as S. antibioticus) was experimentally investigated.
The recovered yields of geosmin and 2-MIB produced by both Streptomyces spp. on all environmental solid substrate media and in sterile reservoir water are shown in Figure 4.25.
Reservoir water and all solid substrates (50 g/L dw) supplemented in 1.5% w/v water agar demonstrated, to different extents, the ability to support geosmin and 2-MIB production by both Streptomyces spp. with quantities produced being comparable to those detected on laboratory growth medium (SCA). For both Streptomyces spp., significantly greater quantities of both compounds were recovered from cultures in sterile marginal reservoir water. For S. coelicolor A3(2), these yields for geosmin and 2-MIB were 26.5 ng/mg and
46.5 ng/mg respectively and for S. antibioticus, 14.8 ng/mg and 25.8 ng/mg respectively.
Similarly, high production yields (particularly for 2-MIB) were also obtained when culturing these bacteria in sterile surface water collected in Grahamstown Reservoir compared to the majority of solid substrates (particularly in the case of 2-MIB). Significant differences in yields for cultures grown on solid environmental substrates were not profound and generally all showed similar capacities to support earthy-musty secondary metabolite production by these organisms. The macrophyte sample supported the lowest production of 2-MIB and 132
geosmin by both Streptomyces spp. (1.0 ng/mg and 0.9 ng/mg and 0.7 ng/mg and 1.1 ng/mg for S. coelicolor A3(2) and S. antibioticus respectively). This compared to the maximal yields of 2-MIB and geosmin detected in cultures grown on solid environmental substrate media
(bottom sediment from Chichester Reservoir), being 2.5 ng/mg and 10.8 ng/mg for S. coelicolor A3(2) and 2.4 ng/mg and 5.2 ng/mg S. antibioticus respectively.
Figure 4.25 Yields of geosmin and 2-MIB recovered from S. coelicolor A3(2) and S. antibioticus (STR0) cultures on growth media containing environmental samples from Chichester (Chi) and Grahamstown (Gra) as the sole source of nutrition. Mean values ± standard error are shown (n=3). Bars not connected by the same letter (2-MIB) or number (geosmin) are significantly different (p<0.05).
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4.3.7 Summary of key findings
With regard to the key research questions and hypotheses, the major findings from the field and laboratory work conducted in this study can be summarised as follows:
Streptomyces are widely distributed, abundant and active in many habitats within and
at the margins of both reservoirs. During wet conditions, following significant rainfall
events, population densities of Streptomyces were significantly greater in all water
samples collected compared to extended dry conditions. Vegetative cells comprised
a large proportion of the total cells in Chichester water samples, particularly during
wet conditions. In Grahamstown, vegetative cells generally represented the major cell
form in water samples during dry conditions whereas spores were predominate during
wet conditions.
Streptomyces were generally found to be more abundant and active in the sediment,
soils and plant debris at the margins of the reservoirs compared to bottom reservoir
sediments. Streptomyces abundance was significantly greater in these marginal
substrates, particularly for the samples collected in Chichester Reservoir during dry
conditions when they were exposed due to water level recession, compared to the
corresponding wet period samples under submerged conditions. In Grahamstown
Reservoir, Streptomyces abundance in marginal soils and sediments was generally
higher during dry conditions, with differences being comparatively less profound
than in Chichester Reservoir. Higher activity levels (vegetative cells) were generally
detected in these samples during dry conditions, particularly in Grahamstown
Reservoir. Macrophyte samples were found to harbour comparatively small
Streptomyces populations.
Streptomyces-like colonies enumerated on selective growth media generally
represented only a small fraction of the qPCR determined abundance of Streptomyces
in the majority of environmental samples. 134
All isolates obtained from a variety of environmental samples and identified through
16S rRNA sequencing (22 Streptomyces spp. and a Nocardia sp.) demonstrated
capacity for geosmin and 2-MIB production.
A wide range of environmental substrates as the sole source of nutrition supported
the growth and geosmin and 2-MIB production by two Streptomyces spp., with
recoveries of both metabolites greatest from samples of reservoir water.
4.4 Discussion
Acquiring evidence to assess the relative significance of Streptomyces as a potential contributor to T&O episodes in drinking water reservoirs formed the primary rationale for undertaking the field and laboratory work presented in this chapter. There has been a general disregard in the literature for these bacteria to be aquatic and lack of evidence to firmly link them to T&O production. Combined, the findings of this research indicate that Streptomyces may indeed represent a significant source of geosmin and 2-MIB in Grahamstown and
Chichester Reservoirs.
4.4.1 The potential of Streptomyces to contribute to T&O
The qPCR protocol for quantifying Streptomyces, along with the temporal and spatial sampling program, enabled comparisons of Streptomyces abundance and activity between diverse habitats within the drinking water reservoirs and between wet and dry conditions.
The results obtained concerning population dynamics of Streptomyces provide insight for assessing their potential to contribute to T&O in drinking water supplies and the likely mechanisms by which they do so. These bacteria were found to be widely distributed, abundant and active within the water mass, bottom sediments and at the margins of the drinking water reservoirs, with the multitude of Streptomyces species isolated from these 135
environments all demonstrating a capacity for geosmin and 2-MIB production. Furthermore, a variety of environmental substrates from the reservoirs, including the water mass itself, were found to support the growth and T&O production by Streptomyces spp.
If Streptomyces are to be considered potentially significant contributors to T&O episodes, what are the primary mechanisms by which they introduce their odorous secondary metabolites into the water mass? The findings of this study suggest that two plausible means by which Streptomyces contribute to T&O episodes can be considered. The following discussion elaborates on the findings, including the molecular and culture assays, to support their feasibility. Firstly, the evidence which supports the notion of the ‘wash-in’ of terrestrial
Streptomyces, theoretically accompanied by their odorous compounds, from marginal habitats of drinking water reservoirs into the water mass is presented. The results of this study imply that the potential means by which Streptomyces contribute to T&O events is certainly not restricted to the wash-in of marginal substrates. The discussion continues by drawing on evidence from this study to support the feasibility of the notion that these bacteria may indeed be metabolically active in the water mass and thus are conceivably also capable of contributing to in situ T&O production. Additionally, the discussion focuses on the culture dependent assays concerning enumeration and isolation of Streptomyces from reservoir habitats and their T&O production capacity.
4.4.2 The ‘wash-in’ of Streptomyces and their T&O secondary metabolites from marginal environments into drinking water supplies
It has been widely accepted in the literature that Streptomyces are predominately terrestrial bacteria and find their way into water bodies via run-off following rain events, accompanied by their odorous secondary metabolites (Cross, 1981; Wood
136
et al., 1983a). The data obtained for both reservoirs provided strong evidence to support the ‘wash-in’ hypothesis of Streptomyces, a potential mechanism of contributing to T&O events. Streptomyces abundance was significantly greater in all water samples collected from both reservoirs during wet sampling conditions compared to those collected during the prolonged dry period, with densities more than 100-fold greater in some samples. The significant trend identified in the abundance data for water samples strongly suggests that the presence of Streptomyces in the water mass of both reservoirs is a result of runoff from terrestrial sources following rain events. The total amount of rainfall that occurred over a period of one month prior to each of the sampling events was 10-fold greater during wet conditions compared to dry conditions (refer to
Appendix E).
This possible mechanism of Streptomyces occurrence in freshwater and their contribution to
T&O episodes has been supported by several studies based on culture assays (Jensen et al.,
1994; Lanciotti et al., 2003; Zaitlin et al., 2003b). Lanciotti et al. (2003) detected an increase in the number of filamentous actinobacteria in the River Arno (Italy) during a period of increased flow rate and turbidity following abundant rainfall, the consequence of washing in of the soil and debris. Jensen et al. (1994) traced the source of annual spring occurrence of earthy-musty odour problems in drinking water derived from the Saskatchewan River
(Canada) to filamentous actinobacteria, which, along with their odorous metabolites, were introduced into these surface waters during snowmelt and subsequent runoff. These bacteria were undetectable before spring melting but rose abruptly during the peak runoff of the watershed area in spring and coincided with increasing flow and colour, parameters associated with spring time odour events. Correlations between parameters indicative of terrestrial runoff (e.g. E.coli, turbidity and suspended sediment) and counts of filamentous actinobacteria have been reported in other Canadian water systems (Elbow River Basin and 137
Lake Ontario) (Zaitlin et al., 2003a; Zaitlin et al., 2003b). In such studies, the majority of isolates (predominately Streptomyces) obtained from these freshwater systems were capable of producing geosmin and/or 2-MIB. Furthermore, Parinet et al. (2010) associated the presence of geosmin and 2-MIB in three Canadian lotic water supplies with leaching from soil, as indicated by elevated levels of nitrate, ammonia and faecal coliforms. In Chichester and Grahamstown Reservoirs, some routinely monitored parameters related to runoff (e.g. turbidity, suspended solids, electrical conductivity, nutrients, total coliforms and E. coli) were elevated during wet conditions, supporting the notion of a terrestrial input into the water mass (see Appendix E).
Streptomyces play an important ecological role as saprophytes and are highly adapted to extensively colonising complex organic substrates (Goodfellow & Williams, 1983;
McCarthy & Williams, 1992). They are recognised as one of the most numerous bacteria of soil microbial communities and have been the predominant genus of actinobacteria isolated from plant debris (Jensen, 1971; Lechevalier, 1981; Goodfellow & Williams, 1983; Wood et al., 1983a; Williams et al., 1984; Alexander, 2002). As major producers of geosmin and 2-
MIB, Streptomyces provide soil with its characteristic earthy-musty odour. Thus soils, sediments and plant debris adjacent to freshwater environments undoubtedly represent a source of these compounds which can be washed into the water mass during runoff, which potentially represents a major mechanism of Streptomyces contribution to T&O (Wood et al., 1983a; Jensen et al., 1994). Indeed, all marginal substrates from the reservoirs, when used as the sole source of nutrition, were found to support the growth and geosmin and 2-
MIB production by two Streptomyces spp. in the laboratory.
138
In addition to supporting the notion of the ‘wash-in’ of Streptomyces and theoretically, their odorous metabolites from marginal environments, this study provided evidence to support the hypothesis that the abundance and activity of Streptomyces in substrates located at the margins of reservoirs would be greater during dry conditions when the substrates in these locations become exposed to air following water level recessions, compared to wet sampling conditions when such substrates are submerged. Previous literature reports have suggested that when marginal substrates become periodically exposed during water level recessions,
Streptomyces, being aerobic bacteria, become active and grow profusely under these aerobic conditions. Theoretically, this results in the accumulation of their odorous secondary metabolites and thus represents a potent source of geosmin and 2-MIB which can enter adjacent water bodies with runoff (Adams, 1929; Thaysen, 1936; Wood et al., 1983b).
Indeed, laboratory studies have shown that increasing atmospheric oxygen concentration enhances geosmin production while lower oxygen levels results in decreased geosmin production by cultures of Streptomyces (Dionigi & Ingram, 1994; Schrader & Blevins, 1999).
During the wet sampling event, the storage capacities were maximum (100%) during which, the marginal sediments and plant debris were submerged and soils were waterlogged. During the prolonged period of dry conditions, Chichester Reservoir dropped to 83% (from 156.21 to 154.13 mAHD) and Grahamstown Reservoir to 87% (from 12.10 to 11.96 mAHD) storage capacity. Recession of the water level in both reservoirs exposed the marginal substrates at the shoreline. The comparative analysis of the qPCR data on Streptomyces abundance and activity levels in marginal soil and sediment samples collected between wet and dry conditions provided evidence to support the hypothesis regarding their increased abundance and activity under exposed conditions. Streptomyces population densities in all the soils and sediments examined at the margins of Chichester Reservoir were significantly greater during dry conditions, compared to wet conditions, with differences in abundance up to 100-fold . 139
The average Streptomyces population density in Grahamstown Reservoir marginal sediments and soils during dry conditions also exceeded that detected during wet conditions, although the difference was comparatively less than Chichester Reservoir (maximum of a 4-fold increase). In Grahamstown Reservoir, the overall average proportion of vegetative cells in marginal soils and sediments was significantly higher during dry conditions (24%, range 9% to 100%) compared to wet conditions (5%, range 1% to 27%). Despite population densities increasing during dry conditions in Chichester Reservoir, activity levels remained relatively constant in samples between sampling events, with overall mean values of 40% and 43% for wet and dry conditions respectively.
In the late 1980s, during an extensive dry period, low water levels in Grahamstown Reservoir led to the exposure of shallow margins, followed by refilling and re-submerging of the sediments and shoreline. Coincidentally, complaints of an earthy-musty odour were recorded soon after this event. Based on the systematic evidence provided in this study, it is highly plausible to consider that geosmin and/or 2-MIB were produced by abundant and active
Streptomyces populations at the margins during the exposed conditions followed by the subsequent transport of these odour compounds into the water mass, causing the earthy- musty T&O event observed. Similarly, during a long drought period in 1987, the Vaal Dam
(South Africa) dropped to a quarter of its storage capacity. Some 32 hectares of previously exposed dam sediment was then flooded and the reservoir rapidly re-filled due to extensive rains in the catchment area. Shortly afterwards, an earthy-musty T&O event occurred. The incident could not be attributed to algae or cyanobacteria (Bailey, 1988; Wnorowski, 1992).
Uwins et al. (2007) reported that during a prolonged dry period between 2001 and 2003, the
Hinze Dam (Gold Coast, Australia) fell to below 30% storage capacity, during which high levels of geosmin and 2-MIB were measured in the reservoir. Considering the low abundance of cyanobacterial biomass at the time, the authors suggested that actinobacteria could be the 140
possible source of these T&O episodes during dry conditions. Wood et al. (1985) reported that reservoir sediment which was exposed during maintenance work developed an earthy odour, containing a geosmin concentration of 46 µg/kg dry weight, whereas none was detected in submerged sediment. Furthermore, a sterilised sample of the sediment was found to support geosmin production when inoculated with a Streptomyces sediment isolate.
Plant debris, which comprises a dominant substrate carpeting the margins of Chichester
Reservoir, was also found to support large and active Streptomyces populations comparable to those obtained in marginal sediments and soils. As observed for marginal soils and sediments, the Streptomyces populations inhabiting plant debris were found to be greater (by
5.5 to 35-fold) during dry conditions following the exposure of these substrates, compared to the wet sampling period when these substrates were covered with water. Furthermore, the proportion of vegetative cells was greater by two to three-fold during dry conditions. This indicates that water level recession and the subsequent exposure of plant debris stimulates
Streptomyces populations colonising these substrates to become metabolically active.
Clearly, the wet-dry regime that this marginal Streptomyces habitat experiences would provide an ideal situation for their synthesis of odorous secondary metabolites in exposed aerobic conditions during dry periods, followed by subsequent removal of these compounds by the action of rain, wind, waves and rising water levels into the water mass (Wood et al.,
1983a). It has been recognised for some time, that decaying vegetation found at the margins of rivers, lakes and reservoirs represents a source of T&O compounds produced by
Streptomyces (Sigworth, 1957; Baker, 1961; Waksman, 1967; Raschke et al., 1975; Wood et al., 1985).
141
Plant communities at the margins of rivers, lakes and reservoirs have also been suggested as possible sites of Streptomyces growth and odour production (Thaysen, 1936; Silvey et al.,
1950; Silvey & Roach, 1975) and given the widespread occurrence of emergent macrophytes along the margins of Grahamstown and Chichester Reservoirs, it was considered that these habitats may harbour large populations of Streptomyces. The findings however, suggested that potentially active, but relatively small (<20,000 cells/g) and stable populations of
Streptomyces inhabit these plant surfaces and are unlikely to represent a significant source of
T&O compounds compared to other substrates examined. Supporting this, Hobson (2010) isolated Streptomyces from the sediments of Grahamstown Reservoir using selective growth media but did not detect these bacteria on plant material (P. repens and spike rushes) collected. While geosmin and 2-MIB have been previously detected in biofilms growing on the surface of macrophytes, Streptomyces were not considered the causal microorganisms
(attributed to cyanobacteria) of the odour compounds (Watson & Ridal, 2004; Ridal et al.,
2007).
This is the first study to provide systematic field evidence to support that exposure of substrates at the margins of reservoirs such as soil, sediment and plant debris, may provide suitable conditions for Streptomyces to become active and grow profusely and thus represent a major source of geosmin and 2-MIB when they are rapidly leached out or washed in by run-off and rising water levels during wet conditions. This potential mechanism of
Streptomyces contribution to T&O is further explored under controlled laboratory conditions in Chapter 5.
In contrast to the generally profound differences in Streptomyces abundance in marginal substrates between wet and dry conditions, the population densities and activity levels in the
142
permanently submerged deep bottom sediments remained comparatively constant. Indeed, this observation lends support to the hypothesis that exposure of substrates at the margins is a factor which stimulates the propagation of Streptomyces.
Considering that Streptomyces are thought to be predominately terrestrial bacteria incapable of growing in aquatic environments, it was hypothesised that their abundance and activity in marginal soil and sediment would greatly exceed population densities and activity levels detected in the deep bottom sediments of the reservoirs. This was largely supported by the data obtained, with population densities in soils and marginal sediments observed to be one to two orders of magnitude greater on average than those found in bottom sediments.
Furthermore, the percentage of vegetative cells detected in the bottom sediments was lower, ranging from 1% to 27% compared to marginal sediments and soils, where up to 100% of the population was estimated to be vegetative in some samples. This indicates that
Streptomyces would predominately exist as spores in bottom sediments (Johnston & Cross,
1976a; Willoughby, 1976). Indeed, the bottom sediments collected (predominately mud with a medium to heavy clay texture) had the characteristic odour of hydrogen sulphide, indicating the prevalence of anaerobic conditions, which would be inimical to the growth of aerobic
Streptomyces. Some authors have reported that the primary mycelium of Streptomyces
(which does not produce secondary metabolites) is facultatively anaerobic and this physiological ability may account for the detection of vegetative cells in the bottom sediments of the reservoirs (Higgins & Silvey, 1966; Francisco & Silvey, 1971).
Although the data obtained supports the notion that Streptomyces are relatively inactive and in low abundance in bottom sediment, some reports have suggested that such habitats may be significant sources of T&O derived from Streptomyces (Adams, 1929; Thaysen, 1936;
Issatchenko & Egorova, 1944; Issatchenko, 1946; Bays et al., 1970; Willoughby et al., 1972).
Zuo et al. (2010) concluded on the basis of in situ analysis and the production of geosmin by 143
Streptomyces isolates, that the high concentrations of geosmin in the deep bottom sediments of the Xionghe Reservoir, China (up to 5,280.1 ng/kg dw) were produced by Streptomyces.
Sugiura et al. (1987) reported that the high abundance of filamentous actinobacteria in the carbohydrate rich sediment of Lake Kasumiguara (Japan) were responsible for the occurrence of 2-MIB. A coincidence between high actinobacteria abundance (10-50 x 106 cells/g) and geosmin and 2-MIB concentrations (500-650 ng/kg) in the bottom sediments of the North
Pine Dam (Australia) when the density of cyanobacteria was low, suggested that these bacteria may be producers of T&O compounds in this benthic environment (Klausen et al.,
2004; Nielsen et al., 2006). The authors suggested that the aeration systems deployed in the dam combined with organic matter availability provides an ideal habitat for these bacteria and therefore T&O production. Interestingly, the genome of S. coelicolor A3(2) contains three operons that encode respiratory nitrate reductase typically involved in anaerobic metabolism, indicating that these bacteria may undertake metabolism in anoxic or in low oxygen conditions (Dyson, 2009). Guttman and van Rijn (2008) recently reported the production of geosmin and 2-MIB by two Streptomyces spp. under anoxic conditions, albeit to a lesser extent than under aerobic conditions system and suggested utilisation of nitrate as an electron acceptor.
The notion that Streptomyces are washed into reservoir waters during wet conditions from marginal substrates was supported by this study. Theoretically, their transport into the water mass would be accompanied by their entrained odorous secondary metabolites, representing a likely mechanism by which these bacteria contribute to T&O episodes. Importantly, the data supported the hypothesis that during dry conditions in which these marginal substrates become exposed, Streptomyces populations become more active and conceivably, represent a greater source of T&O compounds which can enter the adjacent water mass following wash-in. The substrates at the margins undoubtedly represent the environment in which 144
Streptomyces proliferate profusely, in comparison to bottom sediments, where Streptomyces populations appear to represent a minor physiologically active component of microbial communities of freshwater environments and are unlikely to significantly contribute to T&O episodes. What about Streptomyces populations in the water mass of drinking water reservoirs? Do Streptomyces exist as metabolically active forms in this environment, capable of directly contributing to T&O episodes through in situ production of their odorous secondary metabolites? Following is an assessment, based on the findings from this study, of the feasibility of this possible mechanism of Streptomyces contribution to T&O.
4.4.3 The potential for in situ production of T&O secondary metabolites by Streptomyces in the water mass
The data obtained in this study and the evidence within the existing literature both clearly indicate the likely significance of terrestrial wash-in as a major source of Streptomyces in reservoir waters and a potential mechanism of contributing to T&O episodes. As indicated by Zaitlin and Watson (2006), there is still debate surrounding whether Streptomyces washed in from the margins of water bodies continue to be active in the water column and are capable of in situ geosmin and 2-MIB production. Several authors contend that these bacteria occur in water only as metabolically inactive spores (Johnston & Cross, 1976a; Cross, 1981; Jensen et al., 1994; Zaitlin et al., 2003b). However, the results obtained here certainly provide evidence to suggest that they may also be autochthonous, numerous and active components of microbial communities in the water mass, albeit to a much lesser extent than in terrestrial environments. Despite lower population densities in the water mass during dry conditions,
Streptomyces were largely present as vegetative cells. Considering that in the absence of rainfall, the wash-in of Streptomyces from surrounding terrestrial environments would be limited, this suggests that they continue to remain metabolically active in the water mass as opposed to existing solely as inactive spores. Interestingly, Grahamstown Reservoir water 145
samples contained a much greater proportion of vegetative cells during the dry period (overall mean of 54% and range 13-94%) compared to the wet sampling event (<10%, overall mean of 2%), the latter possibly resulting from the wash-in of predominately spores. The overall average proportion of vegetative cells in Chichester Reservoir was similar between wet and dry conditions (82% and 60% respectively).
Several studies using molecular techniques including FISH (fluorescence in situ hybridisation) and FISH with catalysed reporter deposition (CARD) targeting rRNA molecules (indicative of metabolically active cells) have demonstrated that active actinobacteria are ubiquitous and can be important components of planktonic microbial communities (Klausen et al., 2005; Nielsen et al., 2006). A wide range (20% to 70%) of the proportion of filamentous actinobacteria, assumed to be Streptomyces, have been reported to be active in various freshwater systems (Klausen et al., 2005). In Danish streams, Klausen et al. (2005) determined the densities of actinobacteria to be between 4 to 38 x 107 cells/L, corresponding to 3% to 9% of the total bacterial population, of which 2.8% to 38% (average
22%) were filamentous and assumed to be Streptomyces (10-60 x 106 cells/L). Interestingly, the authors reported that the proportion of metabolically active actinobacteria increased two- fold during the passage of stream water through aquaculture ponds in addition to an increase by 55% and 110% in geosmin and 2-MIB concentrations respectively. This, in addition to geosmin and 2-MIB production by Streptomyces isolates and the low abundance of cyanobacterial biomass, may indicate that these bacteria were responsible for the odour.
Using the FISH-CARD protocol, Nielsen et al. (2006) identified that abundances of actinobacteria were greater in bottom waters compared to surfaces waters of the North Pine
Dam (Queensland, Australia), with the proportion of active cells being 35% to 80% higher, which the authors attributed to the effects of an aeration system deployed in the reservoir to minimise stratification. 146
While these studies support that filamentous actinobacteria can be active and abundant organisms in fresh water, the drawback of the molecular assays employed is lack of specificity for Streptomyces, the main geosmin and 2-MIB producing genus (Klausen et al.,
2005; Nielsen et al., 2006). The densities of filamentous actinobacteria assumed to be
Streptomyces (10-2,000 x 106 cells/L) in these studies far exceed the range of densities of this genus in water samples (1,829 cells/L to 846,079 cells/L) determined in this study. This suggests that the above studies using FISH have perhaps overestimated the populations of potentially active Streptomyces, which are certainly not the only filamentous and numerous genus of actinobacteria. Streptomyces population densities detected in the water mass align more closely with those recently reported by Lylloff et al. (2012), who used a TaqMan qPCR assay targeting Streptomyces 23S rRNA sequences to quantify this genus in water samples obtained from a river, a weir and two reservoirs in subtropical Australia. This is the only published report identified in the literature where qPCR has been applied to measure the abundance of Streptomyces in aquatic environments. Mean densities of Streptomyces varied from 225 cells/L to 45, 650 cells/L in the surface water and bottom water (8.5 m deep) of one of the reservoirs respectively. The highest density of Streptomyces at a depth of 8.5 m at the bottom of one reservoir was attributed to a large abundance of Streptomyces that probably exist in the sediment-water interface. In this study, the observation of bottom water samples containing greater abundances of Streptomyces, as both spores and vegetative cells, may be due to favourable nutrient conditions at the bottom of the reservoirs and/or the availability of sufficient DO concentration (see Appendix E) which may support their vegetative growth.
Employing two cell lysis methods for DNA extraction to differentiate between vegetative cells and spores established the presence of vegetative cells in water samples, providing evidence to support their potential growth and activity in the water mass. However, it must 147
be emphasised that detection of vegetative cells in these samples cannot provide absolute certainty that these bacteria were growing in situ, due to the possibility of the wash-in of vegetative forms from surrounding terrestrial environments, resuspension from bottom sediments, or their existence as non-viable mycelial fragments. Nevertheless, this is an important finding as it has been widely surmised that dormant spores represent the dominant form of this genus in the water mass of aquatic environments (Cross, 1981; Wood et al.,
1983a; Zaitlin & Watson, 2006).
Further evidence to support the ability of Streptomyces to be active in the water mass and therefore, potentially contribute to in situ T&O production was obtained in the laboratory study which examined the ability of diverse environmental substrates obtained from the reservoirs, including water itself, to support the growth and production of geosmin and 2-
MIB by two Streptomyces spp. The observation that both Streptomyces spp. could grow and produce both secondary metabolites on all environmental substrates, with production rates comparable to those obtained on laboratory media, was not surprising given their nutritional versatility, allowing them to grow on both nutrient-rich and nutrient-poor carbonaceous substances (Wood et al., 1983a; McCarthy & Williams, 1992). This suggests that T&O episodes resulting from these bacteria could originate from a multiplicity of habitats in freshwater environments as opposed to there being a specific or favourable habitat in which their growth and earthy-musty odour production dominates. However, the most significant finding from this laboratory study was that that sterilised reservoir water supported both vegetative growth and production of geosmin and 2-MIB by both Streptomyces spp. This was an interesting observation, considering that Streptomyces are regarded as being adapted to growth on solid substrates in addition to the fact that their growth and production of odorous secondary metabolites in the water mass has been largely disputed in the literature (Johnston
& Cross, 1976a; Cross, 1981; Zaitlin et al., 2003b). Based on the observed geosmin and 2- 148
MIB production rates by the Streptomyces spp. in sterilised reservoir water, less than one milligram of dry mycelial biomass in one litre of reservoir water could theoretically produce detectable earthy-musty odours (>10 ng/L).
It has been argued that the water mass of most lakes and reservoirs does not provide a suitable environment for the growth of Streptomyces and production of odour causing compounds
(Cross, 1981; Williams et al., 1984; Wood et al., 1985). Wood et al. (1985) demonstrated that natural oligotrophic reservoir water did not support geosmin production by S. albidoflavus, which required a source of nutrient enrichment. However, Grahamstown and
Chichester Reservoirs are generally mesotrophic, which evidently provides sufficient nutrient levels to support the growth of Streptomyces spp. Interestingly, Zaitlin et al. (2003b) found that geosmin production by a Streptomyces isolate was equivalent when grown in sterile river water compared with an optimal laboratory growth medium (Czapek’s agar).
Although the reservoir water was incubated at 28°C, as revealed in Chapter 6, these
Streptomyces spp. were also found to grow and produce both compounds favourably at 24°C, perhaps a more realistic environmental temperature.
As suggested by Lylloff et al. (2012), the significance of Streptomyces as potential contributors to geosmin and 2-MIB episodes in reservoirs can be tentatively estimated from known production rates in laboratory cultures. Klausen et al. (2005) determined cell-specific production rates by selected Streptomyces strains isolated from aquatic environments to be
0.1-30 x 108 g/cell/h for both compounds. Based on these theoretical production rates and densities of vegetative (potentially active) Streptomyces cells detected by qPCR in water samples, the largest concentration produced over the course of a week would be 1.47 ng/L.
Thus, population densities in water samples are probably too low to account for earthy-musty
149
T&O events. However, based on vegetative cell densities, marginal substrates could produce up to 46 ng/g per week, thus representing a significant source of T&O compounds which could enter water via wash-in. However, such theoretical estimates do not take into consideration loss of these compounds through processes such as volatilisation and biodegradation. In addition, it does not take into account that not all Streptomyces produce geosmin and/or 2-MIB (Juttner & Watson, 2007). Calculation of these potential production rates based on Streptomyces abundance measured in samples collected suggests that these bacteria inhabiting the water column, despite being potentially active, are unlikely to be major sources of geosmin and 2-MIB in these reservoirs. Rather, marginal habitats are likely to be sites where these compounds accumulate and are eventually washed into water producing T&O episodes.
4.4.4 Molecular versus culture-dependent detection and quantification of Streptomyces
The qPCR protocol enabled Streptomyces to be detected and quantified in samples from all investigated habitats, from the water mass, bottom sediments to a variety of substrates located at the margins, suggesting a widespread distribution of these bacteria in the reservoirs. This finding is in good agreement with published reports describing habitats where the genus has been enumerated and isolated using culture-dependent techniques (Wood et al., 1983a;
Zaitlin & Watson, 2006). Until recently, the latter has been the predominant approach used for examining and monitoring aquatic Streptomyces populations to assess their potential role as contributors to T&O. The densities of Streptomyces measured by qPCR targeting this genus relative to the colony counts enumerated on selective growth media in this study revealed that the traditional culture-dependent method greatly underestimates the abundance of Streptomyces in the environment. Despite efforts to improve the isolation of environmental
Streptomyces including the use of selective carbon and nitrogen sources and incorporation fungal and bacterial inhibitors in growth media, the vast majority of Streptomyces were 150
unculturable (Lingappa & Lockwood, 1961; Kuster & Williams, 1964; Hsu & Lockwood,
1975; Williams et al., 1984; Kampfer, 2006). Importantly, the dilution plate count method provides no information on the activity of Streptomyces in their environment of isolation as dormant spores and active mycelial forms, which both give rise to colonies, cannot be differentiated. Furthermore, Streptomyces take a long time to develop on growth media and often are outcompeted by other bacteria and fungi, hampering their development. In comparison, the qPCR approach allows for the detection of low cell numbers, unique identification of Streptomyces and a short analytical turnaround period that can be applied for rapid monitoring of Streptomyces populations such as during T&O episodes (Rintala &
Nevalainen, 2005).
Concerning the molecular approach, a potential problem when the 16S rRNA gene is used for microbial quantification attempts on complex environmental samples is the issue of variation in the number of 16S rRNA operons in different species. This issue would introduce some variation in the abundance values obtained in samples containing Streptomyces with different copy numbers of the 16S rRNA gene. As long as the number of operons is not known, the conversions to actual cell numbers have to be interpreted with caution (Smith &
Osborn, 2008). Based on the primer-BLAST results, the majority of Streptomyces strains
(including S. coelicolor A3(2)) contained six copies per genome of the 16S rRNA gene, however the number does vary. Copy number of the Streptomyces isolate (tentatively identified at S. antibioticus) is unknown, but the positive agreement between the two qPCR assay calibration curves (Figure 4.14) suggests the same 16S rRNA copy number in both species. An assumption in this study was the same copy number of 16S rRNA sequences in
Streptomyces detected in environmental samples (i.e. six copies per genome). The primer-
BLAST results also indicated that the primers were not completely complementary to the
16S rRNA sequences of some Streptomyces strains available in the NCBI GenBank database 151
(some mismatches) and may underestimate the abundance of this genus. However, given that
DNA obtained from the 22 confirmed Streptomyces isolates could generate the correct PCR product with these genus-specific primers (Figure 4.22) indicates their reliability. Another potential issue with using qPCR to quantify Streptomyces populations is the detection of the target sequence in non-viable cells or extracellular DNA, as such detections can obviously result in an overestimation of viable cell densities (Bae & Wuertz, 2009).
4.4.5 Streptomyces isolates: identification and geosmin and 2-MIB producing ability
Despite the deficiencies mentioned, culture-dependent methods can indicate the potentially active Streptomyces populations in the environment and importantly, allow for the laboratory isolation of taxa to examine the diversity of species and to test for geosmin and 2-MIB production ability. For these reasons, traditional plate counts are still widely used and in this study, allowed for the isolation and identification of a variety of Streptomyces spp., indicating the diversity of this genus across drinking water reservoir environments (Zaitlin & Watson,
2006).
In this study, all of the 22 individual Streptomyces isolates recovered from reservoir samples were confirmed as producers of both geosmin and 2-MIB. Interestingly, numerous studies investigating the geosmin and 2-MIB producing ability of aquatic Streptomyces isolates have largely reported production of one compound or the other, and less frequently, production of both compounds by any one isolate (Zaitlin et al., 2003b; Tung et al., 2006; Zuo et al., 2009a;
Zuo et al., 2010; Lee et al., 2011; Uwins, 2011). Often, Streptomyces isolates have been found incapable of producing either compound indicating that not all Streptomyces are producers of earthy-musty secondary metabolites. Zaitlin et al. (2003b) found that of the
Streptomyces isolated from Lake Ontario (Canada), 23% produced geosmin only, 14%
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produced 2-MIB only and 41% produced both compounds, whereas the remainder produced no detectable levels of either secondary metabolite. Jensen et al. (1994) isolated 102
Streptomyces-like organisms from the North Saskatchewan River, observing that 30% produced geosmin only, 25% produced both geosmin and 2-MIB, 3% produced 2-MIB only and a large fraction of the isolates (42%) did not produce either secondary metabolite. Despite the relatively low number of isolates examined in this study, confirmation of all of them as geosmin and 2-MIB producers provides strong evidence to suggest that Streptomyces could indeed be contributing to T&O episodes in the drinking water supplies investigated.
Similar to previous studies, considerable variation in the biomass-specific capacity to produce geosmin and 2-MIB among the individual isolates was observed. The largest concentrations of geosmin and 2-MIB produced were in the order of 30 and 84 times greater respectively than the smallest quantity measured. Kenefick et al. (1992) reported 200-fold differences in production rates by Streptomyces whereas Zaitlin et al. (2003b) observed 100- and 300-fold differences in geosmin and 2-MIB respectively. The range of yields produced by the isolates aligns with those reported in previous studies (Sugiura & Nakano, 2000; Pan et al., 2009; Zuo et al., 2009b).
Although all isolates demonstrated capacity to produce both compounds, many produced significantly greater amounts of one compound over the other. The strong ability of a number of isolates to produce 2-MIB indicates that Streptomyces may be a significant source of this compound in the reservoirs, since the historical water quality data analysis presented in
Chapter 3 revealed no relationship between 2-MIB and any other monitored biological parameter. Many of these high 2-MIB producing isolates were obtained from sediment samples, which is a significant finding as a previous study reported that Streptomyces spp.
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isolated from Grahamstown Reservoir sediment were only producers of geosmin, suggesting that these bacteria were not the source of 2-MIB (Hobson et al., 2010). Several studies, based on correlations of biomass with odour intensity and/or in situ production ability, have provided evidence to suggest that Streptomyces are a significant source of 2-MIB in freshwater systems (Sugiura & Nakano, 2000; Pan et al., 2009; Zuo et al., 2009b).
It is also worthy to mention that a number of isolates obtained from different locations within the reservoirs had the same tentative identity based on 16S rRNA sequence comparisons, indicating the widespread distribution of Streptomyces species. Despite being identified as the same strains, many of these isolates varied significantly with respect to odour production.
Klausen et al. (2005) also reported considerable differences in odour production by three
Streptomyces isolates which had similar 16S rRNA gene sequences. Whether this divergence in geosmin and 2-MIB production reflects geno- or phenotypic differences among the isolates is unknown. This study also indicated that Nocardia, another genus of actinobacteria, could potentially contribute to T&O episodes. Among the literature, geosmin producing Nocardia have been reported in relatively few studies (Gerber & Lecheval, 1965; Schrader &
Summerfelt, 2010). Schrader and Summerfelt (2010) isolated two Nocardia spp. (N. cummidenlens and N. fluminea) from biosolids in a RAS, and owing to their high capacity for geosmin production, they were considered responsible for the presence of this earthy odour in the RAS water.
4.5 Conclusions
Combined, the findings obtained from the field and laboratory studies presented in this chapter have provided evidence to suggest that Streptomyces may represent a potentially significant contributor to T&O episodes in Grahamstown and Chichester drinking water 154
reservoirs. Not only were these bacteria found to be widely distributed, abundant and active within the water mass, bottom sediments and at the margins of both drinking water reservoirs, all Streptomyces cultivars isolated from these environments demonstrated the capacity to produce both geosmin and 2-MIB. Additionally, environmental substrates collected from such diverse habitats were all found to support the growth of and T&O production by
Streptomyces spp.
Soils, sediment and plant debris at the margins surrounding the reservoirs undoubtedly support the largest Streptomyces populations. The hypothesis that exposure of such substrates to air during water level recession stimulates the growth and activity of these bacteria was largely supported in this study, in which case they could represent a major source of geosmin and 2-MIB which can enter the adjacent water mass. This potentially important mechanism of Streptomyces contribution to T&O is further explored in Chapter 5 through a controlled laboratory study.
The significantly greater Streptomyces cell densities measured in all water samples during wet conditions supports the hypothesis of the wash-in of these bacteria from marginal environments, conceivably entrained with their odorous secondary metabolites. While this indicates the terrestrial origin and the importance of runoff for the occurrence of these microbes in the water mass, evidence for the potentiality of Streptomyces to exist as metabolically active forms in water and thus contribute to in situ T&O production was identified in this study. The detection of Streptomyces vegetative cells in the water mass of both reservoirs, particularly during dry periods when terrestrial wash-in of these bacteria would be minimal, challenges the opinion of Cross (1981) and other authors (Johnston &
Cross, 1976a; Zaitlin et al., 2003b), that Streptomyces are purely ‘terrigenous’ bacteria.
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Furthermore, the observation of geosmin and 2-MIB production by Streptomyces spp. in sterilised reservoir water indicates that they indeed represent a potentially active genus of bacteria in the water mass with the capacity to contribute directly to T&O.
Previous studies attempting to relate Streptomyces to geosmin and 2-MIB in reservoirs have been undermined by the use of culture-dependent techniques which do not allow the discrimination between dormancy (spores) or actively growing vegetative cells. This study is the first attempt to quantify Streptomyces populations in drinking water reservoir habitats using a genus-specific qPCR assay and further discriminate between vegetative cells and spores by employing a differential DNA extraction protocol, thus allowing an estimate of levels of activity. Future studies would benefit from employing qPCR in natural water systems to identify and quantify specific organisms of interest behind T&O events and correlate such data with geosmin and 2-MIB measurements. The absence of data on geosmin and 2-MIB concentrations in the samples examined is perhaps a limitation of this research in attempting to elucidate the contribution of Streptomyces to T&O episodes in drinking water reservoirs. Nonetheless, the laboratory studies which confirmed production capacity of both geosmin and 2-MIB by all Streptomyces isolates and the ability of environmental substrates to support production of these secondary metabolites by two Streptomyces spp., provides strong evidence to suggest that these bacteria are indeed potential contributors to the occurrence of T&O episodes.
There is still much to be learnt about the breadth and depth of T&O issues in the Australian water industry, despite it being accepted as a common problem. There has particularly been a major deficiency in the literature regarding the role of Streptomyces. The findings of this study certainly indicate a potential role for Streptomyces in T&O events. Aside from the
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wash-in of marginal substrates containing these bacteria and their entrained metabolites, it is possible that their active growth and secondary metabolic activity in the water mass may also contribute to T&O. The results obtained provide suitable scientific knowledge and understanding to enable water utilities to make evidence-based decisions to aid in the management of and prevent or minimise T&O problems due to Streptomyces. Unfortunately, the key factors seem to be exposure of marginal habitats due to water level recession during extended dry periods and subsequent run-off when rainfall returns – factors over which authorities can have little control. Management options to stabilise shorelines may prevent introduction of marginal substrates entrained with Streptomyces and their odorous metabolites into the water mass during rain events. However, there is a high likelihood that other factors may impede or hinder the production of these compounds in the natural environment, and it may be among these that strategies for preventing such T&O episodes can be found. Further examination of the effect of factors including nutrient concentration and other chemical and physical variables on the production of geosmin and 2-MIB by these bacteria is presented in Chapter 6.
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CHAPTER 5 - STREPTOMYCES IN THE MARGINAL SEDIMENTS OF DRINKING WATER RESERVOIRS: A SIGNIFICANT SOURCE OF GEOSMIN AND 2-METHYLISOBORNEOL DURING WATER LEVEL RECESSION?
5.1 Introduction
The water quality data analysis presented in Chapter 3 revealed that the dynamics of geosmin in Grahamstown and Chichester drinking water reservoirs are closely tied with those of
Anabaena spp. abundance. However, not all past episodes of elevated geosmin concentration can be attributed to these organisms. Furthermore, the source of 2-MIB in the reservoirs remained unidentified. Streptomyces have been the suspected causal bacterial genus behind
T&O events which cannot be attributed to cyanobacteria.
The Chapter 4 investigation identified that substrates located at the margins of Chichester and Grahamstown Reservoirs, including soil, sediments and plant debris supported the largest population densities of Streptomyces and would thus represent a significant site of earthy-musty secondary metabolite production. Although occurring in a multiplicity of habitats, such substrates are the greatest natural reservoir of these bacteria. Their filamentous growth habit and extracellular enzyme production capacity enables them to extensively colonise and proliferate on complex polymers (e.g. lignocellulose) that are abundant in these substrates (Goodfellow & Williams, 1983; Hopwood, 2007). A significant finding of the research was that when reservoir water levels recede during extended dry periods, causing marginal substrates to become exposed and thus develop more aerobic conditions, higher abundance and in some samples, higher activity levels of Streptomyces could be detected compared to wet weather conditions when these substrates were submerged or waterlogged.
It has been suggested by several authors (Romano & Safferman, 1963; Goodfellow &
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Williams, 1983; Wood et al., 1985; Bailey, 1988; Wnorowski, 1992) that the former conditions stimulate the profuse growth and production of geosmin and 2-MIB by
Streptomyces and thus represent a significant source of these compounds which can enter the adjacent water mass via seepage, runoff and the subsequent water level rise following rain events, producing T&O episodes. While the data regarding Streptomyces population dynamics presented in Chapter 4 supports this proposition, a limitation of the research was the absence of geosmin and 2-MIB measurements in environmental samples to be able to draw any firm conclusions.
As discussed in Chapter 4, an extended warm dry period during the 1980s caused the lowering of Grahamstown Reservoir water level and consequently, the exposure of the shallow margins. Coinciding soon after refilling and re-submerging of the shorelines were consumer complaints regarding an earthy-musty odour in drinking water. Similar observations have been reported in other reservoirs where the earthy-musty compounds could not be traced to cyanobacteria (Bailey, 1988; Wnorowski, 1992; Uwins et al., 2007).
Additionally, Wood et al. (1985) reported that reservoir sediment which was exposed during maintenance work developed an earthy odour, whereas none was detected in submerged sediment. Early reports by Adams (1929) and Thaysen (1936) suggested that sediment produces earthy odours when exposed. Indeed, Streptomyces are regarded as aerobic bacteria and laboratory studies have shown that increasing atmospheric oxygen concentration promotes geosmin production while lower oxygen levels result in a decrease in its biosynthesis in cultures of these bacteria (Dionigi & Ingram, 1994; Schrader & Blevins,
1999). Thus, conditions of reservoir draw-down may be indicative of periods of increased risk of T&O problems caused by Streptomyces developing in water supplies and warrants further investigation.
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This chapter comprises the details of an experimental-based study, carried out with the objective to further explore this potential means by which Streptomyces inhabiting sediments at the margins of reservoirs may contribute significantly to geosmin and 2-MIB episodes.
The overarching hypothesis of this investigation was that sediments located at the margins of reservoirs represent a significant site of growth and production of geosmin and 2-MIB by
Streptomyces, particularly when such substrates are exposed to air following water level recession, providing the wet aerobic conditions ideal for these bacteria to flourish. Production of these earthy-musty metabolites in submerged shallow sediments at the margins
(representative of when the water level is high), was hypothesised to be comparably less, but not inhibited due to the occurrence of low dissolved oxygen levels. However in submerged sediments under anaerobic conditions, conceivably representative of those located at the bottom of reservoirs, growth and geosmin and 2-MIB production by these obligate aerobic bacteria was expected to be inhibited. These hypotheses were tested in a laboratory simulation using sediment collected from the margins of Chichester Reservoir. Both sterile and non-sterilised sediment was inoculated with the geosmin and 2-MIB producing S. coelicolor A3(2) in addition to an assessment of the capacity of the indigenous microorganisms to produce these compounds.
5.2 Materials and methods
5.2.1 Microorganism
S. coelicolor A3(2) (ATCC BAA-471), a geosmin and 2-MIB producing strain previously employed as a test organism in experiments conducted in Chapter 4, was selected for use in this study. Cultures used for the experiment were maintained on SCA and repeatedly sub- cultured to ensure culture purity. Spores were placed in 20% glycerol solutions for long term storage at -20°C.
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5.2.2 Experimental design
The following experiments were conducted to explore the potential influence of fluctuating water levels in reservoirs on the production of geosmin and 2-MIB by Streptomyces in sediment. Various conditions corresponding to those encountered in marginal and permanently submerged bottom sediments were simulated, including exposed versus submerged and aerobic versus anaerobic, respectively. The sediment used in the experiment was collected from the south marginal sampling location in Chichester Reservoir (dry sampling period 1st October 2013). An analysis of the sediment prior to initiation of the experiment indicated that the initial concentrations of geosmin and 2-MIB were below the limits of detection.
Portions of crude sediment (90 g wet weight) were placed into 250 mL sterile glass beakers to form a sediment depth of approximately 3 cm. The following sediment treatments were applied:
A. Sterilised sediment inoculated with S. coelicolor A3(2)
B. Non-sterilised sediment inoculated with S. coelicolor A3(2)
C. Non-sterilised sediment (biotic control)
Sterilisation of the sediment was achieved by placing 500 g (wet weight) into 1 L Erlenmeyer flasks followed by autoclaving at 121°C for 20 minutes. Sediment treatments A and B were inoculated with 20 mL of a spore suspension of S. coelicolor A3(2) in sterile Milli-Q water prepared from a heavily sporulating culture grown on SCA. Sediment C, which was unsterilised and received no Streptomyces inoculum, served as the biotic control to assess production of geosmin and 2-MIB by the indigenous microorganisms. In triplicate, each of the above sediment treatments were subjected to either submerged conditions (covered by 161
water) or exposed conditions (exposed to air) as shown in Figure 5.1 to examine production of geosmin and 2-MIB. For submerged conditions, beakers were slowly filled with sterile
Milli-Q water free of geosmin and 2-MIB (200 mL), avoiding suspension of the sediment.
For exposed conditions, sediments were supplemented with the small volume of water from the inoculum (20 mL) to moisten the sediment or in the case of the biotic control, sterile water only. Furthermore, sediment treatment A (sterile sediment inoculated with S. coelicolor
A3(2)) was also subjected to anaerobic conditions, representative of those conceivably encountered at the bottom of reservoirs (submerged conditions only). Anaerobic conditions were established in an anaerobic jar (Figure 5.2) using the AnaeroGen™ gas generating system (Oxoid). This involved placing a sachet of oxygen scavenging ascorbic acid inside the jar and immediately clamping the lid. Generation of anaerobic conditions was confirmed using an Anaerotest® methylene blue indicator strip (Merck) placed inside the jar, which turns white upon anaerobiosis within 4-6 hours. All experiments were incubated at 25°C in the dark for seven days (CO2 water jacketed incubator, Forma Scientific).
Figure 5.1 Experimental design for testing the production of geosmin and 2-MIB by each sediment treatment (A, B and C) under submerged and exposed conditions. 162
Figure 5.2 Anaerobic jar used to examine geosmin and 2-MIB production in sterilised sediment inoculated with S. coelicolor A3(2) (Sediment A) under submerged anaerobic conditions.
5.2.3 Dissolved oxygen measurements
After the incubation period, dissolved oxygen (DO) was measured in the submerged sediment experiments using a calibrated probe (HORIBA® D-series pH/DO meter). Measurements were taken by carefully placing the probe 1 cm above the sediment surface to avoid disturbance of the sediment.
5.2.4 Geosmin and 2-MIB measurements
Following the seven day incubation period, the overlying water above the submerged sediments was sucked up slowly avoiding suspension of the sediment, through a glass pipette until the sediment was just exposed. These water samples (50 mL volumes) were analysed for geosmin and 2-MIB measurement. To determine geosmin and 2-MIB concentrations in the sediments, the entire volume of sediment was placed in 250 mL polypropionate bottles
(Beckman) and centrifuged (20 min at 1,270 x g) to obtain interstitial water. Interstitial water 163
(approximately 10-20 mL) was then used for geosmin and 2-MIB extraction. Both interstitial water from the sediments and overlying water samples were extracted using the liquid-liquid extraction (LLE) protocol described in Chapter 4 (section 4.2.7). In brief, water samples were transferred to 100 mL conical flasks and supplemented with 25% (w/v) NaCl. Extraction with 1 mL of hexane as the solvent was achieved using a magnetic stirring device for 30 minutes. To minimise loss of the compounds, flasks were sealed with glass stoppers lined with Teflon (PTFE).
Following extraction, 200 µL of the solvent, supplemented with BSTFA and biphenyl-d10 as the internal standard (20 ng/mL) were transferred to GC vials and sealed with screw cap lids
(Agilent Technologies). Samples were analysed for geosmin and 2-MIB by GC-MS in SIM mode as described previously (section 4.2.6). The analytes were identified based on the retention time and mass spectrum determined using analytical standards. Geosmin, 2-MIB and the internal standard were quantified by integrating the base peak areas using the ions m/z 112, 95 and 164 respectively. The calibration curves presented in section 4.2.6 (Figure
4.9) were used for quantification of the analytes in samples. The concentrations of compounds in sediments were evaluated in terms of the amounts in the interstitial water and expressed on a per dry weight basis of sediment (ng/kg dw). Moisture content of the sediment was determined gravimetrically by freeze-drying to constant weight. The overlying water concentrations were expressed as ng/L.
5.2.5 Quantification of Streptomyces in sediment
The Streptomyces population densities in the sediments for all treatments and under all experimental conditions were determined by qPCR prior to incubation (day 0) and at the end of the seven day incubation period. DNA was extracted from sediment samples (250 mg wet
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weight) using the PowerSoil® DNA isolation kit (MO BIO Laboratories, Inc.). As described previously in Chapter 4 (section 4.2.2), the protocol lyses cells vigorously through mechanical, heat and chemical methods followed by the removal of PCR inhibitors (non-
DNA organic and inorganic material) using patented solutions, binding of DNA to a silica membrane and ethanol washing in spin column format and finally elution of purified DNA.
The densities of Streptomyces in DNA extracted from sediment samples were determined by qPCR based on amplification of the 16S rRNA gene using the genus-specific primers described in Chapter 4. Triplicate qPCR reactions were performed using a LC96 (Roche
Diagnostics) in a volume of 10 µL. The chemical composition and thermal cycling conditions of the qPCR reactions were identical to those described previously (section 4.2.2). CT values were converted to cells using the calibration curves presented in section 4.3.2 which were expressed on a per dry weight basis of sediment (cells/g dw).
5.2.6 Statistical analyses
One way analysis of variance (ANOVA) followed by a Tukey’s Post Hoc test was carried out to test for significant differences in geosmin and 2-MIB produced between the different sediment treatments (A, B and C) under each condition (exposed and submerged) in addition to overlying water concentrations. Student’s t-tests were performed to test for significant differences in geosmin and 2-MIB yields between exposed and submerged conditions for each sediment treatment (A, B and C). The same statistical test was also performed to determine significant differences in Streptomyces abundance including between each condition for each sediment type and to test for significant differences in measured
Streptomyces population densities on day 0 and 7 for each sediment treatment (A, B and C) under each experimental condition (exposed, submerged or submerged anaerobic). ANOVAs and t-tests were carried out using JMP version 7.0 with a confidence level set at 95%.
Regression analyses were conducted using Microsoft Office Excel 2013 to examine the 165
relationship between submerged sediment and overlying water concentration of geosmin and
2-MIB.
5.3 Results
The data pertaining to the significance of marginal reservoir sediment as a site of geosmin and 2-MIB production by Streptomyces are presented in the following results. Analyses include a comparison of geosmin and 2-MIB levels measured in sediment under exposed and submerged conditions, simulating water level recession, hypothesised to provide the wet aerobic conditions ideal for these bacteria to flourish and become a significant source of T&O compounds in marginal sediments. For comparative purposes, permanently submerged, anaerobic conditions representative of bottom reservoir sediments where Streptomyces are considered to be relatively inactive and thus do not produce T&O compounds, were also simulated. The data relating to changes in qPCR-measured population densities of
Streptomyces in the sediment of these experiments is also presented.
5.3.1 Geosmin and 2-MIB production in exposed and submerged sediment
It was hypothesised that when marginal sediments are exposed to air, representative of prolonged dry periods in reservoirs when water level recession occurs, geosmin and 2-MIB production by Streptomyces would be enhanced compared to when these shallow sediments are submerged. Contrary to the hypothesis, no significant differences in the levels of either compounds were observed between exposed and submerged conditions for any of the sediment treatments (Figure 5.3). There were however, differences in the recovered levels of these compounds between sediment treatments under the experimental conditions, with sterilised sediment inoculated with S. coelicolor A3(2) (Sediment A, blue bars) exhibiting the highest level of geosmin in exposed and submerged sediment (173.4 ng/kg dw and 291.1 166
ng/kg dw respectively) compared to inoculated non-sterile sediment (Sediment B, red bars,
64.8 ng/kg dw and 64.5 ng/kg dw respectively) and the biotic control (Sediment C, green bars, 96.5 ng/kg dw and 67.9 ng/kg dw respectively). There were correspondingly significantly greater concentrations detected in the overlying water of the submerged sterile- sediment inoculated with S. coelicolor A3(2) (2,121.5 ng/L) compared to non-sterile sediment and the biotic control (237.2 ng/L and 241.9 ng/L respectively).
Figure 5.3 Levels of geosmin (top) and 2-MIB (bottom) in sediment (ng/kg dw) and overlying water (ng/L) for all sediment treatments examined. Values represent the mean ± standard error (n=3). No detectable quantities of either compound were measured in the sediment or overlying water of sediment A incubated under anaerobic conditions. Values not connected by the same letter for each sediment condition (exposed and submerged) or overlying water are significantly different (p<0.05). Significant differences (p<0.05) between exposed and submerged conditions for each sediment treatment were not identified. 167
The observed concentrations of 2-MIB followed a similar trend, with the S. coelicolor A3(2) in sterilised sediment treatment exhibiting signficantly greater amounts under exposed conditions (291.5 ng/kg dw) compared to inoculated non-sterilised sediment (67.5 ng/kg dw), but did not differ signficantly to the biotic control (227.9 ng/kg dw). As oberved for geosmin levels, no signficant differences between 2-MIB detected under exposed and submerged conditions were found for any of the sediment treatments. Although S. coelicolor A3(2) in sterilised submerged sediment displayed the highest yield of 2-MIB (260.4 ng/kg dw), this was not signficantly greater compared to either inoculated non-sterilised sediment or the biotic control (157.5 ng/kg dw and 110.2 ng/kg dw). There was however, signicantly more
2-MIB detected in the overlying water (1,427.3 ng/L) of the former treatment compared to inoculated non-sterilised sediment and the bitoic control (316.9 ng/L and 284.9 ng/L respectively). Strong correlations were identified between the levels of each compound in submerged sediment and quantities detected in the overlying water as shown in Figure 5.4, with R2 values of 0.999 and 0.919 for geosmin and 2-MIB respectively. For the sterilised submerged sediment inoculated with S. coelicolor A3(2) and incubated under anaerobic conditions, geosmin or 2-MIB could not be detected in the sediment nor in the overlying water.
2500
2000 R² = 0.999 1500 Geosmin 1000 R² = 0.919
Overlyingwater 500 2-MIB concentration(ng/L)
0 0 50 100 150 200 250 300 350 Submerged sediment concentration (ng/kg dw) Figure 5.4 Relationship between geosmin and 2-MIB levels in the submerged sediment and overlying water based on mean values (n=3) from the three different sediment treatments (A [blue], B[red] and C[green]). 168
5.3.2 Dynamics of Streptomyces population densities in exposed and submerged sediment
In addition to measuring the concentrations of geosmin and 2-MIB, Streptomyces population densities in the sediments were determined. Figure 5.5 shows abundances on day 0 and after the 7 day incubation period for each sediment treatment (A, B and C) and experimental condition (exposed, submerged and submerged anaerobic). For sterilised sediment inoculated with S. coelicolor A3(2) (Sediment A, blue columns), no significant changes occurred between day 0 and 7 under submerged conditions (1,588,887 cells/g dw and 1,623,485 cells/g dw respectively), compared to exposed conditions, where a signficant increase in abundance by over two-fold was detected (1,743,697 cells/g dw and 3,919,388 cells/g dw respectively).
These discrepancies in population dynamics contrast with the comparable quantities of geosmin and 2-MIB detected between exposed and submerged conditions in this sediment treatment. By comparison, Streptomyces cell densities in this sediment subjected to submerged anaerobic conditions significantly decreased by more than three-fold during the incubation period (from 1,604,141 cells/g dw to 496,280 cells/g dw), aligning with the absence of geosmin and 2-MIB. Thus overall, significantly greater Streptomyces population densities were observed in the exposed compared to submerged sterile sediment, and under aerobic compared to anaerobic conditions.
For non-sterilised sediment inoculated with this strain (Sediment B, red columns) under both exposed and submerged conditions, significant decreases in abundance by approximately two-fold were detected after 7 days of incubation (from 1,801,898 cells/g dw to 768,885 cells/g dw and from 1,966,039 cells/g dw to 988,858 cells/g dw respectively). This is consistent with the observation of lower concentrations of both geosmin and 2-MIB compared to the sterilised sediment and additionally, with the comparable yields of these metabolites between exposed and submerged conditions. Sediment Streptomyces cell densities in the biotic control (Sediment C, green columns) in contrast, remained 169
Figure 5.5 Mean Streptomyces cell densities (cells/g dw) measured in each sediment treatment (A, B and C) under each experimental condition (exposed, submerged or submerged anaerobic) on day 0 and day 7. The standard error based on triplicate samples and triplicate qPCR results are shown (n=9). Asterisks indicate significantly greater (p<0.05) abundance between day 0 and 7 for each sediment treatment under each experimental condition. Significant differences (p<0.05) in Streptomyces abundance after 7 days for each sediment treatment (A, B and C) between different conditions are shown by letters or numbers (i.e. those not connected by the same letter or number are significantly different).
comparatively constant and similar under both exposed and submerged conditions (changing from 981,800 cells/g dw to 1,000,899 cells/g dw and from 1,029,654 cells/g dw to 1,168,246 cells/g dw respectively). This observation aligns with the statistically similar yields of geosmin and 2-MIB detected between exposed and submerged conditions.
5.3.3 Dissolved oxygen concentration at the surface of submerged sediment
Dissolved oxygen (DO) concentrations were measured above the sediment surface for all treatments under submerged conditions. As shown in Table 5.1, aerobic conditions prevailed.
Slight differences were observed between sediment treatments, being greatest in the sterile
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sediment inoculated with S. coelicolorA3 (2), followed by inoculated non-sterilised sediment and the biotic control. At 25°C, the temperature of incubation, the maximum amount (i.e. saturation) of oxygen that can be dissolved in water is 8.24 mg/L, and thus the sediment surface oxygen concentrations measured were approximately half of the oxygen saturation.
Minute levels of DO were measured above the sediment surface incubated under anaerobic conditions. Although this measurement was taken immediately following removal from the anaerobic jar, detection of low DO levels probably reflects the onset of a rapid equilibration in oxygen concentration between the atmosphere and oxygen levels above the sediment.
Table 5.1 Dissolved oxygen levels (mg/L) measured above the sediment surface for sediment treatments subjected to submerged conditions (n.d. = no data). Mean values ± standard error are presented (n=3). Sediment condition Sediment treatment Aerobic (mg/L) Anaerobic (mg/L) A. Sterilised sediment + S. coelicolor A3(2) 4.63±0.10 0.27±0.03 B. Non-sterilised sediment + S. coelicolor A3(2) 3.97±0.32 n.d. C. Non-sterilised sediment, biotic control 3.20±0.24 n.d.
5.4 Discussion
5.4.1 Geosmin and 2-MIB production by Streptomyces in marginal sediments: a comparison between exposed and submerged conditions
The focus of this study was the possibility that Streptomyces inhabiting the marginal sediments of reservoirs represent a significant source of geosmin and 2-MIB that can contribute to T&O episodes in drinking water. Several researchers have proposed that the aerobic conditions encountered in the wet-dry regime of this marginal habitat are favourable for the growth of Streptomyces (Romano & Safferman, 1963; Wood et al., 1983a, 1985;
Bailey, 1988; Wnorowski, 1992). Specifically, it was hypothesised that when these sediments
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are exposed, representing extended dry periods when water levels recede, the moist aerobic conditions would stimulate the proliferation of Streptomyces and their production of earthy- musty secondary metabolites. Furthermore, the submersion of these marginal sediments, representing a high water level, was hypothesised to be comparatively less favourable for the growth and odour production by these aerobic bacteria due to the lower oxygen availability.
Contrary to the hypothesis, higher levels of geosmin and 2-MIB in each of the sediment treatments in exposed as opposed to submerged conditions was not observed. Previous studies have suggested that Streptomyces inhabiting marginal sediments covered by water are relatively inactive due to lower oxygen levels and that aerobic conditions required for production of geosmin and 2-MIB prevail only when the sediment is exposed during a drop in the water level. The sediments could then become a source of these odorous secondary metabolites when the water level rises or from wash-in during runoff events. Wood et al.
(1985) observed that reservoir sediment exposed during maintenance work developed an earthy odour, containing a geosmin concentration of 46 µg/kg dry weight, whereas none was detected in unexposed sediment, which rather, smelled of hydrogen sulphide, indicative of the prevalence of anaerobic conditions.
However, the DO concentrations measured at the sediment-water interface in the present study confirmed the presence of low concentrations of oxygen (3.20-4.63 mg/L), which may have been sufficient for Streptomyces to produce comparable quantities to those detected in sediment exposed to atmospheric oxygen levels. Silvey and Roach (1975) stated that the minimum DO requirement for the earthy-musty metabolite producing secondary mycelium to grow in water was 1.8 mg/L. Nevertheless, the exact relationship between oxygen availability and production of these compounds remains unclear. Several studies have
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demonstrated that higher oxygen concentrations appear to stimulate greater geosmin production by Streptomyces cultures. Dionigi and Ingram (1994) reported that oxygen enrichment (30% O2 atmosphere) increased production of geosmin in solid cultures of S. tendae compared to ambient atmosphere incubated cultures (21% O2) and that oxygen depletion (5% and 10% O2 atmosphere) decreased the production of geosmin. Schrader and
Blevins (1999) also reported that an oxygen reduced atmosphere decreased geosmin production by liquid cultures of S. halstedii compared to ambient conditions. In contrast with these findings, Sunesson et al. (1997) reported that decreasing the atmospheric oxygen concentration from 16% to 8% increased geosmin production in solid cultures of S. albidoflavus. Thus the findings of equivalent production of geosmin and 2-MIB in exposed and submerged sediments in this study appears to add little in the way of clarity regarding the importance of oxygen concentration to the occurrence of T&O episodes involving
Streptomyces.
One explanation for the detection of lower than expected quantities of these compounds in exposed sediment may be loss to the atmosphere. Being volatile compounds, the high incubation temperature utilised may have encouraged some loss to the atmosphere by volatilisation, as reported by Guttman and van Rijn (2012). Indeed, a high concentration was measured in the water overlying the submerged sediment cultures, which were correlated with the amount produced for each sediment treatment (Figure 5.4). This clearly suggested that geosmin and 2-MIB were being released into the overlying water. Zuo et al. (2010) demonstrated that up to 51.4% of geosmin produced in reservoir sediment was released into the overlying water and that geosmin in the water could be correlated with concentrations in the sediment. Thus loss of these odorous metabolites from the sediment clearly occurs and is likely to occur at a faster rate when exposed to the atmosphere. It must also be acknowledged that the geosmin and 2-MIB concentrations measured in the sediment are probably an 173
underestimate as these were determined by LLE of the interstitial water in the sediment. As they are hydrophobic, it can be assumed that some fraction of these compounds remained strongly adsorbed to sediment surfaces. This was observed by Stahl and Parkin (1994) when using a purge-and-trap technique for analysing geosmin and 2-MIB in soil placed in water.
Low recovery efficiencies (<24%) were reported for added quantities of these compounds due to their sorption to soil. Water on the contrary, is an easier substrate from which to extract these compounds due to their hydrophobic properties (thus low solubility).
Despite finding no significant differences between exposed and submerged sediment yields of geosmin and 2-MIB, the data confirm that both shallow and exposed sediments located at the margins represent a significant source of these compounds produced by Streptomyces.
Given that up to nearly 300 ng/kg of each compound was produced (sterilised sediment inoculated with S. coelicolor A3(2)), which probably represents an underestimate, such sediment washed into the adjacent water mass could produce detectable odours.
Concentrations detected in overlying water (240 ng/L to 2,120 ng/L) would be extremely unpalatable and reaffirms that complete exposure of shallow sediments is certainly not essential for Streptomyces to produce these compounds.
Despite no evidence of greater odour production, the Streptomyces population density did increase (two-fold) in exposed sterilised sediment inoculated with S. coelicolor A3(2) compared to no change under submerged conditions, suggesting that exposure and consequently greater aerobic conditions to air stimulates the growth of these bacteria in sediment. This aligns with the findings in Chapter 4, in which Streptomyces abundance in marginal substrates was found to be significantly greater, particularly at Chichester
Reservoir, during the dry sampling period when the water level had dropped by over 2.0 m,
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exposing the marginal sediments and soils. The Streptomyces population density in sediment from the south marginal site (used for the experimental work in this study) was found to have increased by 18.5 times under exposed compared to submerged conditions. Similarly,
Streptomyces abundance in sediment from the north marginal site when exposed was 23.6 times greater compared to submerged conditions. Furthermore, significant increases in
Streptomyces populations were detected in plant debris and soil (up to 100-fold) at these marginal sampling sites during dry conditions. However, in this laboratory study, differences in population densities between exposed and submerged conditions were not observed for non-sterilised sediment inoculated with S. coelicolor A3(2) or the biotic control. Possible reasons for these discrepancies are further discussed.
Sterilised reservoir sediment inoculated with S. coelicolor A3(2) returned significantly higher geosmin and 2-MIB yields (more than 2-fold greater) under both exposed and submerged conditions compared to the inoculated non-sterilised sediment (with the exception of 2-MIB produced under submerged conditions not being significantly greater). These observations corroborate those reported by Romano and Safferman (1963) and Persson and
Sivonen (1979). To investigate the significance of introduction of odour compounds from a soil origin into a water environment, Romano and Safferman (1963) percolated water continuously through soil columns inoculated with a Streptomyces sp. (S. griseoluteus). The authors identified that considerable quantities of the earthy-musty odour were readily washed from the soil, with significantly more from sterilised soil compared to non-sterilised soil.
Persson and Sivonen (1979) inoculated a Streptomyces isolate into sterilised and unsterilised brackish reservoir water, observing that the abundance of the bacteria declined and the earthy odour disappeared after a few days in the non-sterile water as opposed to the sterilised sample, in which case growth was profuse and the odours strong throughout the experiment.
The disappearance of the odour in the non-sterile water was attributed to the decrease in the 175
Streptomyces population. Similarly in this study, the Streptomyces cell densities determined using qPCR revealed a significant reduction in abundance over the incubation period in non- sterilised sediment inoculated with S. coelicolor A3(2). In comparison, the Streptomyces abundance increased over two-fold in exposed sterilised sediment while the population remained comparatively constant throughout the experiment under submerged conditions.
These observations probably account for the significantly greater levels of geosmin and 2-
MIB obtained in this sediment treatment. As suggested by Persson and Sivonen (1979), the indigenous microorganisms in the non-sterilised sediment may have proliferated during the incubation period and reduced the growth of the introduced Streptomyces strain. It is also feasible to suggest that the native microorganisms in the non-sterilised sediment may have degraded some of the geosmin and 2-MIB produced by S. coelicolor A3(2). A variety of common bacterial genera have been implicated in the biodegradation of these metabolites including Bacillus spp., Candida spp. Enterobacter spp. Flavobacterium spp., Pseudomonas spp. and Rhodococcus spp. (Ho et al., 2012).
The observation of similar quantities in the non-sterilised sediment without the Streptomyces inoculum (the biotic control) suggests that the autochthonous microbial inhabitants may have been capable of producing geosmin and 2-MIB. Indeed, based on the detection of high abundance of Streptomyces, which remained relatively stable throughout the incubation period, these bacteria were likely to be the producers of the earthy-musty odour. Furthermore, several of the Streptomyces that were identified in Chapter 4 and confirmed as geosmin and
2-MIB produces were isolated from this particular sediment. It is however important to acknowledge that other organisms present in the sediment (e.g. fungi) may have also contributed to this.
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5.4.2 Geosmin and 2-MIB production by Streptomyces under anaerobic conditions in submerged sediments
In this study, it was also hypothesised that under anaerobic conditions in submerged sediments, conceivably representing the situation encountered in permanently submerged deep sediments and muds at the bottom of reservoirs, growth and production of geosmin and
2-MIB by these aerobic bacteria would be inhibited. The finding that sterilised sediment inoculated with S. coelicolor A3(2) and incubated under submerged anaerobic conditions did not produce detectable amounts of geosmin or 2-MIB supports the hypothesis. Furthermore, a greater than three-fold reduction in the Streptomyces population density had occurred by the end of the 7 day incubation period, compared to a two-fold increase and no change in the same sediment treatment subjected to exposed and submerged aerobic conditions respectively. These findings certainly lend support to the notion that Streptomyces are aerobic bacteria and require the presence of oxygen for growth and the production of geosmin and 2-
MIB in the sediment environment. However, Higgins and Silvey (1966) reported that when the culture system was anaerobic, spore germination and the development of the primary mycelium could occur in Streptomyces, although growth of the secondary mycelium was inhibited, in which case there was no odour production. The authors however noted that when anaerobic conditions were applied late in the development of the primary mucelium, the secondary mycelium could occasionally grow as a facultative anaerobe.
Several researchers contend that Streptomyces spp. such as S. coelicolor A3(2) may be capable of growth under anaerobic conditions. Indeed, the complex nature of the soil environment means that Streptomyces have had to evolve a broad range of metabolic pathways to enable them to survive in this environment, where extremely variable oxygen tensions are experiened, with little to no oxygen in wet soil. The genome of S. coelicolor
A3(2) contains several enzymes that, in facultative or strict anaerobes, confer the ability to 177
grow by anaerobic respiration. S. coelicolor A3(2) has three respiratory nitrate reductase gene clusters, all of which are expessed based on transript analysis (van Keulen et al., 2005a). This strain has been shown to survive long periods of anaerobiosis (both in a spore and hyphal state) by mainintaing metabolic activity. This survival strategy has not been adopted by all
Streptomyces such as S. avermitilis, which lacks the three nitrate reductase operons (van
Keulen et al., 2007). The capacity to respire with nitrate (use as an electron acceptor) may give these bacteria a selective advantage in the soil environment by delivering enough energy to maintain the membrane potential (proton moptive force) or to sustain growth when oxygen is depleted (Fischer et al., 2010).
Guttman and van Rijn (2008) observed growth and production of geosmin and 2-MIB by two
Streptomyces spp. isolated from a recirculating aquaculture system (RAS) under both anoxic and aerobic culture conditions (although to a significantly lesser extent under anoxic conditions). Under anoxic conditions (established by bubbling argon gas into airtight flasks), a decrease in the medium nitrate concentration (and only slight increase in nitrite and ammonia) indicated a denitrification ability of the bacteria. The authors suggested that the capacity of Streptomyces to generate energy by using nitrate as an electron acceptor in the absence of oxygen may account for the occurrence of geosmin and 2-MIB in anaerobic organic-rich digestion basin of the RAS.
Given these findings, it is plausible to consider that small quantities of T&O metabolites below the analytical detection limits may have been produced in the sediment by S. coelicolor
A3(2) under anaerobic conditions. Biodegradation of these compounds in the sterilised sediment would not be expected to account for their loss due to destruction of the indigenous microorganisms during the autoclaving process. However, Nowak and Wronkowska (1987)
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determined soil sterilisation by autoclaving to be incomplete as bacteria and fungi could be isolated from autoclaved soil, including Bacillus spp. and fungi. Ramsay and Bawden (1983) managed to destroy only 55% of bacteria by sterilizing 100 g of soil samples at 121°C for 30 min. Resistance of endospores from bacterial genera such as Bacillus (a known geosmin and
2-MIB degrader) to high temperature must be acknowledged and the protective effects of the sediment in the large mass used for autoclaving. Khafif et al. (1983) found that bacteria with no property of that kind (e.g. Pseudomonas spp.) could survive in the soil after autoclaving at 121°C for 20 minutes three times. Thus, surviving anaerobic or facultative anaerobic bacteria may have contributed to biodegradation of geosmin and 2-MIB under anaerobic conditions. Furthermore, Guttman and van Rijn (2012) reported that several geosmin and 2-
MIB biodegrading bacteria (e.g. Rhodococcus, Variovorax and Comamonas spp.) isolated from an anaerobic digestion basin in an aquaculture facility could biodegrade these compounds (as the sole carbon source) under anoxic conditions, although at a slower rate than in aerobic conditions.
Overall, the findings indicate that geosmin and 2-MIB are not produced by S. coelicolor
A3(2) under anaerobic conditions. However based on the literature reports discussed above, it seems plausible that they may be produced to some extent during oxygen deprivation, representing conditions similar to those encountered in the bottom sediments of reservoirs.
In Grahamstown and Chichester Reservoirs, measurements of DO recorded weekly at the bottom of the storages average 8.3 mg/L (range 1.8 mg/L to 13.3 mg/L) and 7.7 mg/L (range
0.6 mg/L to 12.2 mg/L) respectively. In this study, production of geosmin and 2-MIB by
Streptomyces in sediment occurred under submerged conditions, where the DO above the sediment was lower (3.20 mg/L to 4.63 mg/L) than those typically encountered in the bottom waters of the reservoirs. Thus it is feasible to suggest that these compounds may be produced by Streptomyces populations inhabiting the bottom substrates of the reservoirs due to 179
sufficient oxygen availability. In Chapter 4, relatively stable populations were indeed detected in the bottom sediments of Chichester and Grahamstown Reservoirs, which were comparatively less numerous with lower proportions of vegetative cells compared to marginal substrates. Several studies have isolated geosmin and 2-MIB producing actinobacteria notably Streptomyces, from the bottom sediments of water bodies which have been considered as a potential source of geosmin and 2-MIB in water (Willoughby, 1969;
Johnston & Cross, 1976a; Sugiura & Nakano, 2000; Zaitlin et al., 2003b; Zuo et al., 2009a).
Large quantities of geosmin (up to 5,280 ng/kg dw) in the bottom sediments of the Xionghe
Reservoir (China) were attributed to Streptomyces on the basis of geosmin and 2-MIB production ability of isolates (Zuo et al., 2010). Sugiura and Nakano (1987) also concluded that the elevated 2-MIB levels in Lake Kasumigaura (Japan) were generated by the activity of actinobacteria in the aerobic, carbohydrate rich sediment layers, based on their high numbers and in vitro observations of geosmin and 2-MIB production by isolates from this habitat.
5.5 Conclusions
The results from this laboratory study indicate that sediments located at the margins of reservoirs represent a site of potentially significant production of geosmin and 2-MIB by
Streptomyces, which occurs irrespective of submerged or exposed conditions. The hypothesis that exposure of marginal sediments, reflecting the situation of water level recession, leads to greater production of these compounds by Streptomyces was not supported. While it was demonstrated that anaerobic conditions do preclude the growth and ability of Streptomyces to produce these compounds in sediments, low oxygen levels in submerged conditions were sufficient for production of comparable quantities to those produced in sediment exposed to the atmosphere. The release of these compounds from submerged sediment into overlying
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water indicates that Streptomyces in shallow sediments represent a significant and possible continuous source of geosmin and 2-MIB that can contribute to T&O episodes. These bacteria inhabiting shallow sediments may be responsible for background concentrations of these compounds found in Chichester Reservoir, particularly when blooms of Anabaena are absent.
The results imply that management of storage levels to avoid exposure of sediments at the margins of reservoirs may not be a necessary measure to prevent increased geosmin and 2-
MIB production by Streptomyces populations inhabiting such substrates. This contradicts the observations of HWC and several other authors concerning earthy-musty odour problems that have occurred after extended warm, dry periods in which exposure of shallow margins occurs followed by refilling and re-submerging during rain events (Bailey, 1988;
Wnorowski, 1992; Uwins et al., 2007). However, considering the evidence for significantly greater growth of S. coelicolor A3(2) under exposed conditions compared to submerged conditions in sediment, it is still conceivable that higher abundances in exposed sediment would translate into significantly higher levels of T&O compounds. Interpretation of the results warrants caution due the absence of measurement of the probable loss of these volatile compounds from exposed sediment and the underestimation of their concentration in sediment. It is recommended that future studies employ more rigorous means to extract these compounds from solid matrices and measure their loss to the atmosphere, and repeat these experiments using other growth substrates found at the margins of reservoirs.
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CHAPTER 6 -MORPHOLOGICAL DIFFERENTIATION AND THE INFLUENCE OF ENVIRONMENTAL FACTORS ON GEOSMIN AND 2-METHYLISOBORNEOL PRODUCTION BY STREPTOMYCES SPP.
6.1 Introduction
Apparent among the existing literature is the distinct paucity of knowledge regarding the ecology of Streptomyces in freshwater environments and their potential role in contributing to the development of T&O episodes involving geosmin and 2-MIB in drinking water supplies (Zaitlin & Watson, 2006; Juttner & Watson, 2007). Thus far, the research presented in this thesis has provided evidence to suggest that Streptomyces in the reservoirs studied are widely distributed, abundant and active and have the capacity to produce T&O causing compounds. These research findings support a terrestrial origin of these bacteria and their odorous secondary metabolites in the water mass, as the primary means by which they may contribute to T&O episodes. The production of geosmin and 2-MIB by these bacteria in the marginal soils, sediment and plant debris surrounding drinking water reservoirs (particularly when exposed to air), aligns with their ecological importance as aerobic saprophytes, thriving ubiquitously on solid substrates (Goodfellow & Williams, 1983). Their existence as metabolically active bacteria in the water mass has, on the contrary, been widely disputed
(Johnston & Cross, 1976a; Cross, 1981; Jensen et al., 1994; Zaitlin et al., 2003b). However, the detection of their vegetative cells in the water mass and their growth and geosmin and 2-
MIB production in reservoir water in the laboratory suggests that Streptomyces may be capable of directly contributing to T&O episodes through in situ production of these secondary metabolites in the water mass.
Having established the potentiality of Streptomyces to contribute to T&O episodes, it was considered that the logical progression to further develop this research was to better
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understand physico-chemical factors, representative of their environment, which may trigger their production of geosmin and 2-MIB. This forms the central focus of experimental work presented in the current chapter.
The effects of several physico-chemical factors on geosmin and 2-MIB biosynthesis by
Streptomyces have been examined here with the objective to better define the role of these bacteria in T&O episodes and to determine the conditions favourable for production of these secondary metabolites. Chapter 2 (section 2.4.2.1) provided a review of previous studies which have examined the effects of major nutrients such as carbon source and phosphorous, nitrogen, potassium and calcium concentration (Weete et al., 1977; Blevins et al., 1995;
Schrader & Blevins, 2001; Uwins, 2011), concentration of micronutrients (Schrader &
Blevins, 2001), changes in temperature (Weete et al., 1977; Dionigi & Ingram, 1994; Blevins et al., 1995; Uwins, 2011), pH (Weete et al., 1977; Dionigi & Ingram, 1994; Blevins et al.,
1995) and salinity levels (Rezanka & Votruba, 1998) on geosmin production by
Streptomyces, with some of these factors demonstrating either a stimulatory or inhibitory effect. Prior to this study, the effects of such factors on 2-MIB production have remained largely unexplored.
The intent of the laboratory-based study presented in this chapter was to gain better understanding of the extent to which changes in certain environmental parameters can influence the production of geosmin and 2-MIB by Streptomyces spp. The specific physico- chemical conditions applied in the study were those previously reported in the literature to have apparently influenced geosmin production by this genus. These included changes in temperature, pH and salinity levels, and the concentration of macronutrients (carbon, nitrogen, phosphorous, potassium, magnesium and calcium) and micronutrients (iron, zinc,
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copper and manganese). Previously published laboratory-based studies regarding the influence of such variables on geosmin production by Streptomyces have used the conventional ‘one-factor-at-a-time’ experimental approach. This experimental methodology can be time-consuming, inefficient and potentially misleading when it comes to discovering new knowledge through experimentation because the possible effects of interactions between the variables are not taken into account (Lawson, 2010). A much better strategy for experimenting with multiple factors is to use a design of experiments (DOE) method referred to as full factorial design. These form the basis for all classical experimental designs used in screening, optimisation and robustness testing (Eriksson et al., 2008). The use of an orthogonal multivariate design allows independent estimates of the effects of changes in the levels of each experimental variable, as well as the effects of interactions of several factors on the response variable. This study is unique in its application of a two-level full factorial design in order to determine simultaneously, the influence of several environmental variables on geosmin and 2-MIB production by Streptomyces spp.
Streptomyces exhibit a complex developmental cycle with physiologically distinct stages starting from spore germination, vegetative hyphal growth, the production of aerial
(secondary) hyphae followed by its differentiation (segmentation) to form chains of spores which are then released for dispersal in the environment (Goodfellow & Williams, 1983;
Williams et al., 1984). As with many other secondary metabolites, numerous reports have identified a relationship between the production of geosmin and degree of morphological differentiation, with the highest levels of production generally associated with cultures of
Streptomyces in the reproductive development stage (aerial hyphae and sporulation) (Bentley
& Meganathan, 1981; Dionigi et al., 1992; Scholler et al., 2002; Tung et al., 2006; Yague et al., 2013). Thus in addition to examining the effect of changes in environmental parameters on geosmin and 2-MIB production by Streptomyces spp., a key objective of this study was 184
to provide a quantitative assessment of the apparent association between production of these compounds and morphological differentiation of the organisms. Based on the literature evidence and the notion that a shift to reproductive activity and dispersal (sporulation) commonly occur when conditions become unfavourable for continued vegetative growth
(Bibb, 2005; van Keulen et al., 2011), two primary hypotheses were proposed:
1. Cultures of Streptomyces spp. exhibiting higher degrees of morphological
differentiation (sporulation) would produce significantly greater yields of geosmin
and 2-MIB than those at the primary vegetative growth stage.
2. Production of geosmin and 2-MIB would be greatest under conditions of major
nutrient (carbon, nitrogen and phosphorous) limitation.
Other factors considered likely to inhibit vegetative growth and therefore be associated with enhanced production of geosmin and 2-MIB included:
Low concentration of other macronutrients (calcium, potassium and magnesium)
Increasing NaCl concentration
Elevated pH (beyond the relatively neutral range of pH 6.5-8)
Temperature variation was also included as a factor for examination, although variations were restricted to within a fairly moderate range and it was unclear based on literature evidence in which direction, if at all, these variations would affect Streptomyces in relation to production of geosmin and 2-MIB.
Due to the number of factors to be examined overall, the study was divided into four separate experiments, each designed to assess the interactive effects of at least three factors on T&O metabolite production by the Streptomyces cultures. Within this scheme, the results of the initial experiments were used to guide the design of the subsequent experiments toward
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conditions favourable to T&O production, effectively pushing the system further in this direction with each successive experiment. While this design has the potential to mask relatively subtle effects in successive experiments, it also adds considerable weight to any statistically significant effects observed. The final experiment involved variation in micronutrients (zinc, iron, manganese and copper) concentration under these optimised conditions and as was the case regarding temperature variation, it was not clear in advance what effects if any could be expected to emerge.
6.2 Materials and methods
6.2.1 Microorganisms
Streptomyces coelicolor A(3)2, the model organism for antibiotic production of the genus, was selected for use in this study (Bentley et al., 2002). The biosynthesis of geosmin has been extensively characterised in this organism (Cane & Watt, 2003; Gust et al., 2003). In addition, the experiments were repeated using a Streptomyces sp. isolated from the marginal sediments of Chichester Reservoir which, based on 16S rRNA sequencing, was tentatively identified as Streptomyces antibioticus. This isolate was chosen as it produced relatively high and equivalent amounts of both geosmin and 2-MIB on SCA (Chapter 4, section 4.3.5).
Conducting the experiments with two Streptomyces spp. was considered important in order to identify differences and similarities in the responses of each to facilitate identification of significant variables influencing geosmin and 2-MIB production across this genus. Cultures used for each experiment were maintained on SCA and repeatedly sub-cultured to ensure culture purity. Spores of both species were placed in 20% glycerol solutions for long term storage at -20°C.
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6.2.2 Full factorial experimental design
Cultures of S. coelicolor A3(2) and S. antibioticus on solid media were exposed to a range of environmental conditions according to full factorial experimental design with two-levels.
Such an experimental design consists of two or more independent factors, each with two levels or discrete values (a high level often given by the ‘+’ symbol and a low level denoted by a ‘-’ symbol) and whose experimental units take on all possible combinations of these levels across all such factors (an orthogonal array of experiments). The design determines the effect of each factor on each response variable in addition to how the effect of each factor varies with the change in the level of the other factors i.e. interactions (synergies and antagonisms). Full factorial two level experiments are referred to as 2k designs, where k denotes the number of factors to be investigated. A full factorial design with k factors requires
2k runs for a single replicate. Thus an experiment with three factors would require 23 (8) treatment combinations and an experiment with four factors would require 24 or (16) treatment combinations. The statistical model of a two-level factorial design is linear. The complete MLR (multiple linear regression) model equation (Equation 1) for 3 factors (x1, x2 and x3) with interaction is given by the second order interaction (polynomial) model:
Y = b0 + b1 x1 + b2 x2 + b3 x3 + b12 x1 x2 + b13 x1 x3 + b23 x2 x3 + residual error (Equation 1)
where Y is the predicted response, the coefficients b1, b2 and b3 account for the main
(independent) effects (the linear coefficients) of the factors x1, x2 and x3 respectively, while b12, b13 and b23 represent the three second order interaction terms between the factors. The independent term b0 represents the the mean of all experiments (i.e. the constant coefficient).
The error is the residual response variation not explained in the model. The aim of the data analysis is to estimate numerical values of the model parameters (the regression coefficients)
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which indicate how the factors and interactions between factors influence the response
(Lundstedt et al., 1998; Morris, 2011).
In this study, three 23 and one 24 independent experiments were conducted in order to investigate the effect of selected environmental variables on geosmin and 2-MIB production by two Streptomyces spp. The two-level full factorial designs were generated using MODDE software version 9.0 (Umetrics, Umeå, Sweden). These experimental designs were conducted as follows:
Experiment 1 (23 factorial design): effects of macronutrient concentration including
carbon (0.42 mM, 208.5 mM and 416.6 mM), nitrogen (3.57 µM, 894.20 µM and
1,784.82 µM) and phosphorous (1.61 µM, 404.42 µM and 807.23 µM).
Experiment 2 (23 factorial design): effects of temperature (24°C, 27°C and 30°C),
pH (6.0, 7.5 and 9.0) and NaCl concentration (0%, 1% and 2%).
Experiment 3 (23 factorial design): effects of macronutrient concentration including
calcium, potassium and magnesium (1 µM, 500.5 µM and 1000 µM)
Experiment 4 (24 factorial design): effects of micronutrient concentration including
iron, zinc, copper and manganese (0.25 µM, 12.51 µM and 25 µM)
The factors were set at two widely spaced levels (high and low), with values generally selected based on previous studies and representing conditions potentially encountered in reservoirs. The experimental design matrices and geometric representation of each factorial experiment used for screening the influence of the environmental parameters on geosmin and
2-MIB production are shown in Figures 6.1, 6.2, 6.3 and 6.4. Two-level factorial experiments are often depicted ‘spatially’ by representing treatments as corners of a cube, where each spatial dimension is associated with a factor. Each experiment was run separately in the order they are listed above and all combinations of factors within each experiment were performed
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simultaneously. For Experiments 1 to 3, a total of 33 experiments were performed according to a three-factor two level full factorial design with eight corner experiments performed in triplicate and nine replicates of the centre point (centroid) where all factors were set to their mid value. For Experiment 4 which involved four factors, 57 experiments were carried out including 16 triplicate corner experiments and nine replicates of the centroid. The centroid,
(often detonated as the ‘0’ level) allows for the detection of non-linear effects between the variables and responses. The level of the independent variables in each experiment that optimised geosmin and 2-MIB production were applied in experiments that followed, which examined the effects of other factors.
Figure 6.1 The two-level full factorial experimental design matrix for Experiment 1 showing the variation in three environmental parameters to investigate the independent and synergistic effects of changes in carbon, nitrogen and phosphorous concentration on geosmin, 2-MIB and biomass production by Streptomyces spp. The design included a mid-point condition (orange) and combinations of the variable levels as represented by the corners of the cube.
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Figure 6.2 The two-level full factorial experimental design matrix for Experiment 2 showing the variation in three environmental parameters to investigate the independent and synergistic effects of changes in temperature, pH and NaCl concentration on geosmin, 2-MIB and biomass production by Streptomyces spp. The design included a mid-point condition (orange) and combinations of the variable levels as represented by the corners of the cube.
Figure 6.3 The two-level full factorial experimental design matrix for Experiment 3 showing the variation in three environmental parameters to investigate the independent and synergistic effects of changes in calcium, magnesium and potassium concentration on geosmin, 2-MIB and biomass production by Streptomyces spp. The design included a mid-point condition (orange) and combinations of the variable levels as represented by the corners of the cube.
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Figure 6.4 The two-level full factorial experimental design matrix for Experiment 4 showing the variation in four environmental parameters to investigate the independent and synergistic effects of changes in iron, copper, zinc and manganese concentration on geosmin, 2-MIB and biomass production by Streptomyces spp. The design included a mid-point condition (orange) and combinations of the variable levels as represented by the corners of the cubes.
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6.2.3 Media composition and culture conditions
Although the majority of previous studies examining geosmin and 2-MIB production by
Streptomyces have employed liquid cultures, due to the simplicity of the metabolite extraction procedure described in Chapter 4 for cultures growing on agar plates, solid medium containing 1.5% w/v agar was utilised throughout. All media were sterilised by autoclaving at 121°C for 20 minutes and after addition of sterile solutions as detailed below,
10 mL aliquots were distributed to sterile polypropylene Petri dishes (60 mm diameter).
Cultures were grown on sterile pre-weighed polycarbonate membranes (45 mm diameter,
0.45 µm pore size, Millipore) placed on the surface of the agar. The prepared solid media used for each experiment were inoculated with 250 µL of a spore suspension of S. coelicolor
A3(2) or S. antibioticus, which was carefully placed in the centre of the polycarbonate membranes. Following drying of the inoculum, the plates were inverted and incubated at the specified temperature for each experiment for 5 days. Preliminary experiments (data not shown) using a variety of solid growth media (SCA, AIA and NA) were undertaken to determine the optimal incubation time for development of aerial mycelium and for sporulation to occur.
6.2.3.1 Experiment 1: The effect of carbon, nitrogen and phosphorous concentration
For the study of the effects of carbon, nitrogen and phosphorous concentration on geosmin and 2-MIB production, defined solid media for each of the nine variable combinations
(Figure 6.1) were made by adding varying concentrations of mannitol (C22H14O14) (0.42 mM,
208.5 mM and 416.6 mM), as a sole carbon source, KNO3 and NH4Cl as equal sources of nitrogen (3.57 µM, 894.20 µM and 1,784.82 µM) and NaH2PO4·H2O as a source of phosphorous (1.61 µM, 404.42 µM and 807.23 µM). Solutions of the carbon, nitrogen and phosphorus sources were made in deionised water and sterilised by passing through 0.22 µm
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bacteriological filters prior to addition to the molten autoclaved media. Each medium received equal amounts of a sterile salt/trace elements solution that has been widely utilised in the laboratory for metabolic studies of Gram-positive bacteria (MgSO4.H20 0.718 mM,
MnSO4·7H2O 16.23 mM, MnSO4·H2O 2.35µM, H3BO3 11.3µM, Cu(NO3)3·3H2O
0.0413µM, ZnSO4·7H2O 0.208µM, NaMoO4·2H2O 1.95µM, (NH4)2Fe(SO4)2·6H2O 0.127
µM and CaCl·2H2O 0.45µM). All media were adjusted to pH 7.5 with the addition of 0.1M
NaOH. Inoculated plates were inverted and incubated at 28°C for 5 days.
6.2.3.2 Experiment 2: The effect of temperature, NaCl concentration and pH
Based on the results of Experiment 1, defined solid media for each of the nine variable combinations for Experiment 2 (Figure 6.2) all received mid-point concentrations of mannitol (208.5 mM) and low concentrations of KNO3 and NH4Cl (3.57 µM) and
NaH2PO4·H2O (1.61 µM). Each combination received the same concentration of a trace elements solution as stated above. Media containing 0%, 1% and 2% NaCl received the appropriate quantity of this ionic compound prior to autoclaving. The medium representing the centroid with a pH of 7.5 and media with a pH value of 9 were adjusted with the addition of 0.1M NaOH. The pH was lowered to 6 by the dropwise addition of 0.1M HCl. Following inoculation with each spore suspension, plates were incubated at the designated temperature according to the experimental design (24°C, 27°C or 30°C) for 5 days.
6.2.3.3 Experiment 3: The effect of calcium, magnesium and potassium concentration
All defined solid media for each of the nine variable combinations for Experiment 3 received concentrations of mannitol, KNO3 and NH4Cl and NaH2PO4·H2O and salts/trace elements solution (without as MgSO4.7H20) as stated above. To study the effect of calcium, magnesium and potassium concentration, varying concentrations of CaCl2, MgCl2 and KCl 193
were added (1 µM, 500.5 µM and 1000 µM) to make up the combination of factors according to the experimental design. Solutions of the chloride salts were made in deionised water and sterilised by passing through 0.22 µm bacteriological filters prior to addition to autoclaved media. Based on the results of Experiment 2, the pH was raised to 9 by the addition of 0.1M
NaOH and plates were incubated at 24°C for 5 days (based on Experiment 2).
6.2.3.4 Experiment 4: The effect of iron, zinc, copper and manganese concentration
Based on the previous three experiments conducted, all defined media for this experiment consisted of mannitol (208.5 mM), KNO3 and NH4Cl (3.57 µM), NaH2PO4·H2O (1.61 µM),
CaCl2 (1000 µM), MgCl2 (1 µM) and KCl (1000 µM) and were raised to pH 9 by the addition of 0.1M NaCl. All experimental combinations received a refined trace elements solution to eliminate micronutrients investigated in this study (H3BO3 11.3µM and NaMoO4·2H2O
1.95µM). Sulphate salts were used to study the effect of iron, copper, zinc and manganese concentration. Varying concentrations of FeSO4.7H20, CuSO4.5H20, MnSO4.H20 and
ZnSO4.7H20 were added (0.25µM, 12.51 µM and 25 µM) to make up the combination of factors according to the experimental design (Figure 6.4). All plates were incubated at 24°C for 5 days.
6.2.4 Biomass measurements and culture extraction of T&O metabolites
After the 5 days incubation period for each experiment, cultures were extracted with 1 mL of n-hexane (Sigma) according to the protocol described in Chapter 4 (section 4.2.6). Two hundred (200) µL of the extract was transferred to a GC vial insert followed by the addition of biphenyl-D10 as an internal standard (20 ng/mL) and BSTFA (50 µL). Vials were sealed with a septum screw cap. The biomass produced by each culture was determined gravimetrically by freeze drying the pre-weighed polycarbonate membranes to constant
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weight and subtracting the membrane weight. Production of geosmin and 2-MIB was expressed on a per dry weight basis (ng/mg).
6.2.5 Geosmin and 2-MIB analysis
Quantitation of geosmin and 2-MIB concentrations in culture extracts was determined using a Hewlett Packard 5973A gas chromatograph and mass spectrophotometer (GC-MS) instrument (Agilent Technologies) equipped with a HP-5ms capillary column (J & W,
Scientific Inc., 30 m x 0.25mm ID x 0.25 µm film thickness). Details of the operating conditions are described in Chapter 4 (section 4.2.6). Analytes were detected using selected ion monitoring (SIM) and the base peak ions m/z 95, 164 and 112 were used for quantification of 2-MIB, biphenyl-d10 and geosmin respectively. Quantification was determined by integrating the base peak area. The analytes were identified based on the retention time and mass spectra of standards. Calibration curves were established using diluted standards of geosmin and 2-MIB for quantification (4.2.6).
6.2.6 Microscopic assessment of morphological differentiation
Following growth under the specific combination of environmental conditions for each experiment, Streptomyces cells were subjected to microscopic morphological investigations.
A small sample of biomass was harvested using a sterile inoculating loop, Gram-stained and observed using light microscopy with oil emersion (1000x magnification). A visual assessment of the degree of differentiation was made according to the criteria presented in
Table 6.1, ranging from no differentiation (presence of vegetative mycelium only) to highly differentiated as observed by a high proportion of spores comprising the total biomass. Figure
6.5 shows a sequence of images detailing an increasing extent of Streptomyces morphological differentiation. The differentiation variable was expressed in values ranging from 0 to 5. The
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relationship between the degree of differentiation and geosmin and 2-MIB production was subsequently investigated.
Table 6.1 Stages of morphological differentiation and corresponding numerical values assigned for the analysis of the relationship between geosmin/2-MIB production and Streptomyces growth phase. Differentiation Description value 0 Vegetative mycelium only. 1 Beginning development of aerial mycelium (some septation); no sporulation. 2 Moderate development of differentiated aerial mycelium and some sporulation (biomass consisting of <10% spores). 3 Majority of the hyphal biomass is differentiated and moderate sporulation (biomass consisting of 10-50% spores). 4 Majority of the hyphal biomass is differentiated and heavy sporulation (biomass consisting of 10-50% spores) 5 Heavy sporulation occurring (biomass consisting of >50% spores)
Figure 6.5 Streptomyces biomass showing different degrees of morphological differentiation.
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6.2.7 Analysis and interpretation of the results
In order to analyse the relationship between the factors of each experiment and the response variables (geosmin, 2-MIB and biomass production), the data generated from the two-level full factorial design for each experiment were evaluated using Multiple Linear Regression
(MLR) as implemented in MODDE software version 9.0 (Umetrics, Umeå, Sweden). MLR is used to estimate the coefficients of the terms in the model (Equation 1). All variables are normalised to vary continuously between -1 and +1. Since variables are normalised in this way, the relative change of a variable is directly related to the size of its regression coefficient. If the model parameters have either a large positive or negative value, the corresponding variable has a large influence on the response variable. For presentation of the results, the regression coefficients for each model were plotted as bar charts with clear confidence intervals to indicate their significance (ɑ = 0.05). Factors with bigger bars are more influential. To interpret the meaning of the variables and their effects on the responses, contour plots were constructed. Furthermore, the raw data was analysed by ANOVA and multiple comparisons post hoc tests (Tukey-Kramer HSD method) using JMP version 7.0 to determine the statistical significance of observed variation between the different combination of variables on the production of geosmin, 2-MIB and biomass (results in Appendix F).
Regression analyses were used to determine the relationships between T&O metabolite yields and morphological differentiation. A confidence level of 95% was used throughout the data analysis.
6.3 Results
Contour plots were drawn to analyse the effects of experimental conditions on the production of geosmin, 2-MIB and biomass by the Streptomyces spp. The statistical significance of the
MLR models developed for each experiment can be interpreted in the regression coefficients
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plots presented. For each experiment conducted, the experimental design matrix and raw data for the end-point measures including geosmin and 2-MIB production (ng/mg), biomass (mg) and the assigned differentiation value for both Streptomyces spp. are presented in Appendix
F in addition to results pertaining to the validity of the models as determined using ANOVA.
Finally, the results concerning the relationship between T&O metabolite production and extent of Streptomyces morphological differentiation are presented.
6.3.1 Experiment 1: The effects of carbon, nitrogen and phosphorous concentration on geosmin and 2-MIB production by Streptomyces spp.
In Experiment 1, cultures of S. coelicolor A3(2) and S. antibioticus were grown under varying combinations of carbon (0.24-416.6 mM), nitrogen (3.57-1,784.82 µM ) and phosphorous
(1.61-807.23 µM) concentration. The independent and interaction effects of these three nutrient factors on the responses (geosmin, 2-MIB and biomass) are depicted in four- dimensional contour plots in Figures 6.6 and 6.7 for S. coelicolor A3(2) and S. antibioticus respectively. The scaled and centred regression coefficients obtained for each factor and the two-way interactions between them with 95% confidence intervals for MLR models of geosmin, 2-MIB and biomass production are shown in the bar charts in Figures 6.8 and 6.9
(S. coelicolor A3(2) and S. antibioticus respectively). The more significant the effect is on the response variable, the larger the regression coefficient and smaller the p-value is for the term in the model, which are both stated in the plots.
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Figure 6.6 Four dimensional contour plots showing the influence of nitrogen, phosphorous and carbon concentration on the production of geosmin (top), 2-MIB (middle) and biomass (bottom) by S. coelicolor A3(2) as modelled with MLR. 199
Figure 6.7 Four dimensional contour plots showing the influence of nitrogen, phosphorous and carbon concentration on the production of geosmin (top), 2-MIB (middle) and biomass (bottom) by S. antibioticus as modelled with MLR.
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Figure 6.8 Scaled and centered MLR coefficients of experimental variables (carbon, nitrogen and phosphorous concentration) and variable interactions for geosmin (top), 2-MIB (middle) and biomass (bottom) production by S. coelicolor A3(2). Coefficients and p-values are stated and the statistically significant (p<0.05) coefficients are indicated with an asterisk.
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Figure 6.9 Scaled and centered MLR coefficients of experimental variables (carbon, nitrogen and phosphorous concentration) and variable interactions for geosmin (top), 2-MIB (middle) and biomass (bottom) production by S. antibioticus. Coefficients and p-values are stated and the statistically significant (p<0.05) coefficients are indicated with an asterisk.
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For S. coelicolor A3(2), the three contour plots (Figure 6.6) of nitrogen versus phosphorous with increasing carbon concentration from left to right depict the negative effect of all three factors on biosynthesis of T&O compounds, as the highest production of geosmin and 2-MIB
(21.80 ± 2.23 ng/mg and 11.13 ± 2.51 ng/mg respectively) occurred when the concentration of all three nutrients were at their lowest level. In comparison, cultures grown on the medium containing the highest concentration of all three nutrients yielded the lowest production, with geosmin and 2-MIB yields reduced by over 10-fold (1.29 ± 0.32 ng/mg and 0.60 ± 0.34 ng/mg respectively). Carbon, phosphorous and nitrogen concentration all had statistically significant negative effects on geosmin and 2-MIB production by S. coelicolor A3(2) (Figure
6.9). The significant positive interaction coefficients between carbon and phosphorous and phosphorous and nitrogen indicate that geosmin and 2-MIB production was greatest under low concentration combinations of these variables. However, the carbon and nitrogen interaction was not found to influence geosmin production significantly, while none of the interaction coefficients were found to be significant in relation to production of 2-MIB.
In terms of biomass production, the contour plots (Figure 6.6) reveal the opposite effect of the factors, with the greatest biomass produced by cultures grown on the medium containing carbon, phosphorous and nitrogen at their highest concentration (7.71 ± 0.76 mg) compared to the combination of their lowest concentration (0.96 ± 0.05 mg). The most significant factor was carbon, followed by phosphorous, nitrogen and the synergy between carbon and phosphorous, for which increasing concentration had positive effects (Figure 6.8). These significant positive regression coefficients indicate that an increase in nutrient concentration favours more biomass development. For S. antibioticus, the regression coefficients shown in
Figure 6.9 indicate the same significant independent and synergistic effect of the factors on biomass development, with maximal production (7.41 ± 0.3 mg) observed when all nutrients
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were in high concentration compared to the combination of their lowest concentration (0.74
± 0.09) as revealed in the contour plots (Figure 6.7).
While nitrogen and phosphorous and the interaction between these variables were found to significantly influence geosmin and 2-MIB production by S. antibioticus (reduced concentrations of both nutrients increased production), the effect of carbon was found to be insignificant (Figure 6.9). The contour plots (Figure 6.7) reveal the effect of the combination of lowest phosphorous and nitrogen concentration favouring higher production of both geosmin and 2-MIB (10-fold greater than the combination of the highest concentration of these factors). Maximum 2-MIB (16.45 ± 2.32 ng/mg) was obtained in cultures provided with the highest carbon concentration and lowest nitrogen and phosphorous concentrations, compared to a yield of 11.60 ng/mg when all three nutrients were in lowest concentration. In the case of geosmin, the insignificant influence of carbon concentration is depicted with statistically similar maximal concentrations obtained for high and low carbon concentrations
(11.31 ± 1.26 ng/mg and 13.15 ± 1.90 ng/mg respectively) as indicated by the similar absolute values between the three contour plots.
6.3.2 Experiment 2: The effects of temperature, pH and NaCl concentration on geosmin and 2-MIB production by Streptomyces spp.
The effects of different levels of pH (6-9), temperature (24-30°C) and NaCl concentration (0-
2%) on geosmin, 2-MIB and biomass production were examined in the second two-level full factorial experiment (Experiment 2). The influence of these three environmental parameters on the response variables are revealed in Figures 6.10 and 6.11 for S. coelicolor A3(2) and
S. antibioticus respectively, depicting the combined effects of NaCl concentration and pH in each contour plot, with increasing temperature from left to right. The regression coefficients
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\ Figure 6.10 Four dimensional contour plots showing the influence of NaCl concentration, pH and temperature on the production of geosmin (top), 2-MIB (middle) and biomass (bottom) by S. coelicolor A3(2) as modelled with MLR.
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Figure 6.11 Four dimensional contour plots showing the influence of NaCl concentration, pH and temperature on the production of geosmin (top), 2-MIB (middle) and biomass (bottom) by S. antibioticus as modelled with MLR.
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for each of the response variables computer by MLR for S. coelicolor A3(2) and S. antibioticus are shown in the bar charts in Figures 6.12 and 6.13 respectively, indicating the effect of each of the terms in the model (i.e. the individual factors and interactions between them).
Figure 6.12 Scaled and centered MLR coefficients of experimental variables (NaCl concentration, pH and temperature) and variable interactions for geosmin (top), 2-MIB (middle) and biomass (bottom) production by S. coelicolor A3(2). Coefficients and p-values are stated and the statistically significant (p<0.05) coefficients are indicated with an asterisk.
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Figure 6.13 Scaled and centered MLR coefficients of experimental variables (NaCl concentration, pH and temperature) and variable interactions for geosmin (top), 2-MIB (middle) and biomass (bottom) production by S. antibioticus. Coefficients and p-values are stated and the statistically significant (p<0.05) coefficients are indicated with an asterisk.
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For both Streptomyces spp. it can be clearly seen that the most influential factor is NaCl concentration. The statistically significant negative regression coefficients (Figures 6.12 and
6.13) for NaCl in terms of geosmin and 2-MIB production indicate that increasing this parameter decreases production of both secondary metabolites, while the significant positive regression coefficients for the biomass models indicates that increasing the NaCl concentration favours biomass development. NaCl concentration exhibited the most significant influence on T&O metabolite biosynthesis independent of the temperature and pH levels. Increasing this factor from its lowest (0%) to highest level (2%) substantially decreased production of these compounds by an order of magnitude for both Streptomyces spp. (Figures 6.10 and 6.11). A statistically significant positive effect of raising the pH from
6 to 9 on production of geosmin and 2-MIB was observed for both species, whereas bacterial growth was greater under conditions of lower pH for S. antibioticus although this was not observed for S. coelicolor A3(2).
The greatest geosmin and 2-MIB production by S. coelicolor A3(2) occurred in the combination of highest pH, lowest NaCl concentration and lowest temperature tested (40.66
± 11.50 ng/mg and 10.40 ± 0.84 respectively) while low yields were produced by all cultures grown under conditions of high NaCl, irrespective of the temperature and pH levels (<1.5 ng/mg for geosmin and <1 ng/mg for 2-MIB). Similarly, maximal yields of geosmin (192.53
± 17.99 ng/mg) also occurred under these conditions in cultures of S. antibioticus as observed for 2-MIB apart from the insignificant differences between cultures incubated under different temperatures (36.34 ± 25 ng/mg and 38.71 ± 41 ng/mg for 24°C and 30°C respectively). The sequence of contour plots from left to right represents the effects observed at increasing temperature levels. Increasing temperature exerted a negative effect on geosmin and 2-MIB production by both species, however this effect was not significant for 2-MIB production by
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S. antibioticus. Higher temperature positively influenced biomass development, but this was only significant for S. antibioticus.
6.3.3 Experiment 3: The effects of calcium, potassium and magnesium concentration on geosmin and 2-MIB production by Streptomyces spp.
The effect of changes in the concentration of three macronutrients including calcium, potassium and magnesium over the specified range (1-1000 µM) on geosmin, 2-MIB and biomass production by Streptomyces spp. was examined in the third two-level full factorial experiment (Experiment 3). The four-dimensional contour plots in Figures 6.14 and 6.15 for
S. coelicolor A3(2) and S. antibioticus respectively, depict the independent and interaction effects of the macronutrient factors on the responses. The series of three contour plots show the combined effects of calcium and potassium concentration, with increasing magnesium concentration from left to right. The regression coefficients and their associated 95% confidence intervals are displayed in Figure 6.16 and 6.17 for S. coelicolor A3(2) and S. antibioticus respectively.
The positive effects of calcium and potassium concentration on geosmin and 2-MIB production by both species are clearly depicted. These positive effects were all statistically significantly with the exception of potassium for 2-MIB produced by S. antibioticus. The synergy between calcium and potassium positively influenced production of both secondary metabolites, although this effect was only significant for S. coelicolor A3(2). Magnesium concentration appeared to exhibit negative effects on T&O metabolite production which were statistically significant with the exception of 2-MIB production by S. antibioticus.
Synergistic effects between magnesium and other macronutrients were all negative, with the
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Figure 6.14 Four dimensional contour plots showing the influence of potassium, calcium and magnesium concentration on the production of geosmin (top), 2-MIB (middle) and biomass (bottom) by S. coelicolor A3(2) modelled with MLR.
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Figure 6.15 Four dimensional contour plots showing the influence of potassium, calcium and magnesium concentration on the production of geosmin (top), 2-MIB (middle) and biomass (bottom) by S. antibioticus.
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Figure 6.16 Scaled and centered MLR coefficients of experimental variables (calcium, potassium and magnesium concentrations) and variable interactions for geosmin (top), 2- MIB (middle) and biomass (bottom) production by S. coelicolor A3(2). Coefficients and p- values are stated and the statistically significant (p<0.05) coefficients are indicated with an asterisk.
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Figure 6.17 Scaled and centered MLR coefficients of experimental variables (calcium, potassium and magnesium concentrations) and variable interactions for geosmin (top), 2- MIB (middle) and biomass (bottom) production by S. antibioticus. Coefficients and p-values are stated and the statistically significant (p<0.05) coefficients are indicated with an asterisk.
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interaction between magnesium and calcium for both compounds and between magnesium and potassium for 2-MIB being significant in S. coelicolor A3(2) cultures.
Maximal yields of geosmin and 2-MIB were produced by cultures grown under the combination of the highest concentration of calcium and potassium and lowest concentration of magnesium in the medium, being 180.92 ± 25.54 ng/mg and 58.75 ± 8.20 ng/mg respectively for S. coelicolor A3(2) and 111.46 ± 46±15.16 ng/mg and 25.12 ± 8.51 ng/mg respectively for S. antibioticus. For S. coelicolor A3(2), the lowest yield of geosmin (65.24
± 10.11 ng/mg) was detected in cultures grown under conditions of highest calcium and magnesium concentration and lowest potassium concentration, whereas for 2-MIB, all cultures grown under conditions of low calcium concentration (irrespective of magnesium and potassium concentration) produced statistically similar low yields of 2-MIB (ranging from 20.60 ± 0.64 ng/mg to 25.04 ± 6.05 ng/mg). For S. antibioticus, geosmin biosynthesis was lowest under conditions of low calcium and potassium concentration and high magnesium concentration (41.57 ± 2.43 ng/mg), however this was not significantly different to amounts produced under all other conditions aside from the combination of factors favouring highest production (lowest magnesium and highest calcium and potassium concentration). In terms of 2-MIB biosynthesis by this species, production was lowest when the concentration of all macronutrients was lowest (9.30±1.15 ng/mg). In general, biomass production by both species did not differ greatly across the different nutrient regimes, although growth by S. coelicolor A3(2) was favourable in the presence of lowest calcium and highest potassium concentration (regardless of magnesium concentration) whereas greater magnesium concentration appeared to be more favourable for growth in the case of S. antibioticus.
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6.3.4 Experiment 4: The effects of iron, zinc, copper and manganese concentration on geosmin and 2-MIB production by Streptomyces spp.
The influence of the concentration of four micronutrients including iron, zinc, copper and manganese over the specified range (0.25 µM to 25 µM) on geosmin, 2-MIB and biomass production by Streptomyces spp. was examined in Experiment 4. The effects of the micronutrient factors and synergies between them on the responses are represented by the contour plots in Figures 6.18, 6.19 and 6.20 and in Figures 6.21, 6.22 and 6.23 for S. coelicolor A3(2) and S. antibioticus respectively. These complex figures present a series of nine contour plots, depicting the combined effects of iron and copper concentration, with an increase in manganese concentration from left to right and an increase in zinc concentration from bottom to top. The regression coefficients, revealing the effects and significance (95% confidence level) of the factors and interactions between them on each of the response variables as determined by MLR for S. coelicolor A3(2) and S. antibioticus are shown in the bar charts in Figures 6.24 and 6.25 respectively.
For both species, the most evident feature of the data was the effect of reduced production of both secondary metabolites in relation to increasing copper concentration. In the case of S. coelicolor A3(2), 10-fold reductions in the concentration of both compounds were observed from this statistically significant negative effect. All cultures containing the highest copper concentration, irrespective of other factor levels, produced the lowest and statistically similar concentrations of geosmin and 2-MIB. For S. coelicolor A3(2), maximal yields of geosmin and 2-MIB were recovered from cultures grown under conditions of highest iron and lowest zinc, copper and manganese concentration (135.26 ± 36.67 ng/mg and 43.95 ± 11.05 ng/mg respectively). The decrease in the absolute value range of the contour plots, observed from bottom to top and from left to right, depict the negative influences of zinc and manganese
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Figure 6.18 Contour plots showing the influence of micronutrient (iron, copper, zinc and manganese) concentration on the production of geosmin by S. coelicolor A3(2).
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Figure 6.19 Contour plots showing the influence of micronutrient (iron, copper, zinc and manganese) concentration on the production of 2-MIB by S. coelicolor A3(2).
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Figure 6.20 Contour plots showing the influence of micronutrient (iron, copper, zinc and manganese) concentration on the production of biomass by S. coelicolor A3(2).
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Figure 6.21 Contour plots showing the influence of micronutrient (iron, copper, zinc and manganese) concentration on the production of geosmin by S. antibioticus.
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Figure 6.22 Contour plots showing the influence of micronutrient (iron, copper, zinc and manganese) concentration on the production of 2-MIB by S. antibioticus. 221
Figure 6.23 Contour plots showing the influence of micronutrient (iron, copper, zinc and manganese) concentration on the production of biomass by S. antibioticus.
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Figure 6.24 Plot of scaled and centered MLR coefficients of experimental variables (micronutrients) and variable interactions for geosmin (top), 2-MIB (middle) and biomass (bottom) by S. coelicolor A3(2). Coefficients and p-values are stated and the statistically significant (p<0.05) coefficients are indicated with an asterisk.
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Figure 6.25 Plot of scaled and centered MLR coefficients of experimental variables (micronutrients) and variable interactions for geosmin (top), 2-MIB (middle) and biomass (bottom) production by S. antibioticus. Coefficients and p-values are stated and the statistically significant (p<0.05) coefficients are indicated with an asterisk.
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concentration respectively on production of both compounds, and the positive but relatively insignificant effect of iron.
Similarly for S. antibioticus, the positive effect of iron on T&O metabolite production can be seen, but this was dependent on low copper concentration and overall the effect was not considered significant. Maximal yields of geosmin and 2-MIB for this species, similar to S. coelicolor A3(2), were recovered from cultures subjected to the combination of highest iron and lowest zinc, copper and manganese concentration (37.53 ± 1.78 ng/mg and 8.94 ± 2.00 ng/mg respectively). The negative effect of zinc can be seen with the reduction in absolute values in the contour plots from the bottom to the top whilst manganese had little effect.
In terms of bacterial growth, the effect of micronutrients and their synergies revealed no clear pattern in S. coelicolor A3(2), with maximal biomass observed under the conditions of highest iron and copper and lowest manganese and zinc whilst also under combined conditions of the other levels of these factors. For S. antibioticus, maximal biomass was observed in cultures grown under conditions of highest concentration of all micronutrients and was lowest with the lowest concentration of all micronutrients aside from iron.
6.3.5 The relationship between morphological differentiation and T&O metabolite production by Streptomyces spp.
Following growth of Streptomyces spp. under the different combination of factors in each experiment, an assessment of the extent of morphological differentiation was made.
Biomass obtained from all experimental treatments was Gram-stained, examined microscopically and assigned a differentiation value according to the criteria presented in
Table 6.1. Scatter plots of geosmin and 2-MIB concentrations versus the differentiation
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value from all experiments are presented in Figures 6.26 and 6.27 for S. coelicolor A3(2) and S. antibioticus respectively. The positive relationship between the extent of biomass differentiation and the yield of geosmin and 2-MIB produced by the Streptomyces spp. cultures is evident in the graphs. The R2 values for the relationship (exponential trend line) in these graphs which combine the data from all experiments were moderately positive. These were substantially higher when the analysis was carried out on data corresponding to individual experiments (see results in Appendix F).
Experiment 1 Experiment 2 Experiment 3 Experiment 4 200 y = 3.359e0.567x 180 Geosmin R² = 0.577 160 140 120 100 80 60
40 Concentration (ng/mg) Concentration 20 0 0 1 2 3 4 5 Differentiation value Experiment 1 Experiment 2 Experiment 3 Experiment 4 70 y = 1.822e0.429x R² = 0.400 60 2-MIB 50 40 30 20
Concentration (ng/mg) Concentration 10 0 0 1 2 3 4 5 Differentiation value
Figure 6.26 Relationship between geosmin (top) and 2-MIB (bottom) production by S. coelicolor A3(2) and differentiation value based on all experimental data.
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Experiment 1 Experiment 2 Experiment 3 Experiment 4 y = 1.619e0.605x 250 Geosmin R² = 0.598
200
150
100
50 Concentration (ng/mg) Concentration 0 0 1 2 3 4 5 Differentiation value
Experiment 1 Experiment 2 Experiment 3 Experiment 4 50 y = 0.816e0.506x 2-MIB R² = 0.569 40
30
20
10 Concentration (ng/mg) Concentration 0 0 1 2 3 4 5
Differentiation value Figure 6.27 Relationship between geosmin (top) and 2-MIB (bottom) production by S. antibioticus and differentiation value based on all experimental data.
Light microscope photographs indicating the differences in morphological differentiation that occurred under each treatment in the experiments (indicated by the experimental ID) for both Streptomyces spp. are presented in Figures 6.28 to 6.33. In Experiment 1, the predominance of vegetative hyphae (differentiation value of 0) observed under the combination of highest (ID 1.4) and mid-level (ID 1.5) concentrations of carbon, nitrogen and phosphorous levels are evident in Figures 6.28 and 6.29 for S. coelicolor A3(2) and
S. antibioticus respectively. Conversely, highly differentiated (sporulating) biomass was observed in both Streptomyces spp. subjected to the combination of lowest concentrations
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of these macronutrients (ID 1.6 in Figures 6.28 and 6.29). Concerning Experiment 2, the observation of a lower degree of morphological differentiation under conditions of highest NaCl concentration (IDs 2.2, 2.4, 2.7 and 2.9) are evident in Figures 6.30 and
6.31 for S. coelicolor A3(2) and S. antibioticus respectively, while cultures grown under low NaCl concentration (IDs 2.1, 2.3, 2.6 and 2.8), exhibited highly differentiated morphologies. Variation in morphological differentiation was not observed across the levels of temperature and pH. Under all combinations of calcium, potassium and magnesium concentrations examined in Experiment 3, variation was not observed, with all cultures exhibiting high degrees of morphological differentiation (photos not shown).
Figures 6.32 and 6.33 show the differences in morphological differentiation observed in
Experiment 4 treatments for S. coelicolor A3(2) and S. antibioticus respectively, with the general observation of predominately vegetative hyphae in cultures exposed to highest copper concentrations (IDs 4.1, 4.2, 4.5, 4.6, 4.10, 4.11, 4.14 and 4.15) while those grown under conditions of lowest copper concentration (IDs 4.3, 4.4, 4.7, 4.8, 4.12, 4.13, 4.16 and 4.17) displayed higher extents of morphological differentiation.
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Figure 6.28 Light microscope (1000x) photographs of Gram-stained S. coelicolor A3(2) biomass growing under the different combinations of carbon, nitrogen and phosphorous concentrations. The experimental ID is indicated which corresponds to the combination of factors specified in the experimental matrix. The differentiation value (0-5) is in italics.
Figure 6.29 Light microscope (1000x) photographs of Gram-stained S. antibioticus biomass growing under the different combinations of carbon, nitrogen and phosphorous concentrations. The experimental ID is indicated which corresponds to the combination of factors specified in the experimental matrix. The differentiation value (0-5) is in italics.
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Figure 6.30 Light microscope photographs of Gram-stained S. coelicolor A3(2) biomass growing under different conditions of temperature, pH and NaCl concentration. The experimental ID is indicated which corresponds to the combination of factors specified in the experimental matrix. The differentiation value (0-5) is also provided in italics.
Figure 6.31 Light microscope photographs of Gram-stained S. antibioticus biomass growing under different conditions of temperature, pH and NaCl concentration. The experimental ID is indicated which corresponds to the combination of factors specified in the experimental matrix. The differentiation value (0-5) is also provided in italics.
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Figure 6.32 Light microscope photographs of Gram-stained S. coelicolor A3(2) biomass growing under different conditions of micronutrient (Fe, Zn, Cu and Mn) concentration. The experimental ID is indicated which corresponds to the combination of factors specified in the experimental matrix. The differentiation value (0-5) is also provided in italics.
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Figure 6.33 Light microscope photographs of Gram-stained S. antibioticus biomass growing under different conditions of micronutrient (Fe, Zn, Cu and Mn) concentration. The experimental ID is indicated which corresponds to the combination of factors specified in the experimental matrix. The differentiation value (0-5) is also provided in italics.
6.3.6 Summary of key findings
This study has demonstrated that geosmin and 2-MIB production by two Streptomyces spp. was influenced by changes in various physico-chemical parameters. Additionally, a relationship between T&O metabolite production and morphological differentiation was established. The major findings from this series of experiments are summarised in Table
6.2 and can be outlined as follows:
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Lower carbon, nitrogen and phosphorous concentrations resulted in greater
production of geosmin and 2-MIB by Streptomyces spp., although carbon levels
did not appear to be influential in this manner in the case of S. antibioticus.
Conversely, the combination of high concentrations of these macronutrients
favoured maximal biomass development.
The amount of geosmin and 2-MIB produced by both Streptomyces spp. was
profoundly reduced under conditions of increased NaCl concentration and this
factor stimulated greater biomass development. Increasing the pH of the growth
media stimulated more geosmin and 2-MIB production by both Streptomyces spp.,
whereas increases in the incubation temperature largely reduced production
yields.
Increases in calcium and potassium concentration were found to positively
influence geosmin and 2-MIB production by both Streptomyces spp., whereas an
increase in magnesium concentration appeared to exhibit the reverse effect.
Of the micronutrients investigated in the final multifactorial experiment, copper
exhibited the greatest effect by significantly inhibiting production of geosmin and
2-MIB. Changes in the concentration of other micronutrients revealed
comparatively minor effects.
The extent of morphological differentiation observed in Streptomyces spp.
cultures was positively related to yields of geosmin and 2-MIB recovered in the
experiments. Generally, cultures that revealed a low degree of morphological
differentiation (mostly vegetative mycelium) developed greater biomass whereas
those that displayed highly differentiated secondary mycelium and the presence
of free spores produced higher yields of the T&O metabolites.
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Table 6.2 Summary of the association of physico-chemical parameters and life cycle stage of Streptomyces with geosmin and 2-MIB production. Factors associated with Factors associated with Factors exhibiting reduced geosmin and 2- elevated geosmin and 2- little effect on geosmin MIB production MIB production and 2-MIB production Vegetative growth Secondary mycelial Zn, Mn and Fe conc. differentiation and sporulation ↑ N, P and C conc. ↓ N, P and C conc. ↑ NaCl conc. ↑pH ↑Temperature ↑ Ca and K conc. ↑ Mg conc. ↑ Cu conc.
6.4 Discussion
A key objective of the research presented in this chapter was to acquire evidence to assess the apparent association between morphological differentiation in Streptomyces and geosmin and 2-MIB production that has been documented in the literature (Bentley &
Meganathan, 1981; Dionigi et al., 1992; Scholler et al., 2002; Tung et al., 2006). The experiments conducted revealed that the level of production of these T&O metabolites by
Streptomyces spp. was closely tied to the growth phase of the organisms and supported the hypothesis that greater yields are produced by cultures exhibiting higher degrees of morphological differentiation compared to those at the primary vegetative growth phase.
It therefore seems logical to infer that an understanding of the physico-chemical factors that prompt Streptomyces to cease vegetative growth and enter the reproductive developmental stage would also aid in elucidating the conditions which trigger significant production of their T&O metabolites. Such conditions were hypothesised to be limiting concentrations of major macronutrients (carbon, nitrogen and phosphorous), the common environmental trigger for Streptomyces to switch to secondary mycelial growth and sporulation. The data obtained from the first multifactorial experiment largely supported this hypothesis. Subsequent experiments examined a number of other factors considered
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likely to inhibit vegetative growth and/or stimulate sporulation and thus trigger production of geosmin and 2-MIB. Contrary to expectations, increasing the NaCl concentration favoured vegetative growth, with significantly more geosmin and 2-MIB obtained in highly differentiated cultures that were subjected to the lowest NaCl concentration. Similar to NaCl, high concentrations of copper significantly inhibited
T&O metabolite production and corresponded to cultures exhibiting low extents of morphological differentiation (vegetative growth). Other micronutrients were comparatively less influential on T&O metabolite production. A preference for the production of T&O metabolites under more alkaline conditions and lower temperature was established, although these parameters did not influence the growth stage, suggesting that these effects of pH and temperature may be independent of morphological differentiation. Changes in the concentration of other macronutrients examined appeared to influence T&O metabolite production, with calcium and potassium revealing positive effects and magnesium, a negative effect. These however, were less significant compared to those of other factors and did not cause variations in the extent of morphological differentiation. The key findings outlined are further elaborated throughout the discussion that follows.
6.4.1 The relationship between geosmin and 2-MIB and morphological differentiation in Streptomyces spp.
The reproductive stage of development, when aerial hyphae form with their eventual differentiation into chains of spores, is known to occur once the opportunity for vegetative growth diminishes. The transition generally occurs as a result of nutrient exhaustion as the vegetative hyphae are broken down to release nutrients to fuel growth of the reproductive secondary mycelium. At this point, metabolic shifts generally occur, setting secondary metabolite production in motion (Bibb, 2005; van Keulen et al., 2011). Thus the observation of relatively low biomass and maximal geosmin and 2-MIB levels when major nutrients (carbon, nitrogen and phosphorous) were lowest is biologically 235
understandable and consistent with hypothetical expectations. In a natural setting,
Streptomyces are particularly vulnerable in this transition phase, needing to defend nutrients released from vegetative hyphae for reproductive development. Secondary metabolites which are produced at this stage in the life cycle of Streptomyces have been proposed to serve the role of antagonising competing microbes in defence of their nutrient supply. Whether or not this is the functional role of geosmin and 2-MIB is unclear and is further investigated in Chapter 7. Indeed a single rationale for explaining the diversity of chemical structures and biological activities of secondary metabolites is probably inadequate (van Keulen et al., 2011).
The notion that nutrient downshift reduces growth and induces differentiation and secondary metabolism as found in this study, has been supported by much of the published literature concerning Streptomyces (Kendrick & Ensign, 1983; Koepsel &
Ensign, 1984; Daza et al., 1989; Karandikar et al., 1997). Culture-based studies have observed that highly differentiated and sporulating Streptomyces cultures have been associated with enhanced T&O metabolite production (Scholler et al., 2002; Tung et al.,
2006). At the molecular level, genetic coregulation of morphological differentiation and earthy-musty secondary metabolite production is also well supported. Yague et al. (2013) compared the transcriptomes (levels of gene expression) between the vegetative mycelium (MI phase) and the aerial mycelium and sporulation phases (MII phase) of the
S. coelicolor A3(2) developmental cycle, reporting that geosmin production genes (geoA, cyclase) were well correlated with MII differentiation in addition to other genes involved in bioactive compound production (e.g. actinorhodin and undecylprodigiosin), aerial mycelium differentiation (e.g. bld) and sporulation (e.g. whi). Gust et al. (2003) also found that mutations in the gene (cyc2) coding the sesquiterpene synthase enzyme
(involved in the initial steps in geosmin biosynthesis) caused reduced sporulation by
Streptomyces. Other reports have shown that Streptomyces mutants which could not produce spores or aerial mycelia lost the ability to produce geosmin (Bentley & 236
Meganathan, 1981; Dionigi et al., 1992). Normal isolates grown on a medium that was not conducive to sporulation reduced their geosmin biosynthesis compared to those grown on a medium that promoted sporulation (Dionigi et al., 1992).
6.4.2 Carbon, phosphorous and nitrogen in relation to geosmin and 2-MIB production by Streptomyces spp.
An important point of consideration was the finding that the combination of lowest carbon, nitrogen and phosphorous concentration elicited the greatest production of both compounds in the case of S. coelicolor A3(2), while the same effect was not observed in cultures of S. antibioticus. Geosmin production was found to be relatively insensitive to carbon concentration over the range tested. Doull and Vining (1990) observed a similar phenomenon whereby production of the antibiotic actinorhodin by S. coelicolor A3(2) was insensitive to carbon (starch) concentration but was elicited by nitrogen (glutamate) and phosphorous (K2HPO4 and KH2PO4) depletion accompanied by a decline in growth rate. Thus actinorhodin biosynthesis was not initiated by carbon source depletion or inhibited by excess amounts of carbon, suggesting that actinorhodin biosynthesis is not subject to carbon catabolite repression (Ruiz et al., 2010)2. A CCR effect was evident in the case of high concentration of mannitol in cultures of S. coelicolor A3(2), although this carbon source did not exhibit CCR on geosmin and 2-MIB production by S. antibioticus. This explanation may also account for lack of carbon control over geosmin biosynthesis by S. antibioticus. For this species, maximal 2-MIB production occurred under conditions of higher carbon concentration. A similar observation was made by
Yagi, Sugiura and Sudo (1987) when they subjected liquid cultures of a Streptomyces reservoir isolate to varying concentrations (0 - 24.6 mM) of starch as a carbon source, with higher concentrations found to favour both biomass and production of geosmin and
2 Sources of carbon such as glucose that are readily used for growth, often repress secondary metabolism, a situation known as carbon catabolite repression (CCR) (Ruiz et al., 2010).
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2-MIB. This is the only other report identified in the literature where the effect of carbon concentration on earthy-musty odour production by Streptomyces has been examined.
Indeed, several studies have investigated the effects of different carbon sources on T&O metabolite production, reporting that some carbon sources such as glucose favoured more biomass development and less production of the compounds whereas others such as glycerol and succinate supported less growth and greater geosmin and 2-MIB production
(Weete et al., 1977; Schrader & Blevins, 2001). Mannitol, utilised as the designated carbon source in this study, has long been associated with T&O production by
Streptomyces with Sivonen (1982) identifying production of an earthy odour by two
Streptomyces spp. when grown in a mannitol based medium, while Schrader and Blevins
(2001) identified that among a range of other carbon sources, this compound promoted maximal geosmin yields by S. halstedii.
Numerous studies have examined the influence of phosphorous concentration on geosmin production by Streptomyces, with contradictory results reported. Schrader and Blevins
(2001) found that maximal geosmin production by S. halstedii occurred at the highest phosphorous concentration tested (36.2 µM) whereas production did not occur at the lowest concentrations tested (0.7 µM and 3.6 µM). Contrary to this, Weete et al. (1977) found that increasing phosphorous concentrations over a similar range (0.83 µM to 25.0
µM), exhibited no stimulatory effects on geosmin production by a Streptomyces isolate, and in the current study, maximal geosmin and 2-MIB production occurred when cultures were subjected to a low phosphorous concentration of 1.6 µM, whereas a high concentration (807.2 µM) repressed production. These results may not be directly comparable due to disparity in the phosphorous concentration ranges examined. The maximal level of phosphorous may not have been sufficient to repress geosmin biosynthesis in the studies by Schrader and Blevins (2001) and Weete et al. (1977) as demonstrated in this study. Results that do align with this study were reported by Yagi et al. (1987). These authors subjected liquid cultures of two Streptomyces reservoir isolates 238
to varying concentrations of phosphorous (20.4 µM, 51.0 µM, 102.1 µM, 306.3 µM,
612.6 µM and 1,020.8 µM), observing maximal production of geosmin and 2-MIB at the low concentrations of 51.0 µM for one isolate and 20.4 µM for the other, with rapid decreases in production as the phosphorous levels were increased.
Contradictory findings were reported by Uwins (2011), who found significantly greater geosmin production by liquid cultures of S. coelicolor A3(2) when grown under the highest phosphorous concentration (13,090 µM), compared to phosphorous limited cultures (0.32 µM, 3.2 µM and 32 µM). The rationale provided by the author for the observation of optimal geosmin production occurring under conditions of high phosphorous is that this is an essential element to cell growth and metabolism (a key component of nucleic acids, phospholipids in cell membranes and energy transfer as
ATP) and is involved in the isoprenoid pathway with pyrophosphates being the precursors to geosmin molecules (see Figure 2.3 in Chapter 2). Indeed the metabolic pathway involved in geosmin and 2-MIB synthesis is intrinsically linked to phosphate, with the precursors being farnesyl diphosphate and geranyl diphosphate for geosmin and 2-MIB respectively. However, the conversion of these terpenes to odorous secondary metabolites requires the removal of the phosphate group, which is then used for other metabolic processes within the cell. It is feasible that geosmin and 2-MIB may be incidental by- products of an organism’s need to sequester phosphate for essential metabolic processes during times of phosphorous limitation. Such a premise is consistent with the findings of the present study. To account for these contradictory results, it may be that two independent mechanisms of phosphorous control on production of these compounds exist, one under phosphorous limiting conditions and another under conditions of phosphorous surplus.
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It makes sense that a phosphorous deficient environment would support more geosmin and 2-MIB production. The negative control of phosphorous on the biosynthesis of a number of secondary metabolites by Streptomyces through the derepression of biosynthetic genes during times of phosphorous limitation has been well documented
(Gersch et al., 1979; Martin et al., 2011; van Wezel & McDowall, 2011). High phosphorous concentrations which are favourable for growth, have been shown to inhibit the production of a number of other secondary metabolites such as streptomycin (>10 mM), candicidin (>5 mM) and rapamycin (>10 mM) (Miller & Walker, 1970; Cheng et al., 1995), similar to the effects of phosphorous on geosmin and 2-MIB observed here.
This control of secondary metabolite production by phosphate appears to be a regulatory system that has evolved in bacteria in response to nutrient scarcity. In soil, the major habitat of Streptomyces, the concentration of phosphorous in its preferred form for metabolism (inorganic orthophosphate) is typically low. When phosphate levels are depleted, production of secondary metabolites is triggered serving to antagonise other competing microbes or acting as biochemical cross-talk signals to enhance survival under harsh nutritional conditions (Martin et al., 2011).
Of recent years, the mechanism of phosphate control over the transcription of biosynthetic and/or regulatory genes associated with secondary metabolism has been unravelled (van
Wezel & McDowall, 2011). In Streptomyces, a two component system called the PhoR-
PhoP system mediates the control of primary and secondary metabolism by phosphate.
When phosphorous becomes limiting for growth, a membrane sensor kinase PhoR phosphorylates PhoP, a DNA-binding response regulator, enabling it to bind strongly to its cognate sites in DNA (called PHO boxes) and therefore regulate transcription. During conditions of phosphorous sufficiency, PhoR is prevented from phosphorylating PhoP and when phosphorous levels drop, PhoR is released and phosphorylates PhoP, activating
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the transcription of phosphate scavenging and transport genes and other genes such as those encoding the enzymes for secondary metabolites. PhoP controls intermediate regulators of these genes in a regulatory cascade that in turn modulates expression of pathway specific regulations (Martin, 2004; van Wezel & McDowall, 2011).
The effect of nitrogen concentration on T&O metabolite production by Streptomyces has previously been investigated and results reported are in general agreement with the findings of the present study, with the observation of cultures under low nitrogen conditions producing more geosmin and 2-MIB. One exception was the study of Yagi et al. (1987) which found that over the range of nitrogen concentrations tested (0-13,698.9
µM), maximal production occurred at relatively high levels between 2,054.8 µM and
4,109.7 µM in liquid cultures of two Streptomyces reservoir isolates. Uwins (2011) reported that significantly more geosmin was produced in liquid cultures of S. coelicolor
A3(2) when nitrogen concentration was reduced (particularly at 74.40 µM and 744.0 µM) compared to when this nutrient was abundant (24,987.5 µM). The author also identified that adenosine tri-phoshphate (ATP) concentration, an indicator of metabolic activity, was aligned with geosmin production (as opposed to biomass production), suggesting that under lower nitrogen conditions, cellular energy was geared toward production of geosmin rather than cell division (Uwins, 2011). Blevins et al. (1995) also observed more geosmin production in nitrogen reduced liquid cultures of S. halstedii, with low concentrations (0.6-1.2 µM) favouring maximal production compared to higher concentrations (up to 93.7 µM). Furthermore, over the range of 4.7 µM to 374.1 µM of nitrogen, Weete et al. (1977) observed that lower concentrations supported greater geosmin production by a Streptomyces reservoir isolate, similar to the low concentrations found to support greatest odour production by both Streptomyces spp. in the present study.
Greater production of geosmin under nitrogen limiting conditions has also been reported in studies examining cyanobacterial spp. (Naes et al., 1988; Wu et al., 1991; Saadoun et al., 2001). 241
A commonality among the aforementioned studies is that they are aligned with respect to higher nitrogen concentration favouring more biomass development. What is not clear from existing data is whether geosmin and 2-MIB production is triggered by low nitrogen concentrations or triggered by other factors and simply inhibited by higher nitrogen concentrations. Production of numerous other secondary metabolites have been shown to be elicited under nitrogen limitation (van Wezel & McDowall, 2011).
In a molecular context, it has been shown that PhoP binds to and reduces transcription from the promoter of glnR, which encodes a major transcriptional regulator of nitrogen metabolism in addition to genes encoding an ammonium transporter, two glutamine synthetases (which assimilate ammonium into primary metabolism) and two post- transcriptional regulators. PhoP represses genes for nitrogen assimilation while activating genes for phosphate assimilation when phosphate is limiting, as there is a reduced demand for nitrogen and thus the continued expression of genes for nitrogen assimilation would be wasteful. Thus phosphorus depletion alone may induce secondary metabolism irrespective of the abundance of nitrogen. It is also known that genes for nitrogen assimilation can be repressed independently of activation of PhoP (van Wezel &
McDowall, 2011). The role of nitrogen in the production of geosmin and 2-MIB is unclear, but is worthy of further investigation.
Overall, the results indicate that control of geosmin and 2-MIB is a multifunctional process in which limiting nutrients, particularly nitrogen and phosphorous play important roles. Carbon, nitrogen and phosphorous, play an important role in the regulation of primary and secondary metabolism in all bacteria and while the different mechanisms of control have been widely studied in an individual context, connections between them have not been investigated (van Wezel & McDowall, 2011). This is the first study that has investigated how different combinations of carbon, nitrogen and phosphorous
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concentration might influence growth and T&O metabolite biosynthesis by Streptomyces, and the findings clearly support the significance of nutrient limiting conditions in the production of geosmin and 2-MIB by these organisms, which is highly coordinated with the process of morphological differentiation.
In order to understand the implications of these experimental results in terms of the potential for Streptomyces to produce geosmin and 2-MIB in drinking water reservoirs, physico-chemical parameters measured in the field need to be taken into consideration.
In this case, the historical water quality data of Grahamstown and Chichester Reservoirs were considered. Average concentrations of total phosphorous measured in the surface waters of both reservoirs, 0.43 (0.05-4.2) µM and 0.78 (0-45.2.0) µM respectively for
Grahamstown and Chichester Reservoirs are of similar concentration to the levels found to stimulate greatest geosmin and 2-MIB production by Streptomyces cultures (1.6 µM).
In addition, the lowest concentration of nitrogen (3.6 µM) in this study which also supported maximal T&O metabolite biosynthesis, are similar to average dissolved inorganic nitrogen levels measured in Grahamstown and Chichester Reservoirs, being
2.62 (0.32-21.6) µM and 4.39 (0.72-32.13) µM respectively. Total organic carbon average concentrations in Grahamstown and Chichester Reservoir waters are 0.60 (0.04-
1.1) mM and 0.25 (0.04-1.61) mM respectively, similar to the lowest carbon concentration provided in the growth media (0.42 mM). In general, it can be concluded that the low phosphorous, nitrogen and carbon concentrations typically found in the reservoir waters are indeed suitable for supporting production of geosmin and 2-MIB by
Streptomyces based on these laboratory results.
6.4.3 Other physico-chemical parameters in relation to geosmin and 2-MIB production by Streptomyces spp.
Beyond nutrient (carbon, nitrogen and phosphorous) limitation, other physico-chemical parameters that were examined in the subsequent multifactorial experiments revealed 243
stimulatory or inhibitory effects on the production of T&O metabolites by the
Streptomyces spp. which are discussed in the following sections. While these effects were also largely tied to the extent of morphological differentiation exhibited by the cultures, this was not always apparent, suggesting that some factors may influence the production of T&O metabolites independent of the extent of morphological differentiation.
6.4.3.1 Physico-chemical factors inhibiting T&O metabolite production by Streptomyces spp.
Increasing the concentration of NaCl was considered likely to inhibit the vegetative growth of Streptomyces spp. and rather, be associated with morphological differentiation corresponding to enhanced production of geosmin and 2-MIB. Contrary to expectations was the significant inhibitory influence of elevated NaCl concentration on production of both odorous compounds, irrespective of the temperature and pH level in the second multifactorial experiment. Increasing the NaCl concentration was found to enhance the growth of the Streptomyces, with the observation of predominately vegetative growth at high NaCl levels (Figures 6.30 and 6.31). High salt tolerance is a documented characteristic of Streptomyces (Tresner et al., 1968) and Sakr et al. (2013) reported that optimum NaCl concentrations for the growth of Streptomyces strains to be between 2.5% to 10%. In contrast, low biomass, highly differentiated mycelium and free spores were observed in the cultures exposed to low NaCl concentration, from which significantly greater yields of geosmin and 2-MIB were recovered. Supporting these findings, Rezanka and Votruba (1998) found that increasing the NaCl concentration up to 0.5% stimulated geosmin production in liquid cultures of S. avermitilis, whereas concentrations above this significantly reduced production (inhibited above 2% NaCl). The authors made no reference to the effects on biomass. Sevckova and Kormanec (2004) reported that production of pigmented antibiotics by S. coelicolor A3(2) were differentially affected by high salt concentration, with production of actinorhodin inhibited whilst production of
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undecylprodigiosin was activated in solid cultures that were salt stressed (2.5% NaCl).
The authors found that this effect was mediated by differential expression at high salt concentrations of genes encoding pathway specific transcription regulators for biosynthesis of these secondary metabolites. The expression of genes associated with geosmin and 2-MIB biosynthesis and/or morphological differentiation may also be repressed under conditions of high salt concentration.
Based on the highest electrical conductivity (EC) values measured in Chichester and
Grahamstown Reservoirs (approximately 350 µS/cm), concentrations of NaCl (if this compound was to be the only contributor to the measurement of EC) would not exceed
0.02% w/v, two-orders of magnitude lower than the maximal concentration used in this study (2%). Thus the findings imply that salinity levels typical of drinking water storages would be favourable for Streptomyces production of T&O metabolites as opposed to waters that are more brackish.
A parameter which undoubtedly controls the metabolic activity of Streptomyces is temperature. Being predominately mesophilic, these organisms generally exhibit optimum growth between 25°C to 30°C (Goodfellow & Williams, 1983). Over the range investigated in this study (24°C to 30°C), the lower temperature level generally favoured higher production of geosmin and 2-MIB by Streptomyces spp. Similarly, Yagi et al.
(1987) and Weete et al. (1977) reported that the optimal temperature for geosmin and 2-
MIB production by Streptomyces isolates was 25°C. Numerous other studies have reported that higher temperatures support greater production of these metabolites by
Streptomyces spp. (between 30 - 40°C) (Dionigi & Ingram, 1994; Blevins et al., 1995;
Uwins, 2011). An important point of consideration in this study is the use of solid as opposed to liquid cultures and the likelihood that greater physical loss of the compounds to the atmosphere occurred at the higher temperature tested due to their high volatility.
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Indeed, preliminary testing revealed that the T&O metabolites could not be recovered from cultures of both Streptomyces spp. when incubated at 37°C. It is plausible to conclude that temperatures over the range examined exhibited little influence on geosmin and 2-MIB production, particularly considering that similar extents of morphological differentiation were observed across the temperature dimension in these experiments
(Figures 6.30 and 6.31). Nonetheless, the lower temperature tested (24°C) that appeared to support greater geosmin and 2-MIB production in this study is more representative of levels encountered in reservoirs during warmer months and thus Streptomyces could indeed be contributing to the presence of T&O compounds in the water column. Although average temperatures in Grahamstown and Chichester Reservoir surface waters are
19.8°C and 18.1°C respectively, temperatures of up to 27.8°C and 30.1°C respectively have been recorded.
Consistent with the effects of the major macronutrients (carbon, nitrogen and phosphorous), increasing the magnesium concentration (1000 µM) significantly reduced geosmin and 2-MIB production by both Streptomyces spp. independent of the calcium and potassium concentration. This however, did not correspond to life cycle stage as all cultures in the experiment exhibited high degrees of morphological differentiation. Yagi,
Sugiura and Sudo (1987) investigated the effect of magnesium concentration (0-480.0
µM) on the production of T&O metabolites in liquid cultures of two Streptomyces reservoir isolates. For one isolate, magnesium had no effect on geosmin production, whilst maximal 2-MIB production occurred in the low concentration of 19.9 µM and dramatically decreased with exposure to higher concentrations. For the other isolate, maximal production of both compounds occurred in the presence of a magnesium concentration between 79.5 µM to 159.0 µM, with higher concentrations decreasing production. These results concur with the findings in this study, with lower magnesium concentration favouring production of geosmin and 2-MIB. Thus increasing magnesium
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concentration appears to be less favourable for T&O secondary metabolite production despite (as indicated in Chapter 2) this cation being a cofactor required for the enzymes which are responsible for geosmin and 2-MIB biosynthesis (Cane et al., 2006; Jiang et al., 2007). It may be that limiting concentrations of magnesium act as a regulatory signal to simulate expression of genes involved the biosynthetic pathways of these compounds.
Gram negative bacteria including Salmonella and Pseudomonas spp. are known to use extracellular magnesium concentration as a primary signal to regulate the activity of the
PhoP-PhoQ system which governs virulence and regulates numerous other cellular activities, with low concentrations of magnesium promoting transcription of PhoP- activated genes (Groisman, 2001).
Micronutrient concentration is known to greatly influence secondary metabolite production in Streptomyces (van Wezel & McDowall, 2011). Based on previous literature reports however, it was not clear what effect such factors would have on geosmin and 2-
MIB production by Streptomyces. The final factorial experiment conducted examined the influence of four micronutrients (zinc, iron, manganese and copper) over the concentration range of 0.25 µM to 25 µM on geosmin and 2-MIB production by the
Streptomyces spp. The most profound result was the strong negative effect of increasing copper concentration on production of both T&O metabolites. This aligns with the results reported by Schrader and Blevins (2001), who observed reduced production of geosmin in liquid cultures of S. halstedii over the range 0 µM to16 µM, with a maximal yield occurring at a concentration of 0.54 µM whilst complete inhibition of geosmin and biomass production occurred at copper concentrations exceeding 10.7 µM. Contradicting these experimental results, Dionigi, Ahten and Wartelle (1996) reported that solid cultures of S. tendae exposed to 15.7 µM and 78.7 µM of copper accumulated more biomass, exhibited higher degrees of sporulation and six-fold greater geosmin production compared to controls and cultures treated with other metals. Although the biomass produced was not greatly influenced by copper in this study, in cultures exposed to highest 247
concentration of this micronutrient, vegetative growth was predominately observed.
Conversely, those subjected to low copper concentrations generally exhibited higher degrees of morphological differentiation (Figures 6.32 and 6.33) aligning with the higher
T&O metabolites recovered. This reaffirmed the strong relationship between T&O metabolite production and morphological differentiation.
In relation to other micronutrients, iron, zinc and manganese exhibited comparatively lesser effects on the production of geosmin and 2-MIB by the Streptomyces spp. Previous reports have observed variable effects of iron concentration on T&O metabolites production by Streptomyces including stimulatory effects (Uwins, 2011) as was generally observed to a small extent in the present study, inhibitory effects (Schrader & Blevins,
2001) and similar yields produced over a broad range of concentrations (0-144.5 µM) as reported by Yagi et al. (1987). Collectively, these findings exemplify the relatively insignificant effect this micronutrient appears to have on T&O metabolite biosynthesis by Streptomyces. Although to a much lesser extent than copper, increasing zinc concentration generally reduced yields of both metabolites, consistent with the observations made by Schrader and Blevins (2001) and Dionigi, Ahten and Wartelle
(1996). In relation to manganese, the results support previous findings which indicate that this micronutrient is not essential for and/or does not significantly influence earthy-musty secondary metabolite production by Streptomyces (Schrader & Blevins, 2001; Uwins,
2011).
Although four micronutrients were investigated in this study, field data was only available for manganese and iron concentrations at Chichester Reservoir. Average values of manganese and iron in the reservoir surface waters are 0.34 (0-6.67) µM and 6.32 (0.018-
181.9) µM respectively which are similar to the range used in this study (0.25 µM to 25
µM), and thus are not likely to influence Streptomyces production of geosmin and 2-MIB
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under natural settings. Considering that the trigger values of copper and zinc concentration specified in the ANZECC (Australian and New Zealand Environment and
Conservation Council) guidelines for lakes and reservoirs are 0.02 µM to 0.04 µM and
0.03 µM to 0.18 µM respectively, these micronutrients are highly unlikely to influence geosmin and 2-MIB production by Streptomyces in reservoirs (ANZECC & ARMCANZ,
2000).
6.4.3.2 Physico-chemical factors stimulating T&O metabolite production by
Streptomyces spp.
The preference for neutral environmental pH for the growth of Streptomyces is well known, yet they are capable of growing over a broad range of pH (Goodfellow &
Williams, 1983). Indeed soil, being their most common habitat, has an average pH range of 3.5 to 6.8 (Kontro et al., 2005). In relation to geosmin and 2-MIB production, it was considered that by increasing the pH (beyond the neutral range optimal for vegetative growth), greater yields would be produced. The results did concur with this expectation, with T&O metabolite production favoured under more alkaline (pH 9) conditions.
However the effect of pH appeared to be independent of the extent of morphological differentiation exhibited by the cultures. That is, the extent of morphological differentiation was similar across the pH range under the same levels of NaCl and temperature (Figures 6.30 and 6.31). This is supported by the findings of Kontro et al.
(2005), where pH (over the range 4.5 to 11.5) had a relatively small effect on the sporulation of numerous Streptomyces spp. grown on different laboratory media. Results documented in the literature support the positive effect of pH on T&O metabolite production by Streptomyces spp., with values of pH 9 to 10 reported to maximise production whereas conditions of neutrality have generally been found to be more favourable for growth (Weete et al., 1977; Yagi et al., 1987; Blevins et al., 1995).
Grahamstown and Chichester Reservoirs generally have neutral pH levels, with average
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values of 7.5 (6.5-8.2) and 6.7 (6.2-7.7) respectively. The typical pH levels in the reservoirs would be suitable for T&O metabolite production by Streptomyces, but are not alkaline enough to stimulate maximal amounts as demonstrated in this study.
Being essential for vegetative growth, macronutrients at their lowest concentration were predicted to be associated with enhanced production of T&O metabolites by
Streptomyces. While this hypothesis was supported in the case of carbon, nitrogen, phosphorous and additionally, magnesium, higher levels of calcium and potassium over the concentration range of 1 µM to 1000 µM were found to maximise production of both secondary metabolites by S. coelicolor A3(2) and S. antibioticus. The positive effect of calcium contradicts the results reported by Schrader and Blevins (2001), who observed that maximal geosmin and biomass occurred in the absence of calcium. Daza et al. (1989) observed that increasing the concentration of calcium in the growth medium increased sporulation by a Streptomyces sp. due to the trapping of phosphate ions as insoluble calcium phosphate, reducing phosphorous levels. Thus the reduction of bioavailable phosphorous by calcium ions may explain the greater production of geosmin and 2-MIB under conditions of higher calcium concentration. Similar to calcium, potassium exhibited a positive effect on geosmin and 2-MIB production by both species which contrasts to the finding of Schrader and Blevins (2001) that potassium had no significant effect geosmin production by S. halstedii, however the concentration range was lower (0-
3.2 µM) compared to this study (1-1000 µM).
Concentrations of these macronutrients have not been monitored in Grahamstown and
Chichester Reservoirs. However, the range of concentrations examined in this study cover those typically encountered in freshwater environments. Given that appreciable quantities of both secondary metabolites were produced under all combinations of concentrations of these macronutrients (including magnesium), fluctuating levels of these factors in
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freshwater environments are unlikely to play a significant role in enhancing or inhibiting production of geosmin and 2-MIB by Streptomyces. Indeed under all combinations of calcium, potassium and magnesium concentrations in this experiment, cultures exhibited a high degree of differentiation, with heavy sporulation observed, suggesting changes in these factors over the concentration ranges examined do not greatly alter secondary metabolism and differentiation.
6.5 Conclusions
Evidence has been acquired in this study to support the notion that the production of geosmin and 2-MIB is highly coordinated with life cycle stage of Streptomyces, being greatest under the conditions which trigger morphological differentiation and sporulation in the organisms. Such conditions included nutrient limitations, the combination of low nitrogen and phosphorous concentration, in addition to low carbon concentration in the case of S. coelicolor A3(2). Conversely, the combination of highest concentration of these macronutrients demonstrated an inhibitory effect on T&O metabolite production, favouring the vegetative growth of the organisms. Generally the effect of changes in the concentration of other macronutrients (calcium, potassium and magnesium) on geosmin and 2-MIB production by the Streptomyces spp. was less significant, exemplified by the similar high degree of morphological differentiation that occurred in cultures subjected to varying concentrations of these factors. Of all micronutrients tested, increasing the concentration of copper proved to have a significant inhibitory effect on the production of geosmin and 2-MIB and stimulated vegetative growth. This inhibitory effect on T&O metabolite production was also apparent in the case of high NaCl concentration, which also favoured vegetative growth of the organisms. An alkaline environment and lower temperature were found to be more favourable for geosmin and 2-MIB production by the
Streptomyces spp., although changes in the levels of these parameters did not affect the extent of morphological differentiation observed.
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In consideration of these results, the levels of physico-chemical characteristics in the surface waters of Grahamstown and Chichester Reservoirs appear to be suitable for
Streptomyces growth, eventual morphological differentiation and consequently, the production of geosmin and 2-MIB. These laboratory studies were conducted using solid media, which as opposed to liquid media, would not be very representative of reservoirs.
However, the fact that both Streptomyces spp. were capable of growing and producing earthy-musty secondary metabolites in sterilised reservoir water (Chapter 4) suggests that they indeed have the potential to produce T&O compounds in the reservoirs.
This study has demonstrated how the application of a full factorial experiment can lead to a systematic overview of the influence of environmental parameters on geosmin and
2-MIB production by Streptomyces. However, it is inherently challenging to relate the results of controlled, pure culture laboratory studies to the manner in which the microorganism may respond in the environment, where uncontrolled and potentially unmeasured, complex interactions occur to influence metabolic processes. Yet the more data that is gathered regarding how an organism’s metabolic processes are influenced by different factors in controlled experimental settings, the more information there is available to assist in deciphering and determining the influence of complex in situ interactions.
The pure culture experiments undertaken in this study attempted to examine interactions of different environmental factors by utilising a factorial experimental design. This differs from previous work published on the influence of factors on geosmin and 2-MIB production by Streptomyces which have examined environmental factors independently.
Future studies should aim to examine the effects of environmental variables simultaneously using chemically defined media and progress from this study by performing a single experiment with a larger number of factors rather than multiple
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factorial experiments examining only three to four factors simultaneously. With the two- level full factorial design, increasing the number of factors beyond four or five dramatically increases the number of experiments, so that only two to four factors are experimentally practical and economically defendable choices. To examine more than four factors, a switch to fractional factorial design is favourable (Eriksson et al., 2008).
This would allow for a wide range of conditions potentially encountered in the environment to be examined in relation to geosmin and 2-MIB production by
Streptomyces. Indeed, the ultimate aim of future studies should be to explore and develop options to control in situ production of these earthy-musty secondary metabolites.
Production of geosmin and 2-MIB is clearly favoured under the conditions which also trigger morphological differentiation and sporulation. This suggests that their possible biological function is related to the reproductive phase of the Streptomyces life cycle, possibly serving as a defence mechanism similar to many other secondary metabolites, to antagonise competing microorganisms in times of adversity (e.g. nutrient limitation).
Such a hypothesis concerning the biological role of geosmin and 2-MIB in the life of
Streptomyces is explored in the next chapter.
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CHAPTER 7 - BIOSYNTHESIS OF GEOSMIN AND 2-METHYLISOBORNEOL BY STREPTOMYCES: WHY?
7.1 Introduction
To date the water industry has tended to treat geosmin and 2-MIB, along with other volatile organic compounds (VOCs), as metabolic by-products. With a view to identifying potential means to control the accumulation of these compounds, research has predominately been focused on their biological sources, triggers of production in drinking water storages and their removal from drinking water. In spite of this research, T&O outbreaks involving these compounds remain highly unpredictable (Watson, 2003; Zaitlin
& Watson, 2006; Juttner & Watson, 2007).
Despite being ubiquitous microbial secondary metabolites in soil environments and the main source of soil odour, comparatively little attention has been paid to the biological purpose served by production of geosmin and 2-MIB by microorganisms. That is, among the existing literature there appears to be little consideration of the functional roles of these metabolites. The purpose of production, if any, of geosmin and 2-MIB by
Streptomyces and indeed other producing organisms, remains unclear. The research presented in this chapter represents an attempt to gain a further understanding of the functional role of these earthy-musty secondary metabolites in the life of these bacteria.
Previous research has focused on determining the toxicity and mutagenicity of geosmin and 2-MIB to a diverse range of test organisms, predominately to gauge the threat they pose to humans regarding the consumption of water contaminated with these compounds.
In vitro assays have demonstrated that they can reduce cell viability and damage DNA in rainbow trout hepatocytes and are toxic, but not mutagenic to sea urchin embryos using concentrations in parts per million (0.45- 100 mg/L) (Nakajima et al., 1996; Gagne et al.,
1999). Over the extensive concentration range of 10 ng/L to 300 mg/L, geosmin and 2- 254
MIB were found to exhibit no cytotoxic effects against cultured mammalian cells (human, monkey and dog cells) (Mochida, 2009). More recently, Burgos et al. (2014) assessed the genotoxicity of 2-MIB and demonstrated using an in vivo test system (SMART) that 2-
MIB was unable to induce gene and chromosome mutations or events associated with mitotic recombinations in Drosophila melanogaster (concentrations up to 500 mg/L). The authors also demonstrated through in vitro assays that 2-MIB is not associated with chromosomal instability in Chinese Hamster Ovary (CHO) cells. Together, these studies suggested that apart from some isolated exceptions involving relatively substantial concentrations, geosmin and 2-MIB are largely non-toxic to higher-order organisms.
Some authors have suggested that geosmin and 2-MIB may exhibit an antimicrobial activity similar to that reported for other terpene derived alcohols (Dionigi et al., 1993;
Watson, 2003). However, the excessively high concentrations used in toxicity studies far exceed those levels normally encountered in soil and freshwater environments.
Nevertheless, concentrations where they may exhibit antimicrobial activity may be encountered by microorganisms in close proximity to the producing population or at the micro-scale level in soil, sediments, and biofilm matrices (Watson, 2003; Zaitlin &
Watson, 2006).
While the biological functions of many of the secondary metabolites such as geosmin and
2-MIB produced by Streptomyces are not known, it seems inconceivable that they do not play an adaptive ecological role. It is difficult to reconcile them as waste products or shunt metabolites, given that they have complex underlying genetic information and are biosynthesised by many separate enzymatic steps through pathways which are energetically expensive. It therefore follows that their production must be important to the survival of the organisms in their natural habitat. In fact, although regarded as non- essential compared to primary metabolites required for cell growth, secondary
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metabolites are often considered molecules of adaptation, intimately linked with the ecology of the producing organisms, that have evolved for a purpose other than primary metabolism (O’Brien & Wright, 2011). That is, the producing microorganism has retained the capacity to biosynthesise secondary metabolites due to selective advantages conferred by the functions of those compounds (Maplestone et al., 1992; Marinelli & Marcone,
2011). In the case of geosmin and 2-MIB production by Streptomyces, these functions are yet to be established.
Microbial populations in nature consist of complex mixtures of different species which interact with and respond to each other through processes such as competition, predation, parasitism, symbiosis and neutralism. It is these microbial interactions that are thought to be the driving force for the production of many secondary metabolites. Soil in particular is inhabited by an immense variety of microbial species, all in competition for the limited nutrient resources available in this nutritionally heterogeneous environment (Tarkka &
Hampp, 2008). Streptomyces are ubiquitous in soil and as saprophytes, play an important role in nutrient recycling. Upon nutrient and moisture availability, their spores germinate, growing by tip extension to form a dense vegetative mycelium. In response to nutrient depletion, Streptomyces enter the reproductive stage of their complex life cycle. It is at this stage, the transition from vegetative growth to the initiation of the aerial mycelium which differentiates into chains of spores, that the vast array of secondary metabolites that this genus is capable of biosynthesising are produced (Bibb, 2005; Hopwood, 2007).
This was demonstrated in Chapter 6, with low concentrations of nutrients including carbon, nitrogen and phosphorous stimulating maximal geosmin and 2-MIB production and morphological differentiation in solid cultures of Streptomyces spp.
Considering that secondary metabolites are produced by sessile Streptomyces as the secondary mycelium develops at the expense of nutrients released by the breakdown of
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vegetative cells, it has been proposed that these small molecules are produced to defend this nutrient supply from competitors (Marinelli & Marcone, 2011). Thus, the competition hypothesis contends that the antimicrobial activities of secondary metabolites provide the producing organism a selective advantage over competitors in nutritionally poor environments (Maplestone et al., 1992). Indeed, since the 1940s, Streptomyces have been utilised for their production of an impressive variety of secondary metabolites which have antibacterial, antifungal and other biological activities. They are a key genus in many industrial applications, with over two-thirds of all known antibiotics produced by
Streptomyces (Hopwood, 2007). Perhaps however, the focus on identification of secondary metabolites for their exploitation as therapies for human diseases, rather than determination of their actual function in the producing organism, has led to oversight of possible roles aside from that as competitive weapons of inter-microbial chemical warfare
(O’Brien & Wright, 2011).
Nevertheless, in this study it was hypothesised that, similar to many other Streptomyces secondary metabolites that have been characterised, geosmin and 2-MIB are biosynthesised as antimicrobial compounds to antagonise competing microorganisms in their natural soil environment. Indeed, at respective concentrations of 45.2 mg/L and 18.1 mg/L, geosmin and 2-MIB were shown by Dionigi et al. (1993) to inhibit the growth of the bacterium Salmonella typhimurium, suggesting high concentrations of these compounds can exhibit antimicrobial activity. The antifungal activity of VOCs extracted from cultures of Streptomyces alboflavus, of which 2-MIB and geosmin were particularly abundant (51.34% and 4.28% of total VOCs respectively), was also recently reported
(Wang et al., 2013a). In a subsequent study (Wang et al., 2013b), these same authors directly demonstrated that 2-MIB, the most abundant VOC, exhibited significant inhibitory effects on the mycelia growth and sporulation by the fungus Fusarium moniliforme by fumigation (at concentrations exceeding 10 µg per plate).
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Contrary to these findings, Schulz et al. (2010) reported that among other bacterial volatile compounds in agar diffusion assays, geosmin and 2-MIB (at a high concentration of 20 µg/µL) were not inhibitory against selected test microorganisms (Aspergillus fumigatus, Botrytis cinerea, Candida albicans, Pythium debaryanum, Hansenula anomala, Saccharomyces cerevisiae, Escherichia coli, Pseudomonas aeruginosa,
Klebsellia pneumonia, Staphylococcus aureus, Micrococcus luteus and Mycobacterium phlei). No other identified reports have investigated the antimicrobial effects of geosmin and 2-MIB and given the contradictory results reported, this possible biological function remains inconclusive.
In the laboratory, it is common procedure to study bacteria such as Streptomyces in pure cultures, which in the absence of interacting microorganisms that are present in the natural microbial environment, masks the real biosynthetic potential of the organism under investigation (Onaka et al., 2011). In nature, it is not common to find one type of microorganism growing in isolation from others. Inter-species interactions through co- culturing may induce the unexpressed or enhance the expressed biosynthetic pathways of some compounds compared to growth as a mono-culture. This would be true for those secondary metabolites that function as antimicrobial compounds. A series of studies by
Luti and Mavituma (2011a, 2011b, 2011c) demonstrated the effect of interaction of
Streptomyces coelicolor A3(2) with other bacteria on enhancing the production of the antibiotic undecylprodigiosin, as compared to pure cultures. When interacted with live or heat-killed cells of Bacillus subtilis, liquid cultures of S. coelicolor A3(2) increased production of undecylprodigiosin by up to 256% in comparison to pure cultures of this strain (Luti & Mavituma, 2011c). In another study, pure cultures of S. coelicolor A3(2) produced higher concentrations of actinorhodin compared with undecylprodigiosin, but elicitation with E.coli altered this pattern, such that undecylprodigiosin production, a more potent antibiotic was enhanced, while actinorhodin was repressed (Luti &
Mavituma, 2011b). Furthermore, introducing heat-killed cells of B. subtilis and 258
Staphylococcus aureus to S. coelicolor A3(2) cultures stimulated an earlier onset of production and higher yields of undecylprodigiosin by 3-fold and 5-fold respectively compared to pure cultures (Luti & Mavituma, 2011a). The interaction between the predatory myxobacterium Myxococcus xanthus with S. coelicolor A3(2) in liquid co- cultures also increased production of actinorhodin by up to 20-fold compared to pure cultures (Perez et al., 2011). The direct cell-to-cell interaction between Streptomyces spp. and mycolic acid-containing bacteria induced secondary metabolism, with a novel antibiotic (alchivemycin A) produced by Streptomyces endus S-522 when co-cultured with Tsukamurella pulmonis (Onaka et al., 2011).
The use of interspecies interaction for enhancing secondary metabolite production has been demonstrated with other microbial taxa, such as the production of a new antibiotic, pestalone by liquid cultures of a marine fungus (Pestalotia sp.) when challenged with a mixture of marine bacteria (Cueto et al., 2001). Rateb et al. (2013) observed that by co- culturing Aspergillus fumigatus with Streptomyces bullii led to the isolation of diverse secondary metabolites that could not be detected in pure cultures of either microorganism.
Are geosmin and 2-MIB over- or under-expressed by Streptomyces under unnatural mono-culture conditions used in the laboratory? Such a phenomenon has not previously been investigated in relation to these metabolites. To address the hypothesis concerning the possibility that Streptomyces production of geosmin and 2-MIB serve an antagonistic purpose as antimicrobial compounds, a co-culturing protocol was adopted to partially mimic inter-microbial competition found in nature. For the hypothesis to hold true, it was expected that the interaction between Streptomyces with other diverse microorganisms, some of which are common soil inhabitants, would stimulate greater production of geosmin and 2-MIB compared to the absence of inter-microbial competition under pure culture conditions. To further test the hypothesis, the antimicrobial effects of these
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secondary metabolites were screened against a range of bacterial and fungal species using the agar diffusion method with pure analytical standards of these compounds.
7.2 Materials and methods
7.2.1 Microorganisms
The geosmin and 2-MIB producing Streptomyces spp. employed in earlier experimental work in Chapters 4, 5 and 6 including S. coelicolor A3(2) (ATCC BAA-471) and the sediment isolate S. antibioticus, were used in the present study. Working Streptomyces cultures for the co-culture experiments were maintained on SCA incubated at 28°C.
A diverse range of microorganisms were employed to interact with Streptomyces spp. under co-culturing conditions including Gram-negative bacteria (Pseudomonas fluorescens, Serratia marscesens and Escherichia coli), Gram-positive bacteria (Bacillus subtilis, Micrococcus luteus and Staphylococcus aureus) in addition to two fungi
(Penicillium notatum and an unidentified fungal isolate recovered from a soil sample also exhibiting Penicillium-like characteristics). These test microorganisms were also used in the assays testing the antimicrobial effects of geosmin and 2-MIB. S. aureus (ATCC
29213) was purchased from ATCC and the other microbes were obtained from stock culture collections maintained in The School of Environmental and Life Sciences at the
University of Newcastle. Stock cultures of these bacteria and fungi were maintained on nutrient agar (NA) (Oxoid) and refrigerated at 4°C. Prior to use, all microorganisms were tested for geosmin and 2-MIB production ability on NA and confirmed as non-producers of both secondary metabolites. Despite some species in the Penicillium genus being implicated as geosmin producers, this metabolite could not be detected in the culture extract of this particular species.
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7.2.2 Co-culture experiments: the effect of interaction with other microorganisms on the production of geosmin and 2-MIB by Streptomyces spp.
A co-culturing protocol was developed similar to the method described by Charusanti et al. (2012) to investigate how Streptomyces interaction with other microorganisms on solid medium influences the production of geosmin and 2-MIB. NA was employed as the growth medium in this investigation as preliminary testing revealed its suitability for supporting profuse growth of all test microorganisms and both Streptomyces spp. A schematic overview of the methodology developed is summarised in Figure 7.1. Using a sterile cotton tip, each of the test bacteria and fungi were spread over the surface of NA in polystyrene Petri dishes (60 mm x 10 mm) to establish dense lawns of biomass. A space approximately 2 cm in diameter in the centre of each plate where the test microorganisms were not spread was established to enable the growth of the Streptomyces spp. to occur without being overgrown, but to still allow direct contact between the
Streptomyces and test microorganism. An inoculum (250 µL) containing spores for each
Streptomyces spp. was carefully deposited onto the remaining exposed agar surface at the center. Once the inoculum had dried, the plates were inverted and incubated at 28°C for 5 days. The temperature was selected based on preliminary assessment of its suitability for the growth of the diverse range of microorganisms utilised in this study over the incubation period. Each Streptomyces spp. was co-cultured with all test microorganisms in triplicate in addition to three replicates of a control, containing only the Streptomyces inoculum (pure culture).
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Figure 7.1 Co-culturing scheme developed to test the effect of Streptomyces spp. interaction with other microorganisms on the production of geosmin and 2-MIB.
Following the incubation period, geosmin and 2-MIB were extracted from all co-cultures and the controls using the hexane extraction protocol described in Chapter 4. The culture extracts were supplemented with internal standard (biphenyl-d10) and BSTFA, analysed by GC-MS and quantified as previously described (4.2.6). The production was expressed as ng per culture. The absence of biomass measurements was due to the difficulty in separation of Streptomyces biomass from that of the competing organism.
In order to determine the extent of morphological differentiation, Streptomyces cultures were examined with light microscopy (1000x magnification) after Gram-staining and were assigned differentiation values ranging from 0 (vegetative mycelium only) to 5
(highly differentiated, biomass >50% spores) according to the criteria developed in
Chapter 6 (Table 6.1). The relationship between T&O metabolite production and differentiation was assessed. Following the incubation period, images of all plates were taken with a digital camera.
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7.2.3 Antimicrobial assays of geosmin and 2-MIB
To further examine whether the function of geosmin and 2-MIB may be to antagonise competing organisms, the antimicrobial (antibacterial and antifungal) effects of these compounds were assessed. Analytical standards of geosmin (2 mg/mL) and 2-MIB (10 mg/mL) dissolved in methanol (MeOH) were purchased from Sigma-Aldrich. The standards were diluted in MeOH to concentrations of 1 mg/mL to use in antimicrobial assays.
The agar diffusion method using agar plugs was used to examine the antimicrobial activity of these compounds. A concentration of 35 µg/mL of each compound, similar to those employed in previous toxicity tests of geosmin and 2-MIB was used. Such a concentration of these compounds would greatly exceed those normally observed in water and soil, but conceivably would represent concentrations encountered by organisms in close proximity to the producing Streptomyces population at the micro-scale level in soil or other substrates. Solid media containing geosmin and 2-MIB were prepared by inoculating 350 µL of the working solutions (1 mg/mL) into 9.65 mL of autoclaved molten water agar (1.5%) cooled down to 50°C. Additionally, a medium was prepared containing both compounds. The molten water agar containing the compounds were distributed (10 mL) to polystyrene Petri dishes (90 mm x 10 mm) and immediately cooled at 4°C for 10 minutes to minimise loss of the compounds through volatilisation and to increase the firmness of the agar for ease in removing cores.
The diverse range of bacterial and fungal species used in the co-culture study was employed to test the potential scope of antimicrobial activities of geosmin and 2-MIB.
Single colonies of the microorganisms growing on NA were transferred and spread over the surface of fresh NA plates with a sterile cotton tip to establish dense growth. Agar wells were created in each plate using an alcohol-sterilised metal cork-borer (10 mm
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diameter). Agar cores were cut with the borer from the water agar plates containing geosmin, 2-MIB and both compounds and were carefully removed aseptically with forceps and transferred to the NA wells (in duplicate). Plates were then incubated at 28°C.
The antimicrobial effect of geosmin, 2-MIB and both compounds together, was assessed by examination of a zone of inhibition around the introduced agar plugs. Plates were examined after 24 and 48 hours for the formation of a zone of inhibition. The absence of a zone of inhibition indicated a negative result. As a control, water agar plugs containing no standards (only MeOH) were used. A schematic overview of the protocol described is detailed in Figure 7.2.
Figure 7.2 Schematic of the protocol developed for testing the antimicrobial (antibacterial and antifungal) effects of geosmin, 2-MIB and the combination of both compounds against the test microorganisms.
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7.2.4 Statistical analysis
The statistical significance of the data from the co-culturing experiment at a 95% confidence level was assessed by conducting one way ANOVA followed by Tukey HSD test for post hoc pairwise multiple comparison using the statistical program JMP version
7.0. Regression analyses were used to determine the relationship between earthy-musty secondary metabolite yields and differentiation using Microsoft Office Excel 2013.
7.3 Results
In this study, the hypothesis that geosmin and 2-MIB are produced by Streptomyces to function as antimicrobial compounds was tested through a combination of direct analyses using analytical standards of both compounds to assess their antimicrobial capacity, and by examining the effects of interaction of Streptomyces spp. with other microorganisms on the production of these secondary metabolites.
7.3.1 The effect of interaction with other microorganisms on the production of geosmin and 2-MIB by Streptomyces spp.
Geosmin and 2-MIB production by both Streptomyces spp. was examined on NA under pure culture conditions and co-culturing conditions to determine the influence of interaction with other microorganisms on the production of these secondary metabolites.
Figures 7.3 and 7.4 show the yields of geosmin and 2-MIB produced by S. coelicolor
A3(2) and S. antibioticus respectively under these conditions. Production of each compound is expressed as ng per culture. Overall, the major trend depicted is that pure cultures produced greater yields of both secondary metabolites, contrary to the hypothesis that Streptomyces interaction with other microorganisms, partially representing the situation of inter-microbial competition encountered in natural settings, would stimulate greater production of geosmin and 2-MIB.
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600 Geosmin A 500
400
300 ng/culture 200 B BC BCD 100 BCD BCD BCD CD D 0
60 2-MIB 50 A
40
30 B
ng/culture 20 C C C C C 10 C C 0
Figure 7.3 Geosmin (top) and 2-MIB (bottom) production by S. coelicolor A3(2) when grown as a pure culture and when co-cultured on NA plates with other bacteria and fungi. Results not connected by the same letter represent significant differences (p<0.05). Values represent the mean ± SE (n=3).
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Geosmin 50 45 A 40 35 AB 30 BC CD 25 20
ng/culture DE 15 E E E 10 E 5 0
2-MIB 8
7 A 6 5 AB 4 AB B B 3 B ng/culture B B 2 B 1 0
Figure 7.4 Geosmin (top) and 2-MIB (bottom) production by S. antibioticus when grown as a pure culture and when co-cultured on NA plates with other bacteria and fungi. Results not connected by the same letter represent significant differences (p<0.05). Values represent the mean ± SE (n=3).
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This trend was particularly profound in the case of S. coelicolor A3(2). Mean geosmin production by the pure culture was 503.5 ng, being significantly greater than under co- culturing conditions with all test microorganisms. Production of geosmin under co- culture conditions ranged from 8.6 ng when interacted with the fungal isolate to 108.4 ng when cultured with S. aureus, representing quantities of 58-fold and 4.6-fold less than that detected in the pure culture respectively. Regarding 2-MIB production, the highest and lowest yields under co-culturing conditions were obtained when S. coelicolor A3(2) was co-cultured with these same two organisms, with 2.5 ng and 22.3 ng produced in the presence of the fungal isolate and S. aureus respectively. In comparison, the pure culture produced a significantly greater yield of 48.7 ng, being 19.2 and 2.2-fold greater than the lowest and highest yield respectively generated under co-culturing conditions.
Pure cultures of S. antibioticus also generally produced greater amounts of both metabolites compared to when this species was co-cultured with other microorganisms.
However, the discrepancy between the yields was less profound than observed for S. coelicolor A3(2) and for some of the co-cultures did not vary significantly. Geosmin production by the pure culture reached 41.1 ng, which was not significantly greater compared to that produced when co-cultured with P. notatum (30.3 ng). Geosmin yields produced by S. antibioticus in all other co-cultures were significantly less compared to the pure culture, ranging from 6.82 ng to 28.9 ng when interacted with B. subtilis and M. luteus respectively. For 2-MIB, maximal production was detected in the pure culture (5.7 ng), which did not differ significantly to production yields obtained when interacted with
S. aureus (3.7 ng) and M. luteus (3.3 ng). Production of 2-MIB by S. antibioticus in other co-cultures were significantly less, ranging from 1.4 ng to 2.7 ng when interacted with P. notatum and the fungal isolate respectively. Both Streptomyces spp. produced greater yields by up to an order of magnitude of geosmin compared to 2-MIB under all conditions.
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For both species, the extent of differentiation observed was positively correlated with geosmin and 2-MIB production as shown by the moderate to high correlation coefficients in Figure 7.5. Greater yields (by up to an order of magnitude) of both compounds were obtained in the culture extracts of S. coelicolor A3(2). As seen in the S. coelicolor A3(2) photomicrographs of biomass (Figure 7.6) extensive differentiation was observed as indicated by the predominance of spores in addition to the co-culture photos (Figure 7.7) revealing the well-developed grey/white powdery surface of the cultures, indicative of a differentiating secondary mycelium. This was most apparent in the S. coelicolor A3(2) cultures which yielded the highest amounts of T&O metabolites, particularly the pure culture and those co-cultured with E. coli and S. aureus. On the contrary, cultures interacted with B. subtilis and the fungal isolate which produced the lowest yields revealed comparatively lower extents of differentiation as indicated by the presence of mostly undifferentiated mycelium observed in the photomicrographs (Figure 7.6) and by the absence of a well-developed grey secondary mycelium (Figure 7.7). For S. antibioticus, the predominance of mostly undifferentiated mycelium (Figure 7.8) and comparatively translucent cultures due to the absence of a mature secondary mycelium
(Figure 7.9) aligned with the lower production yields relative to S. coelicolor A3(2).
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Geosmin 2-MIB 2 -
600 60 MIBproduced culture by (ng) y = 8.09e0.57x y = 3.88e0.31x 500 R² = 0.739 R² = 0.475 50
400 40
300 30
200 20
100 10 Geosmin produced by culture (ng) culture by produced Geosmin 0 0 0 1 2 3 4 5
Differentiation value
Geosmin 2-MIB
y = 10.56e0.58x y = 1.93e0.35x 2 - 45 R² = 0.576 R² = 0.422 7 MIBproduecd culture by (ng) 40 6 35 5 30 25 4 20 3 15 2 10
5 1 Geosmin produced by culture (ng) culture by produced Geosmin 0 0 0 1 2 3 4 5 Differentiation value
Figure 7.5 Relationship between geosmin and 2-MIB production by S. coelicolor A3(2) (top) and S. antibioticus (bottom) and differentiation value under conditions of pure and co-cultures.
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Figure 7.6 Microscopic photos of co-cultures for studying production of geosmin and 2- MIB by S. coelicolor A3(2). The scale bar is 20 µm. Photos taken at 1000 x magnification under oil immersion. Note the absence of Streptomyces spores in the ‘fungal isolate’ image.
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Figure 7.7 Co-cultures for studying the effect of interaction with other microorganisms on the production of geosmin and 2-MIB by S. coelicolor A3(2). The blue pigment is the antibiotic actinorhodin produced by this strain. Note: secondary aerial mycelia are only partially evident in the B. subtilis, M. luteus and P. notatum plates and absent from the ‘fungal isolate’ plate.
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Figure 7.8 Microscopic photos of co-cultures for studying production of geosmin and 2- MIB by S. antibioticus. The scale bar is 20 µm. Photos taken at 1000 x magnification under oil immersion. Note the predominately vegetative hyphal form of S. antibioticus in these images.
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Figure 7.9 Co-cultures for studying the effect of interaction with other microorganisms on the production of geosmin and 2-MIB by S. antibioticus, revealing little evidence of powdery secondary mycelia.
7.3.2 Antimicrobial activity of geosmin and 2-MIB
To further test the hypothesis, the possibility that geosmin and 2-MIB exert antimicrobial effects on the diverse bacteria and two fungal species used in the co-culturing experiment was evaluated on NA using agar-plugs containing analytical standards of these compounds (35 µg/mL). In the case of all test microorganisms, dense growth was
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established across the entire surface of the NA plates with no zones of inhibition being observed around the agar plugs containing either geosmin, 2-MIB or the combination of both compounds (and the control) after 24 and 48 hours of incubation.
7.4 Discussion
Geosmin and 2-MIB are ubiquitous secondary metabolites in nature, produced particularly in soil by the abundant Streptomyces genus, providing this complex environment with its characteristic earthy-musty odour. To suggest that they serve no biological function seems inconceivable, thus raising the question of exactly what purpose the production of these compounds with potent odorous characteristics may serve. In this study, it was hypothesised that they, like many other secondary metabolites produced by the Streptomyces genus, function as antimicrobial compounds to supress the growth of other competing soil microorganisms.
It was considered that if geosmin and 2-MIB function in antagonistic capacities, by interacting Streptomyces spp. with a diverse range of bacterial and fungal species through co-culture conditions, partially simulating inter-microbial competition encountered in the soil environment, production of these metabolites would be enhanced compared to pure culture conditions. This has been previously demonstrated under co-culturing conditions for other secondary metabolites produced by Streptomyces which function as antimicrobial compounds (Luti & Mavituma, 2011a, 2011b, 2011c; Onaka et al., 2011;
Perez et al., 2011). However, the observation of a significant reduction in geosmin and
2-MIB production by Streptomyces spp. when co-cultured with other microorganisms compared to pure cultures suggests they are not part of the metabolic response of these bacteria to inter-microbial competition and thus are unlikely to function as antimicrobial compounds. The decrease in production of both secondary metabolites when interacted with other microorganisms could perhaps be attributed to use of cell resources (e.g.
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antibiotic precursors such as acetyl-CoA) in other metabolic pathways that are more active under such conditions, such as those for secondary metabolites that are necessary for supressing the growth of competing microorganisms (i.e. antimicrobial compounds).
The magnitude of reduction of geosmin and 2-MIB production compared to the pure culture varied significantly between co-cultures, probably reflecting different metabolic responses of the Streptomyces spp. to each of the competing microbes (perhaps differing demands for antimicrobial secondary metabolite production). The greatest reduction in geosmin and 2-MIB production by Streptomyces spp. tended to occur when co-cultured with organisms that are typically highly abundant in soil such as the fungal isolate and B. subtilis, which would represent the natural competitors of Streptomyces. Thus the metabolism of Streptomyces spp. when interacted with these organisms would presumably be geared towards antimicrobial secondary metabolite rather than geosmin and 2-MIB production, with precursor molecules being channelled into biosynthetic pathways for antimicrobial compounds. This compares to other organisms, such as S. aureus, which would not represent the natural competitors of Streptomyces, providing explanation for why geosmin and 2-MIB production and morphological differentiation more closely resembles that of the control, where no competition is occurring.
It is important to note, particularly in the case of S. coelicolor A3(2), that the extent of morphological differentiation was greatest in the control. This seems somewhat contradictory to expectations based on the notion supported in Chapter 6, that under conditions of nutrient limitation and thus potential competitive stress, Streptomyces shift to reproductive growth and differentiation. Such conditions would be expected to be enhanced when Streptomyces are subjected to interaction with another organism through co-culturing and not when grown as a mono-culture with an ample supply of nutrients provided in complex laboratory growth medium. The pure Streptomyces spp. cultures in
Chapter 6 grown when under the highest concentration of the major macronutrients carbon, nitrogen and phosphorous in the defined medium exhibited vegetative growth 276
only. In this study, when subjected to no competition under mono-culture conditions, profuse vegetative growth of Streptomyces on the more nutrient rich complex medium, may have ultimately induced nutrient limitation and possibly cell density signals, triggering formation of the secondary mycelium and sporulation and thus greater geosmin and 2-MIB production (Flardh & Buttner, 2009).
The absence of any zone of inhibition surrounding high concentrations of geosmin and/or
2-MIB by the diverse range of test microbes employed in the antimicrobial screening assay, provides strong evidence to suggest that they do not function as chemical weapons to antagonise competing microorganisms. This finding is in agreement with the absence of inhibitory effects of these compounds against selected bacteria and fungi in agar diffusion assays as reported by Schulz et al. (2010). The possibility of loss of the volatile compounds from the agar plugs during the incubation period must be considered, although the intense earthy and musty odours that could be detected after incubation suggested minimal loss had occurred. Taken together, the findings suggest that the role of geosmin and 2-MIB in the life of Streptomyces is unlikely to be as antimicrobial compounds.
The question then must be asked, if these metabolites do not serve as a chemical defence mechanism in a competitive situation, what selective advantage has the production of geosmin and 2-MIB provided Streptomyces with in their complex natural environment of soil? They are indeed produced ubiquitously, being the predominant odours associated with soil. The energetic costs and the complexity of the biosynthetic pathways of producing these low molecular weight compounds (described in Chapter 2) indicates they must play an important role in increasing the fitness of Streptomyces for survival
(Maplestone et al., 1992).
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The simultaneous production of geosmin and 2-MIB and sporulation in these bacteria perhaps points to a functional role of these secondary metabolites in relation to these reproductive structures. Secondary metabolites are generally produced at the onset of
Streptomyces aerial mycelia growth, as a defence mechanism at a time when they are particularly vulnerable to competition from other organisms, expending most of their metabolic energy on morphological differentiation (Maplestone et al., 1992). As observed in Streptomyces cultures in the present study and the work presented in Chapter 6, maximal production of geosmin and 2-MIB tends to occur during the differentiation process of sporulation that follows the establishment of the aerial mycelium.
Speculatively, this indicates that they function to assist these reproductive units in some manner to ensure the continuation of the species.
Zaitlin and Watson (2006) suggest that the increased production under certain circumstances indicates that rather than an overflow waste-disposal mechanism, these compounds may function as quorum sensing signal molecules. Perhaps geosmin and 2-
MIB function exogenously in cell-to-cell communication to stimulate and allow
Streptomyces to collectively control gene expression associated with the process of sporulation (Demain & Fang, 2000). In support of this proposition, Gust et al. (2003) found that mutations in the sesquiterpene synthase gene (cyc2) (involved in the initial steps of geosmin biosynthesis) which prevented geosmin production also reduced sporulation by S. coelicolor A3(2). Thus geosmin may not be obligatory for sporulation, but clearly it may stimulate the sporulation process. Numerous other studies have reported the association between T&O metabolite production by Streptomyces cultures and the process of sporulation (Dionigi et al., 1992; Scholler et al., 2002; Tung et al., 2006) and in some cases other secondary metabolites such as A-Factor (a γ-butyrolactone) in S. griseus and pamamycin produced by S. alboniger have also been suggested to induce sporulation in Streptomyces (Demain & Fang, 2000; Bibb, 2005).
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Considering that both geosmin and 2-MIB are potent odours in nature, with organisms including humans having developed extremely sensitive olfactory receptors to detect them, raises the question of whether they may serve as interspecies signalling molecules
(Marinelli & Marcone, 2011). VOCs are generally acknowledged as important groups of infochemicals (Rowan, 2011). These potent and ubiquitous earthy-musty odours may act as invertebrate (and indeed possibly vertebrate) attracting molecules, being infochemicals advertising moist soil, the conditions under which Streptomyces profusely grow and subsequently sporulate. The ingestion of soil by invertebrates entrained with the spores of Streptomyces would facilitate spore dispersal to other environments. Attraction and grazing responses of certain worms (Enchytraeus crypticus) and springtails (Collembola spp.) in the presence of Streptomyces species was thought to be induced by geosmin production, serving as a spore dispersal mechanism (Takahashi et al., 1984; Takahashi et al., 1991). There are numerous instances of microbial VOCs acting as insect semiochemicals, being closely associated with insect feeding behaviours, oviposition, inciting insect aggregations, indicating habitat suitability in addition to some being powerful insect repellents (Davis et al., 2013). Plants utilise VOCs to attract pollinators and seed dispersers and species of a few cactus genera scent their flowers with an earthy- smelling sesquiterpenoid similar to geosmin called dehydrogeosmin. Schlumpberger et al. (2004) observed that the emissions of the odour followed a diurnal rhythm, following the opening of the flowers, with the highest emission during maximum anthesis. Based on the timing and location of odour emission, being correlated with the timing of floral visitation, the authors hypothesised this earthy odour to play a role in plant-pollinator interactions. Furthermore, no antimicrobial effects of the volatile compound against selected bacteria and fungi were observed (Schlumpberger, 2002). In line with this finding, endogenous synthesis of geosmin has been confirmed in aseptically grown red beets (Beta vulgaris L.) (Lu et al., 2003). Collectively these observations support the proposition that this compound could be functionally relevant to plant pollen and/or seed dispersal. 279
Indeed, camels are thought to be particularly sensitive to geosmin and can sense
Streptomyces growing in moist soil for many miles in the desert. As the camels slake their thirst, the microbes would benefit by spore adherence to the camel for dispersion as they are carried to the next oasis (Hopwood, 2007). Thus the smell of geosmin and/or 2-MIB may be signature smell of moisture availability and a way of luring animals to disperse
Streptomyces spores, thus acting as mediators of cross species mutualism. Evolutionarily speaking, it makes sense for organisms such as humans and camels to be able to locate moisture, particularly if they live in a dry environment. This proposition provides a delightful explanation for the functional role of these compounds in nature and accounts for why their production is highly coordinated with the process of sporulation.
Another interesting proposal concerning a possible role of geosmin was provided by Tosi and Sola (1993). These authors examined the behaviour of migrating glass eels (Abguilla anguilla) to geosmin, observing high sensitivity to low concentrations, avoidance elicited by the unnatural combination of salinity and earth smell and evidence of a strong attraction as the level of salinity was reduced. It was concluded that geosmin is an important inland water marker involved in the orientation of glass eels towards freshwater, given that it is commonly found in freshwater, which further emphasises the high sensitivity of organisms to these compounds and their role as infochemicals.
While the aforementioned hypothesis makes biological sense, with some organisms being attracted to the potent odours of geosmin and 2-MIB which may benefit the dispersal of
Streptomyces spores, some recent studies have provided evidence to suggest that geosmin acts as an innate repelling odour to fruit flies. Becher et al. (2010) found that the addition of geosmin to vinegar significantly reduced positive chemotaxis in Drosophila melanogaster’s response to this innately attractive odour. More recently, Stensmyr et al.
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(2012) examined the functional significance of geosmin to the fly (D. melanogaster) and showed that geosmin activates a single class of olfactory sensory neurons which express an odour receptor that is exclusively tuned to geosmin. The authors showed that the functional significance of the segregated pathway was to detect and hence avoid microbes
(e.g. Streptomyces) that produce toxic compounds, some of which have insecticidal activity. Because geosmin itself is non-toxic to invertebrates and mammals, it may be used by flies as a universal warning signal for the presence of toxic compounds that are comorbid with geosmin and inhibits positive chemotaxis, oviposition (breeding) and feeding. The authors showed that flies avoided oviposition on a medium with S. coelicolor and feeding from a sucrose solution washed with the bacteria, but by abolishing geosmin production using a S. coelicolor strain (J3001) which carries a deletion in a geosmin synthesis gene, these avoidance behaviours were eliminated. The evolutionary significance of this neural pathway that is activated exclusively by geosmin is conserved in Drosophila and provides flies with a sensitive and specific means to identify suitable hosts (Stensmyr et al., 2012). Whether or not this innate avoidance behaviour is found among other invertebrates is unclear.
Although the primary function of production of geosmin and 2-MIB by Streptomyces may be to attract other organisms to facilitate the dispersal of their spores, it is highly plausible to suggest that organisms have evolved unique and diverse innate responses to these non- toxic potent odours, such as the fruit fly’s avoidance behaviour to geosmin signalling the presence of toxin-producing microbes, or the camel’s exploitation of these earthy-musty odours in the desert as infochemicals indicating the availability of water. Chemical sensing of these microbial metabolites has been conserved during evolution and they have clearly been exploited in diverse ways to benefit the survival of organisms in various environments.
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Another, less convincing hypothesis concerning their biological function is that these compounds may be used to lure other organisms such as motile microbes and invertebrates in the soil to the differentiating Streptomyces population, where they could be subsequently killed by the action of other antimicrobial and insecticidal metabolites, thus contributing to the Streptomyces nutrient source to facilitate completion of their life cycle (i.e. sporulation) (Hopwood, 2007).
An interesting possibility concerning the function of these metabolites produced by
Streptomyces is a potential role in controlling spore germination. Germination inhibitors have been identified in Streptomyces, such as germicidin, an antibiotic that is a specific inhibitor of ATP synthase and appears to be responsible for maintaining dormancy of spores (Demain & Fang, 2000). Geosmin and 2-MIB have been shown to specifically inhibit the germination of plant seeds (Brassicaceae species) exposed to concentrations of 100 µg/mL. Growth assays identified that the inhibition of seed germination was not due to the toxicity of geosmin and 2-MIB, as germination of seeds exposed to these compounds could be restored by stratification, gibberellin A3 treatment or seed coat removal, suggesting they act as growth-regulating substances during the germination process (Ogura et al., 2000). Hung et al. (2014) on the otherhand, demonstrated that geosmin did not inhibit germination of Arabidopsis thaliana seeds, although the concentration tested was comparatively lower (1 ppm). Perhaps Streptomyces produce these earthy-musty secondary metabolites to inhibit the germination of seeds and spores of other organisms, which represent competition and potentially could threaten their own survival and/or to inhibit germination of their own spores, thus delaying the production of the next generation until they have been dispersed to a new environment (where geosmin and 2-MIB may be absent or in lower concentration).
These metabolites may have a role in stimulating the germination of spores. Interestingly,
Carpenter-Boggs et al. (1995) showed that volatile compounds, particularly 2-MIB and 282
geosmin produced by cultures of Streptomyces isolates stimulated and positively correlated with spore germination of the arbuscular mycorrhizal fungus Gigasporu margarita. When placed in a common headspace, but physically separated from the
Streptomyces in a divided Petri dish, fungal spore germination rates were up to 73% compared to control treatments lacking these bacteria (23%). Whether these metabolites stimulate germination of the spores belonging to Streptomyces themselves is unclear.
Geosmin and 2-MIB are produced by other soil microorganisms including some fungi
(Kikuchi et al., 1981; Borjesson et al., 1993; Larsen & Frisvad, 1995), myxobacteria
(Yamamoto et al., 1994; Schulz et al., 2004; Dickschat et al., 2005) and amoeba (Hayes et al., 1991) in addition to plants such as liverworts (Sporle et al., 1991) and red beets (Lu et al., 2003). Given their functionally similar morphological characteristics to
Streptomyces (i.e. spore and seed producers) perhaps suggests a similar biological purpose of production of geosmin and 2-MIB in the life of these taxonomically diverse microorganisms. Indeed, geosmin production has also been associated with sporulation in the fungus Penicillium expansum (Judet-Correia et al., 2013). Cyanobacteria, which are regarded as the major producers of geosmin and 2-MIB in aquatic environments
(Juttner & Watson, 2007), also have complex multicellular morphologies and produce spore-like survival structures called akinetes. Intracellular pools of these compounds are known to be released from the cells of these photosynthetic bacteria at the end of their growth cycle (late exponential to stationary phase when cell lysis occurs), when differentiation of akinetes is also initiated (Fay et al., 1984; Rosen et al., 1992; Li et al.,
1997; Watson, 2003; Zhang et al., 2009). Whether or not a common biological function of geosmin and 2-MIB exists between Streptomyces, other soil organisms and aquatic cyanobacteria should be further investigated.
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7.5 Conclusions
Geosmin and 2-MIB, like other secondary metabolites produced by Streptomyces, are products of evolutionary selection over millennia, suggesting that they must play some important functional role in the competitive survival of these bacteria. The findings reported in this study contradict the hypothesis that geosmin and 2-MIB function as bacterial chemical weapons in inter-microbial competition. Several alternative propositions based on literature evidence, have been suggested for the biological purpose of production of these compounds including assisting the formation, germination and/or dispersal of their spores. It is clear that geosmin and 2-MIB production is highly coordinated with the process of sporulation and it seems plausible to consider, as proposed by Gust et al. (2003), that they may indeed act as signalling molecules and control gene expression associated with this process. Based on some partial evidence in the literature, they could be considered to function in regulating (stimulating or inhibiting) the germination of their spores, or indeed the spores or seeds of other organisms. The proposition that geosmin and 2-MIB function as infochemicals, attracting other organisms which in turn facilitate the dispersal of Streptomyces spores, seems rather convincing and accounts for why their production is highly coordinated with sporulation.
Indeed, some plants have been found to produce a similar compound to geosmin which has been associated with attracting pollinators and seed dispersers (Schlumpberger,
2002). Chemical sensing of these microbial metabolites by other organisms has been conserved during evolution, with humans themselves possessing extremely sensitive olfactory receptors to detect these ubiquitous earthy-musty odours in the environment.
There is mutual benefit arising from other organisms being attracted to the scent of geosmin and 2-MIB produced by Streptomyces. The spores of these bacteria can be dispersed to new environments while those agents of dispersal attracted to earthy-musty scented spores in moist soil are provided with a reward (i.e. availability of moisture) enhancing their survival.
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While this study does not provide information concerning the management of
Streptomyces- derived earthy-musty odours in drinking water supplies, it does broaden the view of the possible biological purpose of production of these compounds, a concept which has been thus far neglected. Rather than being regarded simply as waste products of metabolism, a widespread opinion held in the water industry (Watson, 2003), it seems apparent that they are more likely produced for a biological purpose to benefit the survival of Streptomyces. In this respect, the concept that these metabolites may function to lure other organisms for spore dispersal is highly plausible and certainly warrants further investigation.
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CHAPTER 8: CONCLUSIONS AND RECOMMENDATIONS FOR FUTURE RESEARCH
Despite the wealth of literature that has accumulated over recent decades on T&O involving geosmin and 2-MIB in drinking water supplies, there is still much to be learnt about the breadth and depth of this issue, particularly in the Australian water industry.
There is especially a deficiency in the literature concerning the potential involvement of
Streptomyces, which have been speculated to be a major contributor, particularly where
T&O events cannot be attributed to cyanobacteria.
This research project was initiated in response to past geosmin and 2-MIB episodes which were unrelated to cyanobacteria in HWC’s primary drinking water supplies,
Grahamstown and Chichester Reservoirs. The historical water quality data analysis presented in Chapter 3 revealed that the majority (but not all) geosmin episodes were related to the dynamics of the cyanobacterial genus Anabaena, while 2-MIB episodes could not be attributed to any biological parameter routinely monitored in the reservoirs.
This prompted research concerning the potential role of Streptomyces in contributing to these unexplained T&O events.
The study presented in Chapter 4 comprised both field and laboratory work to assess the potentiality of Streptomyces to contribute to T&O in these reservoirs. This involved an investigation of Streptomyces population dynamics (abundance and activity) in a variety of habitats in the reservoirs, identifying and assessing the geosmin and 2-MIB producing ability of Streptomyces spp. isolated from these environments, and examining the capacity of environmental substrates to support their growth and T&O production.
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The results of the detailed spatial and temporal sampling program undertaken at both reservoirs indicated that Streptomyces were significantly more abundant in the water mass during wet conditions compared to extended dry conditions, supporting the notion of the terrestrial wash-in of these bacteria, and potentially, their odorous secondary metabolites, from marginal environments. This represents a highly probable means by which
Streptomyces contribute to T&O episodes. Streptomyces were found to populate the soils, sediments and plant debris at the margins surrounding the reservoirs, being significantly less abundant and active in bottom sediments. They are indeed renowned terrestrial microorganisms. During dry conditions, exposed marginal substrates (soil, sediment and plant debris) were generally found to harbour significantly greater Streptomyces populations. This provided evidence to support the hypothesis that exposure of marginal sediments stimulates the growth and conceivably, odour production of these aerobic bacteria, which may therefore represent a significant source of geosmin and 2-MIB which can enter adjacent water bodies via wash-in.
The laboratory study presented in Chapter 5 however, found that levels of geosmin and
2-MIB in marginal sediments obtained from Chichester Reservoir and inoculated with a
Streptomyces sp. were similar irrespective of exposed or submerged conditions, although evidence for increased growth under exposed conditions was provided. This simulation experiment suggested that sufficient oxygen availability in the shallow submerged sediments of reservoirs may permit the growth and production of these compounds by these aerobic bacteria and may represent a constant background source of geosmin and
2-MIB. However, the possibility that a fraction of these compounds were lost due to volatilisation under the laboratory conditions in the exposed sediment must be considered, particularly before drawing any firm conclusions regarding management implications.
Based on the findings, the management options for controlling T&O potentially originating from these bacteria relate to minimising water level fluctuations and the stabilisation of shorelines to prevent introduction of marginal substrates entrained with 287
Streptomyces and their odorous metabolites into the water mass during rain events.
Managers may consider that stabilisation of the marginal sediments of reservoirs by establishing populations of macrophyte species (such as torpedo grass) may be an appropriate strategy to prevent Streptomyces induced T&O events. Macrophyte species assessed in both reservoirs were found to support significantly low Streptomyces population densities and are therefore an unlikely substrate to support T&O metabolite production by these bacteria.
The ability of these terrestrially-derived bacteria to continue to actively grow in the water mass and contribute directly to T&O has been largely disputed in the literature (Cross,
1981; Jensen et al., 1994; Zaitlin et al., 2003b). However, the detection of Streptomyces vegetative cells in the water mass, particularly during dry periods when the wash-in of these bacteria would be minimal, in addition to the observation that sterilised reservoir water supported the growth and odour production by Streptomyces spp., implies their potential capacity for in situ T&O production.
Previous studies attempting to relate Streptomyces to geosmin and 2-MIB occurrence in reservoirs have been undermined by the use of culture-dependent techniques which do not allow the discrimination between dormancy (spores) or actively growing vegetative cells. This study is the first attempt to quantify Streptomyces populations in drinking water reservoir habitats using a 16S rRNA-targeting, genus-specific qPCR assay and to further discriminate between vegetative cells and spores by employing differential cell lysis procedures for the extraction of environmental DNA. Future studies at other locations in Australia or indeed worldwide where T&O events are problematic, could employ this molecular approach and correlate such data with geosmin and 2-MIB measurements in freshwater systems, which were absent in the present study.
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Furthermore, such studies could involve more intensive spatial and temporal sampling programs to provide firmer evidence linking Streptomyces to T&O.
The multitude of Streptomyces isolates obtained in the field study and identified by 16S rRNA sequence analysis in Chapter 4 were all found to be capable of producing geosmin and 2-MIB, adding further evidence to support the potential ability of Streptomyces found within the reservoirs to contribute to T&O episodes. It should be emphasised that
Streptomyces are certainly not the only geosmin and 2-MIB producing genus of actinobacteria in freshwater environments. Indeed, a Nocardia sp. was isolated in this study and found to be a producer of both compounds. In addition to primers targeting
Streptomyces-specific 16S rRNA sequences, a primer pair with a broader taxonomic specificity targeting 16S rRNA sequences unique to the class actinobacteria was also applied to environmental DNA (Schafer et al., 2010). The data for this molecular analysis is presented in Appendix D. Comparisons of the Streptomyces and actinobacteria qPCR data (Appendix D, Figure D.3) revealed that Streptomyces comprised a small proportion of the total actinobacterial population in most samples (<0.1% of the total abundance).
This indicates the presence of other genera of the class actinobacteria in both reservoirs which may also contribute to T&O episodes (as previously outlined in Chapter 2), such as Nocardia, Micromonospora, Microbispora, Actinomadura, Actinosynnema,
Kitasatospora, Rothia, Frankia and Saccharopolyspora (Gerber, 1969, 1979; Giglio et al., 2008; Schrader & Summerfelt, 2010; Citron et al., 2012). Published reports investigating the potential contribution of these genera to T&O episodes involving geosmin and 2-MIB are lacking. It is recommended that future studies target the isolation of such genera from drinking water supplies to assess their T&O producing capacity and utilise genus-specific primers in qPCR assays to determine their population dynamics.
Such systematic investigations may reveal causal microorganisms of T&O that have been previously overlooked.
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The study in Chapter 4 employed a 16S rRNA gene-based assay for detecting and quantifying Streptomyces. Although the qPCR protocol developed to target this taxonomic marker proved to be a sensitive, reliable and relatively rapid method for detecting and measuring the abundance of Streptomyces in environmental samples, a disadvantage of this molecular assay is that it cannot differentiate between producers and non-producers of geosmin and 2-MIB. It is well established that not all Streptomyces produce these compounds (Jensen et al., 1994; Zaitlin et al., 2003b). Examining the T&O producing ability of this genus was achieved by screening reservoir isolates that were cultivated on selective isolation media. In recent years, qPCR protocols have been developed to target the functional genes that encode enzymes responsible for geosmin and 2-MIB biosynthesis in a number of taxa. These assays have demonstrated their utility in monitoring and predicting T&O events, particularly for the detection of geosmin- producing Anabaena spp. (Su et al., 2013; Kutovaya & Watson, 2014; Tsao et al., 2014a).
Various authors (Auffret et al., 2011; Du et al., 2013) have developed primers for qPCR assays targeting Streptomyces based on multiple sequence alignment of the geosmin synthase gene from this genus, which have successfully detected the biosynthetic gene and correlated its abundance with geosmin production in cultures of Streptomyces.
However, when applied to environmental DNA, such primers have failed to detect sequences or have amplified those of other taxa associated with geosmin production such as Sorangium and Nannocystis (Myxococcales), suggesting their low specificity for
Streptomyces (Auffret et al., 2013; Kutovaya & Watson, 2014). This issue is likely attributable to the low sequence conservancy of these genes. Future studies should aim to improve qPCR assays for the detection of geosmin and 2-MIB producing Streptomyces or other actinobacteria, by developing more taxa-specific primers targeting the genes involved in the biosynthesis of these compounds. Such information could provide insight on the abundance and distribution of microorganisms with the genetic potential to contribute to T&O events in various environments. 290
The multivariate laboratory studies presented in Chapter 6 demonstrated how several physico-chemical parameters and interactions between them influence the production of geosmin and 2-MIB by Streptomyces spp. and established that the production of these
T&O metabolites was highly coordinated with the reproductive (sporulation) stage of the
Streptomyces life cycle. Thus physico-chemical factors that trigger Streptomyces to cease vegetative growth and enter the reproductive developmental stage would conceivably allow elucidation of the conditions which also trigger significant production of their T&O metabolites. Morphological differentiation and T&O metabolite production was greatest in the combined presence of low nitrogen and phosphorous concentration (and low carbon concentration in the case of S. coelicolor A3(2)), whereas highest concentration of NaCl and copper favoured vegetative growth, thus inhibiting T&O metabolite production.
Greater production of T&O metabolites under alkaline conditions and lower temperature was established, although these parameters did not influence the growth stage, suggesting that their effects were independent of morphological differentiation. Other factors were comparatively less influential over the ranges investigated. In consideration of the results from this laboratory study, the levels of physico-chemical characteristics of Grahamstown and Chichester Reservoirs appear to be suitable for Streptomyces growth, differentiation and production of geosmin and 2-MIB. It is recommended that future studies examine the effects of all environmental variables simultaneously rather than in separate two-level full factorial experimental designs used in this study. A switch to fractional factorial design is favourable for this purpose, enabling a wide range of conditions potentially encountered in the environment to be examined in relation to geosmin and 2-MIB production by Streptomyces which may assist in the development of options to control in situ production of these earthy-musty secondary metabolites.
Finally, the research presented in Chapter 7 explored the biological purpose of production of these terpenoids by Streptomyces. The experimental work provided evidence to challenge the hypothesis that they are biosynthesised to serve an antagonistic function in 291
inter-microbial competition (i.e. antimicrobial compounds). The close association between production of these secondary metabolites and the process of sporulation as demonstrated in Chapter 6 and 7 and previously described in the literature, suggests a functional role related to spores, possibly in controlling spore formation, spore germination or attracting other organisms for spore dispersal. While management implications for Streptomyces- derived T&O problems were not provided from this particular study, the possible biological purpose of production of these compounds, a concept which has been thus far largely neglected in the literature, has been extended.
The propositions concerning the possible functional role of these secondary metabolites in the life of Streptomyces as regulators of spore formation, germination and/or dispersal, certainly warrant further investigation.
Overall, this research has illuminated a number of issues in relation to determining the potential significance of Streptomyces as contributors to problematic geosmin and 2-MIB associated T&O events in drinking water supplies. The information gathered in this research together with recommended future work as outlined, will assist in unravelling the extent and mechanisms by which these filamentous bacteria and other actinobacteria contribute to the occurrence of geosmin and 2-MIB in freshwater systems. Such knowledge would provide substantial aid to water utilities in making evidence-based decisions in the management of Streptomyces induced T&O problems.
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REFERENCES Adams, B. A. (1929). Odours in the water of the Nile River. Water and Water Engineering, 31, 309-314. Ahmad, I., Sajjad, M., Khan, A., Aqil, K., & Singh, M. (2011). Microbial applications in agriculture and the environment: A broad perspective. In I. Ahmad, F. Ahmad & J. Pichtel (Eds.), Microbes and microbial technology: Agricultural and environmental applications (pp. 1-28). New York: Springer. Alexander, D. B. (2002). Soil bacteria. In G. Bitton (Ed.), Encyclopedia of environmental microbiology (Vol. 5, pp. 2880-2894). New York: John Wiley & Sons, Inc. Alferova, I. V., Terekhova, L. P., & Prauzer, K. H. (1989). Selective medium with nalidixic acid for isolating antibiotic-producing Actinomyces. Antibiot Khimioter, 34(5), 344-348. ANZECC, & ARMCANZ. (2000). Australian and New Zealand Guidelines for fresh and marine water quality. Volume 1: The guidelines. Artarmon, NSW: Australian and New Zealand Environment and Conservation Council Agriculture and Resource Management Council of Australia and New Zealand. Aoyama, K. (1990). Studies on the earthy-musty ddors in natural-water. 4. Mechanism of earthy-musty odor production of actinomycetes. Journal of Applied Bacteriology, 68(4), 405-410. Auffret, M., Pilote, A., Proulx, E., Proulx, D., Vandenberg, G., & Villemur, R. (2011). Establishment of a real-time PCR method for quantification of geosmin- producing Streptomyces spp. in recirculating aquaculture systems. Water Research, 45(20), 6753-6762. Auffret, M., Yergeau, E., Pilote, A., Proulx, E., Proulx, D., Greer, C. W., . . . Villemur, R. (2013). Impact of water quality on the bacterial populations and off-flavours in recirculating aquaculture systems. Fems Microbiology Ecology, 84(2), 235-247. Bae, S., & Wuertz, S. (2009). Discrimination of viable and dead fecal Bacteroidales bacteria by quantitative PCR withpropidium monoazide. Applied and Environmental Microbiology, 75(9), 2940-2944. Bailey, A. J. (1988). Odours in drinking water: a case study. Water, Sewage and Effluent, 8(4), 13-16. Baker, R. A. (1961). Taste and odor in water: A critical review (pp. 159 p). Philadelphia: The Franklin Institute. Bartholomew, K. A. (1958). Control of earthy, musty odours in water by treatment with residual copper. Journal American Water Works Association, 50(1), 481. Bays, L. R., Burman, N. P., & Lewis, W. M. (1970). Taste and odour in water supplies in Great Britain: a survey of the present position and problems for the future. Water Treatment and Examination, 19, 136-160. Becher, P. G., Bengtsson, M., Hansson, B. S., & Witzgall, P. (2010). Flying the fly: long- range flight behavior of Drosophila melanogaster to attractive odors. Journal of Chemical Ecology, 36(6), 599–607. Bentley, R., & Meganathan, R. (1981). Geosmin and methylisoborneol biosynthesis in streptomycetes. Evidence for an isoprenoid pathway and its absence in non- differentiating isolates. Febs Letters, 125(2), 220-222. Bentley, S. D., Chater, K. F., Cerdeno-Tarraga, A. M., Challis, G. L., Thomson, N. R., James, K. D., . . . Hopwood, D. A. (2002). Complete genome sequence of the model actinomycete Streptomyces coelicolor A3(2). Nature, 417(6885), 141-147. Bibb, M. (2005). Regulation of secondary metabolism in streptomycetes Current Opinion in Microbiology, 8(2), 208-215.
293
Blevins, W. T., Schrader, K. K., & Saadoun, I. (1995). Comparative physiology of geosmin production by Streptomyces halstedii and Anabaena sp. Water Science and Technology, 31(11), 127-133. Borjesson, T. S., Stollman, U. M., & Schnurer, J. L. (1993). Off-odourous compounds produced by moulds on oatmeal agar: identification and relation to other growth characteristics Journal of Agricultural and Food Chemistry, 41(11), 2104-2111. Bowmen, K. H., Padovan, A., Oliver, R. L., Korth, W., & Ganf, G. G. (1992). Physiology of geosmin production by Anabaena circinalis isolated from the Murrumbidgee River, Australia. Water Science & Technology, 25(2), 259-267. Bredholt, H., Fjaervik, E., Johnsen, G., & Zotchev, S. B. (2008). Actinomycetes from sediments in the Trondheim Ford, Norway: Diversity and biological activity. Marine Drugs, 6(1), 12-24. Burch, M., & Baker, P. (2000). Common algal species in South Australian waters, potential toxicity and odours Australian Water Quality Centre Report: Australian Water Quality Centre. Burgos, L., Lehmann, M., de Andrade, H. H. R., de Abreu, B. R. R., de Souza, A. P., Juliano, V. B., & Dihl, R. R. (2014). In vivo and in vitro genotoxicity assessment of 2-methylisoborneol, causal agent of earthy-musty taste and odor in water. Ecotoxicology and Environmental Safety, 100, 282-286. Butakova, E. A. (2013). Specific features of odor-causing compounds (geosmin and 2- methylisoborneol) as secondary metabolites of cyanobacteria. Russian Journal of Plant Physiology, 60(4), 507-510. Cane, D. E., He, X. F., Kobayashi, S., Omura, S., & Ikeda, H. (2006). Geosmin biosynthesis in Streptomyces avermitilis. Molecular cloning, expression, and mechanistic study of the germacradienol/geosmin synthase. Journal of Antibiotics, 59(8), 471-479. Cane, D. E., & Watt, R. M. (2003). Expression and mechanistic analysis of a germacradienol synthase from Streptomyces coelicolor implicated in geosmin biosynthesis. Proceedings of the National Academy of Sciences of the United States of America, 100(4), 1547-1551. Carpenter-Boggs, L., Loynachan, T. E., & Stahl, P. D. (1995). Spore germination of Gigaspora margarita stimulated by volatiles of soil-isolated actinomycetes. Soil Biology and Biochemistry, 27(11), 1445-1451. Cees, B., Zoeteman, B. C. J., & Piet, G. J. (1974). Cause and identification of taste and odour compounds in water. Science of the Total Environment, 3(1), 103-115. Charusanti, P., Fong, N. L., Nagarajan, H., Pereira, A. R., Li, H. J., Abate, E. A., . . . Palsson, B. O. (2012). Exploiting adaptive laboratory evolution of Streptomyces clavuligerus for antibiotic discovery and overproduction. Plos One, 7(3), 1-12. Cheng, Y. R., Hauck, L., & Demain, A. L. (1995). Phosphate, ammonium, magnesium and iron nutrition of Streptomyces hygroscopicus with respect to rapamycin biosynthesis. Journal of Industrial Microbiology, 14(5), 424-247. Citron, C. A., Gleitzmann, J., Laurenzano, G., Pukall, R., & Dickschat, J. S. (2012). Terpenoids are widespread in actinomycetes: A correlation of secondary metabolism and genome data. Chembiochem, 13(2), 202-214. Grahamstown Dam water quality & aquatic ecological functioning (2011). Cotarlet, M., Bahrim, G., Negoita, T., & Stougaard, P. (2010). Comparative study for establishing the efficiency of some methods for chromosomal DNA extraction from cold adapted streptomycetes. Romanian Biotechnological Letters, 15(4), 5482-5486.
294
Cresswell, N., Saunders, V. A., & Wellington, E. M. H. (1991). Detection and quantification of Streptomyces violaceolatus plasmid DNA in soil. Letters in Applied Microbiology, 13(4), 193-197. Cross, T. (1981). Aquatic actinomycetes: A critical survey of the occurrence, growth and role of actinomycetes in aquatic habitats. Journal of Applied Bacteriology, 50(3), 397-423. Cueto, M., Jensen, P. R., ., Kauffman, C., Fenical, W., Lobkovsky, E., & Clardy, J. (2001). Pestalone, a new antibiotic produced by a marine fungus in response to bacterial challenge. Journal of Natural Products, 64(11), 1444-1446. Davis, T. S., Crippen, T. L., Hofstetter, R. W., & Tomberlin, J. K. (2013). Microbial volatile emissions as insect semiochemicals Journal of Chemical Ecology, 39(7), 840-859. Daza, A., Martin, J. F., Dominguez, A., & Gil, J. A. (1989). Sporulation of several species of Streptomyces in submerged cultures after nutritional downshift. Journal of General Microbiology, 135(9), 2483-2491. Demain, A. L., & Fang, A. (2000). The natural functions of secondary metabolites. In A. FIechter (Ed.), History of modern biotechnology I (ed., Vol. 69, pp. 1-39). Berlin, Heidelberg: Springer-Verlag. Denisova, L. Y., Bel'kova, N. L., Tulokhonov, I. I., & Zaichikov, E. F. (1999). Bacterial diversity at various depths in the southern part of Lake Baikal as revealed by 16S rDNA sequencing. Mikrobiologiia, 68(4), 475-483. Dickschat, J. S., Bode, H. B., Wenzel, S. C., Muller, R., & Schulz, S. (2005). Biosynthesis and identification of volatiles released by the myxobacterium Stigmatella aurantiaca. Chembiochem, 6(11), 2023-2033. Dickschat, J. S., Nawrath, T., Thiel, V., Kunze, B., Muller, R., & Schulz, S. (2007). Biosynthesis of the off-flavor 2-methylisoborneol by the myxobacterium Nannocystis exedens. Angewandte Chemie-International Edition, 46(43), 8287- 8290. Dionigi, C. P., Ahten, T. S., & Wartelle, L. H. (1996). Effects of several metals on spore, biomass, and geosmin production by Streptomyces tendae and Penicillium expansum. Journal of Industrial Microbiology, 17(2), 84-88. Dionigi, C. P., & Ingram, D. A. (1994). Effects of temperature and oxygen concentration on geosmin production by Streptomyces tendae and Penicillium expansum. Journal of Agricultural and Food Chemistry, 42(1), 143-145. Dionigi, C. P., Lawlor, T. E., Mcfarland, J. E., & Johnsen, P. B. (1993). Evaluation of geosmin and 2-methylisoborneol on the histidine dependence of Ta98 and Ta100 Salmonella typhimurium tester strains. Water Research, 27(11), 1615-1618. Dionigi, C. P., Millie, D. F., Spanier, A. M., & Johnsen, P. B. (1992). Spore and geosmin production by Streptomyces tendae on several media. Journal of Agricultural and Food Chemistry, 40(1), 122-125. Doull, J. L., & Vining, L. C. (1990). Nutritional control of actinorhodin production by Streptomyces coelicolor A3(2): Suppressive effects of nitrogen and phosphate Applied Microbiology and Biotechnology, 32(4), 449-454. Downing, J. A., Watson, S. B., & McCauley, E. (2001). Predicting cyanobacteria dominance in lakes Canadian Journal of Fisheries and Aquatic Sciences, 58(10), 1905-1908. Du, H., Lu, H., Xu, Y., & Du, X. W. (2013). Community of environmental Streptomyces related to geosmin development in Chinese liquors. Journal of Agricultural and Food Chemistry, 61(6), 1343-1348.
295
Dyson, P. (2009). Streptomyces In S. Mosello (Ed.), Encyclopedia of microbioloy (3rd ed., pp. 318-332). Oxford: Academic Press. Dzialowski, A. R., Smith, V. H., Huggins, D. G., deNoyelles, F., Lim, N. C., Baker, D. S., & Beury, J. H. (2009). Development of predictive models for geosmin-related taste and odour in Kansas, USA, drinking water reservoirs. Water Research, 43(11), 2829-2840. Ebner, J. B., & Frea, J. I. (1970). Heat resistance during life cycle of Streptomyces fradiae. Microbios, 2(5), 43-48. Elliot, M. A., & Flardh, K. (2012). Streptomycete spores Encyclopedia of life sciences (eLS). Chichester John Wiley & Sons Ltd. Eriksson, L., Johansson, E., Kattenah-Wold, N., Wikstrom, C., & Wold, S. (2008). Design of experiments: principles and applications (3rd ed.). Sweden: Umetrics Academy Fay, P., Lynn, J. A., & Majer, S. C. (1984). Akinete development in the planktonic blue- green alga Anabaena circinalis. British Phycological Journal, 19(2), 163-173. Filippova, S. N., Gorbatyuk, E. V., Poglazova, M. N., Soina, V. S., Kuznetsov, V. D., & El'-Registan, G. I. (2005). Endospore formation by Streptomyces avermitilis in submerged culture. Microbiology, 74(2), 204-214. Fischer, M., Alderson, J., van Keulen, G., White, J., & Sawers, R. G. (2010). The obligate aerobe actinomycete Streptomyces coelicolor A3(2) synthesizes three active respiratory nitrate reductases. Microbiology and Molecular Biology Reviews, 156(10), 3166-3179. Flardh, K., & Buttner, M. J. (2009). Streptomyces morphogenetics: dissecting differentiation in a filamentous bacterium. Nature Reviews Microbiology, 7(1), 36-49. Francisco, D. E., & Silvey, J. K. G. (1971). The effect of carbon monoxide inhibition on the growth of an aquatic streptomycete. Canadian Journal of Microbiology, 17(3), 347-351. Franco-Correa, M., Quintana, A., Duque, C., Suarez, C., Rodriguez, M. X., & Barea, J. M. (2010). Evaluation of actinomycete strains for key traits related with plant growth promotion and mycorrhiza helping activities. Applied Soil Ecology, 45(3), 209-217. Gagne, F., Ridal, J., Blaise, C., & Brownlee, B. (1999). Toxicological effects of geosmin and 2-methylisoborneol on rainbow trout hepatocytes. Bulletin of Environmental Contamination and Toxicology, 63(2), 174-180. Geladi, P., & Kowalski, B. R. (1986). Partial least-squares regression: A tutorial. Anlytica Chimica Acta, 185, 1-17. Geldreich, E. E. (2002). Controlling the microbial quality of drinking water in distribution systems. In G. Bitton (Ed.), Encyclopedia of environmental microbiology (Vol. 2, pp. 941-952). New York: John Wiley & Sons, Inc. Gerber, N. N. (1969). A volatile metabolite of actinomycetes, 2-methylisoborneol. Journal of Antibiotics, 22(10), 508-509. Gerber, N. N. (1979). Volatile substances from actinomycetes: their role in odour pollution of water. CRC Critical Reviews in Microbiology, 7, 191-214. Gerber, N. N., & Lecheval, H. A. (1965). Geosmin an earthy-smelling substance isolated from actinomycetes. Applied Microbiology, 13(6), 935-938. Gersch, D., Skurk, A., & Romer, W. (1979). Phosphate inhibition of secondary metabolism in Streptomyces hygroscopicus and its reversal by cyclic AMP. Archives of Microbiology, 121(1), 91-96.
296
Ghimire, G. P., Oh, T. J., Lee, H. C., Kim, B. G., & Sohng, J. K. (2008). Cloning and functional characterization of the germacradienol synthase (spterp13) from Streptomyces peucetius ATCC 27952. Journal of Microbiology and Biotechnology, 18(7), 1216-1220. Giglio, S., Chou, W. K. W., Ikeda, H., Cane, D. E., & Monis, P. T. (2011). Biosynthesis of 2-methylisoborneol in cyanobacteria. Environmental Science & Technology, 45(3), 992-998. Giglio, S., Jiang, J. Y., Saint, C. P., Cane, D. E., & Monis, P. T. (2008). Isolation and characterization of the gene associated with geosmin production in cyanobacteria. Environmental Science & Technology, 42(21), 8027-8032. Glazebrook, M. A., Doull, J. L., Stuttard, C., & Vining, L. C. (1990). Sporulation of Streptomyces venezuelae in submerged cultures. Journal of General Microbiology, 136(3), 581-588. Goodfellow, M., & Williams, S. T. (1983). Ecology of actinomycetes. Annual Review of Microbiology, 37, 189-216. Groisman, E. A. (2001). The pleiotropic two-component regulatory system PhoP-PhoQ. Journal of Bacteriology, 183(6), 1835-1842. Gust, B., Challis, G. L., Fowler, K., Kieser, T., & Chater, K. F. (2003). PCR targeted Streptomyces gene replacement identifies a protein domain needed for biosynthesis of the sesquiterpene soil odor geosmin. PNAS, 100(4), 1541–1546. Guttman, L., & van Rijn, J. (2008). Identification of conditions underlying production of geosmin and 2-methylisoborneol in a recirculating system. Aquaculture, 279(1- 4), 85-91. Guttman, L., & van Rijn, J. (2012). Isolation of bacteria capable of growth with 2- methylisoborneol and geosmin as the sole carbon and energy sources. Applied and Environmental Microbiology, 78(2), 363-370. Hayes, S. J., Hayes, K. P., & Robinson, B. S. (1991). Geosmin as an odourous metabolite in cultures of a free-living amoeba, Vannella species (Gymnamoebia, Vannellidae). Journal of Protozoology, 38(1), 44-47. Henatsch, J., & Juttner, F. (1990). Seasonal dynamics of organic volatile substances (odors) - Lake Schlein. Ecology and Management of Small Bodies of Standing Waters, 3, 93-126. Higgins, M. L., & Silvey, J. K. G. (1966). Slide culture observations of two freshwater actinomycetes. Transactions of the American Microscopical Society, 85(3), 390- 398. Hirsch, P. R., Mauchline, T. H., & Clark, I. M. (2010). Culture-independent molecular techniques for soil microbial ecology Soil Biology & Biochemistry, 42(6), 878- 887. Ho, L., Hoefel, D., Bock, F., Saint, C. P., & Newcombe, G. (2007). Biodegradation rates of 2-methylisoborneol (MIB) and geosmin through sand filters and in bioreactors. Chemosphere, 66(11), 2210-2218. Ho, L., Sawade, E., & Newcombe, G. (2012). Biological treatment options for cyanobacteria metabolite removal: A review. Water Research, 46(5), 1536-1548. Hobson, P., Fazekas, C., House, J., Daly, R., Kildea, T., Giglio, S., . . . Chen, Y.-M. (2010). Taste and odours in reservoirs (W. Q. R. Australia Ed.). Adelaide, SA Water Quality Research Australia Ltd. Hopwood, D. A. (2007). Streptomyces in nature and medicine: the antibiotic makers. New York Oxford University Press, Inc.
297
Hsu, S. C., & Lockwood, J. L. (1975). Powdered chitin agar as a selective medium for enumeration of actinomycetes in water and soil. Applied Microbiology, 29(3), 422-426. Hung, R., Lee, S., Rodriguez-Saona, C., & Bennett, J. W. (2014). Common gas phase molecules from fungi affect seed germination and plant health in Arabidopsis thaliana. AMB Express, 4(53), 1-7. Issatchenko, V. L. (1946). Flavours and tastes of water. Priroda Mosk, 3, 26-33. Issatchenko, V. L., & Egorova, A. (1944). Actinomycetes in reservoirs as one of the causes responsible for the earthy smell of their waters. Microbiologia, 13(5), 216- 225. Izaguirre, G., & Taylor, W. D. (2007). Geosmin and MIB events in a new reservoir in southern California. Water Science and Technology, 55(5), 9-14. Izaguirre, G., Taylor, W. D., & Pasek, J. (1999). Off-flavor problems in two reservoirs, associated with planktonic Pseudanabaena species Water Science and Technology, 40(6), 85-90. Jahnichen, S., Jaschke, K., Wieland, F., Packroff, G., & Benndorf, J. (2011). Spatio- temporal distribution of cell-bound and dissolved geosmin in Wahnbach Reservoir: Causes and potential odour nuisances in raw water. Water Research, 45(16), 4973-4982. Jensen, P. R., Dwight, R., & Fenical, W. (1991). Distribution of actinomycetes in near- shore tropical marine-sediments. Applied and Environmental Microbiology, 57(4), 1102-1108. Jensen, P. R., Mincer, T. J., Williams, P. G., & Fenical, W. (2005). Marine actinomycete diversity and natural product discovery. Antonie Van Leeuwenhoek International Journal of General and Molecular Microbiology, 87(1), 43-48. Jensen, S. E., Anders, C. L., Goatcher, L. J., Perley, T., Kenefick, S., & Hrudey, S. E. (1994). Actinomycetes as a factor in odor problems affecting drinking-water from the North Saskatchewan River. Water Research, 28(6), 1393-1401. Jensen, V. (1971). The bacterial flora of Beech leaves. In T. F. Preece & C. H. Dickinson (Eds.), Ecology of leaf surface micro-organisms (pp. 463-469). London & New York: Academic Press Inc. . Jiang, C. L., & Xu, L. H. (1996). Diversity of aquatic actinomycetes in lakes of the Middle Plateau, Yunnan, China. Applied and Environmental Microbiology, 62(1), 249- 253. Jiang, J. Y., He, X. F., & Cane, D. E. (2007). Biosynthesis of the earthy odorant geosmin by a bifunctional Streptomyces coelicolor enzyme. Nature Chemical Biology, 3(11), 711-715. Johnston, D. W., & Cross, T. (1976a). Actinomycetes in lake muds - dormant spores or metabolically active mycelium. Freshwater Biology, 6(5), 465-470. Johnston, D. W., & Cross, T. (1976b). Occurrence and distribution of actinomycetes in lakes of English Lake District. Freshwater Biology, 6(5), 457-463. Jones, G. J., & Korth, W. (1995). In-situ production of volatile odour compounds by river and reservoir phytoplankton populations in Australia. Water Science and Technology, 31(11), 145-151. Journey, C. A., Beaulieu, K. M., & Bradley, P. M. (2013). Environmental factors that influence cyanobacteria and geosmin occurrence in reservoirs. In P. M. Bradley (Ed.), Current perspectives in contaminant hydrology and water resources sustainability (pp. 27-55). Croatia: InTech. Judet-Correia, D., Bensoussan, M., Charpentier, C., & Dantigny, P. (2013). Influence of temperature, copper and CO2 on spore counts and geosmin production by 298
Penicillium expansum. Australian Journal of Grape and Wine Research, 19(1), 81-86. Juttner, F., & Watson, S. B. (2007). Biochemical and ecological control of geosmin and 2-methylisoborneol in source waters. Applied and Environmental Microbiology, 73(14), 4395-4406. Kampfer, P. (2006). The family Streptomycetaceae. Part I: Taxonomy. In M. Dworkin (Ed.), The prokaryotes: a handbook on the biology of bacteria (3rd ed., pp. 538- 604). Berlin: Springer. Kämpfer, P., & Wagner, M. (2002). Filamentous bacteria in activated sludge: current taxonomic status and ecology. In G. Bitton (Ed.), Encyclopedia of environmental microbiology (Vol. 3, pp. 1287-1306). New York: John Wiley and Sons Inc. Karandikar, A., Sharples, G. P., & Hobbs, G. (1997). Differentiation of Streptomyces coelicolor A3(2) under nitrate-limiting conditions. Microbiology, 143(11), 3581- 3590. Kendrick, K. E., & Ensign, J. (1983). Sporulation of Streptomyces grieus in submerged culture. Journal of Bacteriology, 155(1), 357-366. Kenefick, S. L., Hrudey, S. E., Prepas, E. E., Motkosky, N., & Peterson, H. G. (1992). Odorous substances and cyanobacterial toxins in prairie drinking water sources. Water Science and Technology, 25(2), 147-154. Khafif, T., Goetz, N., Coste, C. M., & Bastide, J. (1983). Sterilization thermique des sols et degradation des herbicides. Soil Biology & Biochemistry, 15(2), 205-209. Kikuchi, T., Kadota, S., Suehara, H., Nishi, A., & Tsubaki, K. (1981). Odourous metabolites of a fungus, Chaetomium globosum Kinze Ex Fr - Identification of geosmin, a musty-smelling compound. Chemical & Pharmaceutical Bulletin, 29(6), 1782-1784. Kikuchi, T., Mimura, T., Moriwaki, Y., Negoro, K., Nakazawa, S., & Ono, H. (1973). Metabolites of an Actinomyces Biwako-B strain, isolated from sediment in Lake Biwa - identification of odourous compounds, geosmin and 2-methylisoborneol, and analysis of fatty-acids. Yakugaku Zasshi-Journal of the Pharmaceutical Society of Japan, 93(5), 658-663. Klausen, C., Jørgensen, N. O. G., Burford, M., & O'Donahue, M. (2004). Actinomycetes may also produce taste and odour Water, 31, 45-49. Klausen, C., Nicolaisen, M. H., Strobel, B. W., Warnecke, F., Nielsen, J. L., & Jørgensen, N. O. G. (2005). Abundance of actinobacteria and production of geosmin and 2- methylisoborneol in Danish streams and fish ponds. Fems Microbiology Ecology, 52(2), 265-278. Koepsel, R., & Ensign, J. C. (1984). Microcycle sporulation of Streptomyces viridochromogenes Archives of Microbiology, 140(1), 9-14. Koksal, M., Chou, W. K. W., Cane, D. E., & Christianson, D. W. (2012a). Structure of 2-methylisoborneol synthase from Streptomyces coelicolor and implications for the cyclization of a noncanonical C-methylated monoterpenoid substrate. Biochemistry, 51(14), 3011-3020. Koksal, M., Chou, W. K. W., Cane, D. E., & Christianson, D. W. (2012b). Structure of geranyl diphosphate C-methyltransferase from Streptomyces coelicolor and implications for the mechanism of isoprenoid modification. Biochemistry, 51(14), 3003-3010. Komatsu, M., Tsuda, M., Omura, S., Oikawa, H., & Ikeda, H. (2008). Identification and functional analysis of genes controlling biosynthesis of 2-methylisoborneol. Proceedings of the National Academy of Sciences of the United States of America, 105(21), 7422-7427. 299
Kontro, M., Lignell, U., Hirvonen, M.-R., & Nevalainen, A. (2005). pH effects on 10 Streptomyces spp. growth and sporulation depends on nutrients. Letters in Applied Microbiology, 41(1), 32-38. Krishnani, K. K., Ravichandran, P., & Ayyappan, S. (2008). Microbially derived off- flavor from geosmin and 2-methylisoborneol: sources and remediation. Reviews of Environmental Contamination and Toxicology, 194, 1-27. Kuster, E., & Williams, S. T. (1964). Selection of Media for Isolation of Streptomycetes. Nature, 202(493), 928-929. Kutovaya, O. A., & Watson, S. B. (2014). Development and application of a molecular assay to detect and monitor geosmin-producing cyanobacteria and actinomycetes in the Great Lakes. Journal of Great Lakes Research, 40(2), 404-414. Lanciotti, E., Santini, C., Lupi, E., & Burrini, D. (2003). Actinomycetes, cyanobacteria and algae causing tastes and odours in water of the River Arno used for the water Supply of Florence. Journal of Water Supply Research and Technology-Aqua, 52(7), 489-500. Lange, B. M., Rujan, T., Martin, W., & Croteau, R. (2000). Isoprenoid biosynthesis: The evolution of two ancient and distinct pathways across genomes. Proceedings of the National Academy of Sciences of the United States of America, 97(24), 13172- 13177. Lange, C. L., & Wittmeyer, S. A. (1997). The contribution of Zebra Mussel (Dreissena spp.) feces and pseudofeces production to taste and odor episodes in the Niagara River and Lake Erie. In F. M. D'Itri (Ed.), Zebra Mussels and Aquatic Nuisance Species (pp. 225-243). Chelsea, MI: Ann Arbor Press. Larkin, M. A., Blackshields, G., Brown, N. P., Chenna, R., McGettigan, P. A., McWilliam, H., . . . Higgins, D. G. (2007). ClustalW and ClustalX version 2. Bioinformatics, 23(21), 2947-2948. Larsen, T. O., & Frisvad, J. C. (1995). Characterization of volatile metabolites from 47 Penicillium taxa. Mycological Research, 99(10), 1153-1166. Lawson, J. (2010). Factorial designs. In B. P. Carlin, J. J. Faraway, M. Tanner & J. Zidek (Eds.), Design and analysis of experiments with SAS (pp. 53-114). Boca Raton, FL: Chapman & Hall/CRC Press. Lechevalier, M. P. (1981). Ecological associations involving actinomycetes. Zentralblatt Fur Bakteriologie Mikrobiologie Hygiene I Abteilung, Suppl. 11, 159-166. Lee, G. C., Kim, Y. S., Kim, M. J., Oh, S. A., Choi, I., Choi, J., . . . Lee, C. H. (2011). Presence, molecular characteristics and geosmin producing ability of actinomycetes isolated from South Korean terrestrial and aquatic environments. Water Science and Technology, 63(11), 2745-2751. Li, H., Waiariabe, M., & Watanabe, M. M. (1997). Akinete formation in planktonic Anabaena spp. cyanobacteria by treatment with low temperature. Journal of Phycology, 33(4), 576-584. Lingappa, Y., & Lockwood, J. L. (1961). Chitin media for selective isolation and culture of actinomycetes. Nature, 189(475), 158-&. Lu, G., Edwards, C. G., Fellman, J. K., Mattinson, D. S., & Navazio, J. (2003). Biosynthetic origin of geosmin in red beets (Beta vulgaris L.). Journal of Agricultural and Food Chemistry, 51(4), 1026-1029. Ludwig, W., Euzéby, J., Schumann, P., Busse, H. J., Trujillo, M. E., Kämpfer, P., & Whitman, W. B. (2012). Road map of the Actinobacteria In M. Goodfellow, P. Kämpfer, H. J. Busse, M. E. Trujillo, K. I. Suzuki, W. Ludwig & W. B. Whitman (Eds.), Bergey’s Manual® of Systematic Bacteriology (Vol. Five The Actinobacteria, Part A, pp. 1-28). New York: Springer. 300
Lundstedt, T., Seifert, E., Lisbeth, A., Bernt, T., Nystrom, A., Pettersen, J., & Bergman, R. (1998). Experimental design and optimization. Chemometrics and Intelligent Laboratory Systems, 42(1), 3-40. Luti, K. J. K., & Mavituma, F. (2011a). Elicitation of Streptomyces coelicolor with dead cells of Bacillus subtilis and Staphylococcus aureus in a bioreactor increases production of undecylprodigiosin Applied Microbiology and Biotechnology, 90(2), 461-466. Luti, K. J. K., & Mavituma, F. (2011b). Elicitation of Streptomyces coelicolor with E.coli in a bioreactor enhances undecylprodiosin production. Biochemical Engineering Journal, 53(3), 281-285. Luti, K. J. K., & Mavituma, F. (2011c). Streptomyces coelicolor increases the production of undecylprodigiosin when interacted with Bacillus subtilis. Biotechnology Letters, 33(1), 113-118. Lylloff, J. E., Mogensen, M. H., Burford, M. A., Schlüter, L., & Jørgensen, N. O. G. (2012). Detection of aquatic streptomycetes by quantitative PCR for prediction of taste-and-odour episodes in water reservoirs. Journal of Water Supply Research and Technology-Aqua, 61(5), 272-282. Ma, X., Gao, N., Chen, B., Li, Q., Zhang, Q., & Gu, G. (2007). Detection of geosmin and 2-methylisoborneol by liquid-liquid extraction-gas chromatograph mass spectrum (LLE-GCMS) and solid phase extraction-gas chromatograph mass spectrum (SPE-GCMS). Frontiers of Environmental Science & Engineering in China, 1(3), 286-291. Mackay, S. J. (1977). Improved enumeration of Streptomyces spp. on a starch casein salt medium. Applied and Environmental Microbiology, 33(2), 227-230. Mahmoudi, N., Slater, G. F., & Fulthorpe, R. R. (2011). Comparison of commercial DNA extraction kits for isolation and purification of bacterial and eukaryotic DNA from PAH-contaminated soils. Canadian Journal of Microbiology, 57(8), 623-628. Makkar, N. S., & Cross, T. (1982). Actinoplanetes in soil and on plant litter from freshwater habitats. Journal of Applied Microbiology, 52(2), 209-218. Manteca, A., Alvarez, R., Salazar, N., Yague, P., & Sanchez, J. (2008). Mycelium differentiation and antibiotic production in submerged cultures of Streptomyces coelicolor. Applied and Environmental Microbiology, 74(12), 3877-3886. Maplestone, R. A., Stone, M. J., & Williams, D. H. (1992). The evolutionary role of secondary metabolites - a review. Gene, 115(1-2), 151-157. Marinelli, F., & Marcone, G. L. (2011). 3.26 Microbial Secondary Metabolites. In M. Moo-Young (Ed.), Comprehensive Biotechnology (2nd ed., Vol. Volume 3: Industrial Biotechnology and Commodity Products, pp. 285-297). Amsterdam, The Netherlands: Elsevier. Martin, J. F. (2004). Phosphate control of the biosynthesis of antibiotics and other secondary metabolites is mediated by the PhoR-PhoP system: an unfinished story. Journal of Bacteriology, 186(16), 5197-5201. Martin, J. F., Sola-Landa, A., Santos-Beneit, F., & Rodriguez, G., A. (2011). Network mechanisms of phosphate control of primary and secondary metabolism. In P. Dyson (Ed.), Streptomyces molecular biology and biotechnology (pp. 137-150). Norfolk, UK: Caister Academic Press. McCarthy, A. J., & Williams, S. T. (1992). Actinomycetes as agents of biodegradation in the environment - a review. Gene, 115(1-2), 189-192. Miller, A. L., & Walker, J. B. (1970). Accumulation of streptomycin-phosphate in cultures of streptomycin producers grown on a high-phosphate medium Journal of Bacteriology, 104(1), 8-12. 301
Mincer, T. J., Fenical, W., & Jensen, P. R. (2005). Culture-dependent and culture- independent diversity within the obligate marine actinomycete genus Salinispora. Applied and Environmental Microbiology, 71(11), 7019-7028. Mochida, K. (2009). Evaluation of the cytotoxicity of geosmin and 2-methylisoborneol using cultured human, monkey, and dog cells. Biocontrol Science, 14(1), 35-38. Moran, M. A., Rutherford, L. T., & Hodson, R. E. (1995). Evidence for indigenous Streptomyces populations in a marine-environment determined with a 16s ribosomal-RNA probe. Applied and Environmental Microbiology, 61(10), 3695- 3700. Morris, M. D. (2011). Two-level factorial experiments: basics. In B. P. Carlin, J. J. Faraway, M. Tanner & J. Zidek (Eds.), Design of experiments: an introduction based on linear models (pp. 187-206). Boca Raton, FL: Chapman & Hall/CRC Press. Morris, R. L. (1962). Actinomycetes studies as taste and odor cause. Water & Sewage Works, 109(2), 76-77. Naes, H., Utkilen, H. C., & Post, A. F. (1988). Factors influencing geosmin production by the cyanobacterium Oscillatoria brevis. Water Science and Technology, 20(8- 9), 125-131. Nakajima, M., Ogura, T., Kusama, Y., Iwabuchi, N., Imawaka, T., Araki, A., . . . Sunairi, M. (1996). Inhibitory effects of odour substances, geosmin and 2- methylisoborneol, on early development of sea urchins. Water Research, 30(10), 2508-2511. Newcombe, G., & Cook, D. (2002). Influences on the removal of tastes and odours by PAC. Journal of Water Supply Research and Technology-Aqua, 51(8), 463-474. Nielsen, J. L., Klausen, C., Nielsen, P. H., Burford, M., & Jørgensen, N. O. G. (2006). Detection of activity among uncultured Actinobacteria in a drinking water reservoir. Fems Microbiology Ecology, 55(3), 432-438. Niemi, R. M., Knuth, S., & Lundstrom, K. (1982). Actinomycetes and fungi in surface waters and in potable water. Applied and Environmental Microbiology, 43(2), 378-388. Nowak, A., & Wronkowska, H. (1987). On the efficiency of soil sterilization by autoclave. Zentralblatt Fur Mikrobiologie, 142(7), 521-525. O’Brien, J., & Wright, G. D. (2011). An ecological perspective of microbial secondary metabolism. Opinion in Biotechnolog, 22(4), 552–558. Ogura, T., Sunairi, M., & Nakajima, M. (2000). 2-methylisoborneol and geosmin, the main sources of soil odor, inhibit the germination of Brassicaceae seeds. Soil Science and Plant Nutrition, 46(1), 217-227. Onaka, H., Mori, Y., Igarashi, Y., & Furumai, T. (2011). Mycolic acid-containing bacteria induce natural-product biosynthesis in Streptomyces species. Applied and Environmental Microbiology, 77(2), 400-406. Pan, Y., Xu, L. P., Cao, W. X., Yin, S. R., Wang, Z. W., & Zhou, Q. H. (2009). Actinomycetes and earthy-musty odourous compounds in brackish fishponds in Tianjin, China. Water Science and Technology, 59(6), 1185-1194. Parinet, J., Rodriguez, M. J., & Serodes, J. (2010). Influence of water quality on the presence of off-flavour compounds (geosmin and 2-methylisoborneol). Water Research, 44(20), 5847-5856. Parinet, J., Rodriguez, M. J., & Serodes, J. B. (2013). Modelling geosmin concentrations in three sources of raw water in Quebec, Canada. Environmental Monitoring and Assessment, 185(1), 95-111.
302
Park, D.-K., Maeng, J., Ahn, C.-Y., Chung, A.-S., Lee, J.-H., & Oh, H.-M. (2001). Geosmin concentration and its relation to environmental factors in Daechung Reservoir, Korea. Korean Journal of Limonology, 34(4), 319-326. Perez, J., Munoz-Dorado, J., Brana, A. F., Shimkets, L. J., Sevillano, L., & Santamaria, R. I. (2011). Myxococcus xanthus induces actinorhodin overproduction and aerial mycelium formation by Streptomyces ceolicolor Microbial Biotechnology, 4(2), 175-183. Persson, P.-E., & Sivonen, K. (1979). Notes on muddy odour .V. actinomycetes as contributors to muddy odour in water. Aqua fenn, 9, 57-61. Persson, P. E. (1995). 19th-century and early-20th-century studies on aquatic off-flavors: A historical review. Water Science and Technology, 31(11), 9-13. Peter, A., Koster, O., Schildknecht, A., & von Gunten, U. (2009). Occurrence of dissolved and particle-bound taste and odor compounds in Swiss lake waters. Water Research, 43(8), 2191-2200. Qi, M., Chen, J., Sun, X., Deng, X., Niu, Y., & Xie, P. (2012). Development of models for predicting the predominant taste and odour compounds in Taihu Lake, China. Plos One, 7(12), 1-10. Ramsay, A. J., & Bawden, A. D. (1983). Effects of sterilization and storage on respiration, nitrogen status and direct counts of soil bacteria using acridine organge. Soil Biology & Biochemistry, 15(3), 263-268. Raschke, R. F., Carroll, B., & Tebo, L. B. (1975). The relationship between substrate content, water quality, actinomycetes and musty odours in the Broad River basin. Journal of Applied Ecology, 12(2), 535-560. Rastogi, G., & Sani, R. K. (2011). Molecular techniques to assess microbial community structure, function and dynamics in the environment. In I. Ahmad, F. Ahmad & J. Pichtel (Eds.), Microbes and Microbial Technology: Agricultural and Environmental Applications (pp. 29-57). New York: Springer Science+Business Media LLC. Rateb, M. E., Hallyburton, I., Houssen, W. E., Bull, A. T., Goodfellow, M., Santhanam, R., . . . Ebel, R. (2013). Induction of diverse secondary metabolites in Aspergillus fumigatus by microbial co-culture. Rsc Advances, 3(34), 14444–14450. Rezanka, T., & Votruba, J. (1998). Effect of salinity on the formation of avermectins, odour compounds and fatty acids by Streptomyces avermitilis. Folia Microbiologica, 43(1), 47-50. Ridal, J. J., Watson, S. B., & Hickey, M. B. C. (2007). A comparison of biofilms from macrophytes and rocks for taste and odour producers in the St. Lawrence River. Water Science and Technology, 55(5), 15-21. Rintala, H., & Nevalainen, A. (2005). Quantitative measurement of streptomycetes using real-time PCR. Journal of Environmental Monitoring, 8(7), 745-749. Romano, A. H., & Safferman, R. S. (1963). Studies on actinomycetes and their odours. Journal American Water Works Association, 55(2), 169-176. Rosen, B. H., Macleod, B. W., & Simpson, M. R. (1992). Accumulation and release of geosmin during the growth phases of Anabaena circinalis (Kutz) Rabenhorst. Water Science and Technology, 25(2), 185-190. Rowan, D. D. (2011). Volatile metabolites. Metabolites, 1(1), 41-63. Ruiz, B., Chavez, A., Forero, A., Garcia-Huante, Y., Romero, A., Sanchez, M., . . . Langley, E. (2010). Production of microbial secondary metabolites: regulation by carbon source. Critical Reviews in Microbiology, 36(2), 146-167.
303
Saadoun, I. (2005). Production of 2-methylisoborneol by Streptomyces violaceusniger and its transformation by selected species of Pseudomonas. Journal of Basic Microbiology, 45(3), 236-242. Saadoun, I. M. K., Schrader, K. K., & Blevins, W. T. (2001). Environmental and nutritional factors affecting geosmin synthesis by Anabaena sp. Water Research, 35(5), 1209-1218. Saitou, N., & Nei, M. (1987). The neighbor-joining method: a new method for reconstructing phylogenetic trees. Molecular Biology and Evolution, 4(4), 406- 425. Sakr, A. A., Ghaly, M. F., & Ali, M. F. (2013). The relationship between salts and growth of Streptomyces isolated from mural paintings in some ancient Egyptian tombs. Conservation Science in Cultural Heritage, 13, 313-330. Schafer, J., Jackel, U., & Kampfer, P. (2010). Development of a new PCR primer system for selective amplification of Actinobacteria. Fems Microbiology Letters, 311(2), 103-112. Schlatter, D. C., Samac, D. A., Tesfaye, M., & Kinkel, L. L. (2010). Rapid and specific method for evaluating Streptomyces competitive dynamics in complex soil communities. Applied and Environmental Microbiology, 76(6), 2009-2012. Schlumpberger, B. O. (2002). Dehydrogeosmin produzierende Kakteen: Untersuchungen zur Verbreitung, Duftstoff-Produktion und Besta¨ubung. (Doctor of Philisophy PhD dissertation), University of Tu¨ bingen, Grauer. Schlumpberger, B. O., Jux, A., Kunert, M., Boland, W., & Wittmann, D. (2004). Musty- earthy scent in cactus flowers: Characteristics of floral scent production in dehydrogeosmin-producing cacti. International Journal of Plant Sciences, 165(6), 1007–1015. Scholler, C. E. G., Gurtler, H., Pedersen, R., Molin, S., & Wilkins, K. (2002). Volatile metabolites from actinomycetes. Journal of Agricultural and Food Chemistry, 50(9), 2615-2621. Schrader, K. K., & Blevins, W. T. (1999). Effects of selected environmental conditions on biomass and geosmin production by Streptomyces halstedii. Journal of Microbiology, 37(3), 159-167. Schrader, K. K., & Blevins, W. T. (2001). Effects of carbon source, phosphorus concentration, and several micronutrients on biomass and geosmin production by Streptomyces halstedii. Journal of Industrial Microbiology & Biotechnology, 26(4), 241-247. Schrader, K. K., & Summerfelt, S. T. (2010). Distribution of off-flavor compounds and isolation of geosmin-producing bacteria in a series of water recirculating systems for rainbow trout culture North American Journal of Aquaculture, 72(1), 1-9. Schulz, S., Dickschat, J. S., Kunze, B., Wagner-Dobler, I., Diestel, R., & Sasse, F. (2010). Biological activity of volatiles from marine and terrestrial bacteria. Marine Drugs, 8(12), 2976-2987. Schulz, S., Fuhlendorff, J., & Reichenbach, H. (2004). Identification and synthesis of volatiles released by the myxobacterium Chondromyces crocatus. Tetrahedron, 60(17), 3863-3872. Schummer, C., Delhomme, O., Appenzeller, B. M. R., Wennig, R., & Millet, M. (2009). Comparison of MTBSTFA and BSTFA in derivatization reactions of polar compounds prior to GC/MS analysis. Talanta, 77(4), 1473-1482. Sekar, R., Pernthaler, A., Pernthaler, J., Warnecke, F., Posch, T., & Amann, R. (2003). An improved protocol for quantification of freshwater Actinobacteria by
304
fluorescence in situ hybridization. Applied and Environmental Microbiology, 69(5), 2928-2935. Sevcikova, B., & Kormanec, J. (2004). Differential production of two antibiotics of Streptomyces coelicolor A3(2), actinorhodin and undecylprodigiosin, upon salt stress conditions. Archives of Microbiology, 181(5), 384–389. Sigworth, E. A. (1957). Control of odor and taste in water supplies. Journal American Water Works Association, 49(12), 1507-1521. Silvey, J. K. G. (1963). Quality aspects of water distribution systems: actinomycetes in water distribution systems. Paper presented at the Proc. 5th Sanitary Eng. Conf, University of Illinois, England Silvey, J. K. G., & Roach, A. W. (1953). Actinomycetes in the Oklahoma City water supply. Journal American Water Works Association, 45(4), 409-416. Silvey, J. K. G., & Roach, A. W. (1975). The taste and odour producing aquatic actinomycetes. CRC Critical Reviews in Environmental Control, 5(2), 233-273. Silvey, J. K. G., Russell, J. C., Redden, D. R., & McCormick, W. C. (1950). Actinomycetes and common tastes and odours. Journal American Water Works Association, 42(11), 1018-1026. Singh, B., Oh, T. J., & Sohng, J. K. (2009). Exploration of geosmin synthase from Streptomyces peucetius ATCC 27952 by deletion of doxorubicin biosynthetic gene cluster. Journal of Industrial Microbiology & Biotechnology, 36(10), 1257- 1265. Singh, K. P., Malik, A., Basant, N., & Saxena, P. (2007). Multi-way partial least squares modeling of water quality data. Analytica Chimica Acta, 584(2), 385-396. Sivonen, K. (1982). Factors influencing odor production by actinomycetes. Hydrobiologia, 86(1-2), 165-170. Smith, C. J., & Osborn, A. M. (2008). Advantages and limitations of quantitative PCR (Q-PCR)-based approaches in microbial ecology. Fems Microbiology Ecology, 67(1), 6-20. Sohler, A., Romano, A. H., & Nickerson, W. J. (1958). Biochemistry of the actinomycetales. III Cell wall composition and the action of lysozyme upon cells and cell walls of the actinomycetales. Journal of Bacteriology, 75(3), 283-290. Spiteller, D., Jux, A., Piel, J., & Boland, W. (2002). Feeding of [5,5-H-2(2)]-1-desoxy- D-xylulose and [4,4,6,6,6-H-2(5)]-mevalolactone to a geosmin-producing Streptomyces sp and Fossombronia pusilla. Phytochemistry, 61(7), 827-834. Sporle, J., Becker, H., Allen, N. S., & Gupta, M. P. (1991). Occurrence of (-) geosmin and other terpenoids in an axenic culture of the liverwort Symphyogyna brongiartin. Z. Naturforsch., 46, 183-188. Srinivasan, R., & Sorial, G. A. (2011). Treatment of taste and odour causing compounds 2-methyl isoborneol and geosmin in drinking water: a critical review. Journal of Environmental Sciences-China, 23(1), 1-13. Stackebrandt, E., & Goebel, B. M. (1994). Taxonomic note: a place for DNA-DNA reassociation and 16S rRNA sequence analysis in the present species definition in bacteriology. International Journal of Systematic Bacteriology, 44(4), 846-849. Stahl, P. D., & Parkin, T. B. (1994). Purge-and-trap extraction of geosmin and 2- methylisoborneol from soil. Soil Science Society of America Journal, 58(4), 1163- 1167. Stensmyr, M. C., Dweck, H. K. M., Farhan, A., Ibba, I., Strutz, A., Mukunda, L., . . . Hansson, B. S. (2012). A conserved dedicated olfactory circuit for detecting harmful microbes in Drosophila Cell, 151(6), 1345–1357.
305
Su, M., Gaget, V., Giglio, S., Burch, M., An, W., & Yang, M. (2013). Establishment of quantitative PCR methods for the quantification of geosmin-producing potential and Anabaena sp. in freshwater systems. Water Research, 47(10), 3444-3454. Sugiura, N., Inamori, Y., Hosaka, Y., Sudo, R., & Takahashi, G. (1994). Algae enhancing musty odour production by actinomycetes in Lake Kasumigaura. Hydrobiologia, 288(1), 57-64. Sugiura, N., & Nakano, K. (2000). Causative microorganisms for musty odour occurrence in the eutrophic Lake Kasumigaura. Hydrobiologia, 434(1-3), 145-150. Sugiura, N., Utsumi, M., Wei, B., Norio, I., Okano, K., Kawachi, Y., & Maekawa, T. (2004). Assessment for the complicated occurence of nuisance odours from phytoplankton and environmental factors in a eutrophic lake Lakes and Reservoirs: Research and Management, 9, 195-201. Sugiura, N., Yagi, O., & Sudo, R. (1987). Effect of carbohydrate in the sediment on the musty odour production by actinomycetes. Environmental Technology Letters, 8(2), 95-103. Sun, D. L., Yu, J. W., An, W., Yang, M., Chen, G. G., & Zhang, S. J. (2013). Identification of causative compounds and microorganisms for musty odor occurrence in the Huangpu River, China. Journal of Environmental Sciences-China, 25(3), 460- 465. Sunesson, A. L., Nilsson, C. A., Carlson, R., Blomquist, G., & Andersson, B. (1997). Production of volatile metabolites from Streptomyces albidoflavus cultivated on gypsum board and tryptone glucose extract agar: Influence of temperature, oxygen, and cardon dioxide levels The Annuals of Occupational Hygeine, 41(4), 393-413. Suurnakki, S., Gomez-Saez, G. V., Rantala-Ylinen, A., Jokela, J., Fewer, D. P., & Sivonen, K. (2015). Identification of geosmin and 2-methylisoborneol in cyanobacteria and molecular detection methods for the producers of these compounds Water Research, 68(1), 56–66. Suutari, M. H., Rintala, H., Ronka, E., & Nevalainen, A. (2002). Oligonucleotide primers for detection of Streptomyces species. World Patent No. WO 02/092848 A1. World Intellectual Property Organisation Takahashi, Y., Iwai, Y., & Omura, S. (1991). Mode of submerged spore formation in Kitasatosporia setae. Journal of General and Applied Microbiology, 37(3), 261- 266. Takahashi, Y., Kuwana, T., Iwai, Y., & Omura, S. (1984). Some characteristics of aerial and submerged spores of Kitasatosporia setalba. Journal of General and Applied Microbiology, 30(3), 223-229. Takizawa, M., Colwell, R. R., & Hill, R. T. (1993). Isolation and diversity of actinomycetes in the Chesapeake Bay. Applied and Environmental Microbiology, 59(4), 997-1002. Tamura, K., Nei, M., & Kumar, S. (2004). Prospects for inferring very large phylogenies by using the neighbor-joining method. Proceedings of the National Academy of Sciences (USA), 101(30), 11030-11035. Tamura, K., Stecher, G., Peterson, D., Filipski, A., & Kumar, S. (2013). MEGA6: Molecular evolutionary genetics analysis version 6.0. Molecular Biology and Evolution, 30(12), 2725-2729. Tarkka, M., & Hampp, R. (2008). Secondary metabolites of soil Streptomycetes in biotic interactions. In P. Karlovsky (Ed.), Secondary metabolites in soil ecology (pp. 107-126). Berlin, Heidelberg: Springer-Verlag.
306
Thaysen, A. C. (1936). The origin of an earthy or muddy taint in fish. Annals of Applied Biology, 23(1), 99-104. Thermo Fisher Scientific. (2013). Assessment of Nucleic Acid Purity. T042-Technical Bulletin Nano Drop Spectrophotometers. Retrieved 26th September 2013, from http://www.nanodrop.com/Library/T042-NanoDrop-Spectrophotometers- Nucleic-Acid-Purity-Ratios.pdf Tosi, L., & Sola, C. (1993). Role of geosmin, a typical inland water odour, in guiding Glass Eel Anguilla anguilla (L.) migration. Ethology, 95(3), 177-185. Tresner, H. D., Hayes, J. A., & Backus, E. J. (1968). Differential tolerance of streptomycetes to sodium chloride as a taxonomic aid. Applied Microbiology, 16(8), 1134-1136. Tsao, H.-W., Michinaka, A., Yen, H.-K., Giglio, S., Hobson, P., Monis, P. T., & Lin, T.- F. (2014a). Monitoring of geosmin producing Anabaena circinalis using quantitative PCR. Water Research, 49(1), 416-425. Tsao, H. W., Michinaka, A., Yen, H. K., Giglio, S., Hobson, P., Monis, P., & Lin, T. F. (2014b). Monitoring of geosmin producing Anabaena circinalis using quantitative PCR. Water Research, 49, 416-425. Tsujimura, S., & Okubo, T. (2003). Development of Anabaena blooms in a small reservoir with dense sediment akinete population, with special reference to temperature and irradiance Journal of Plankton Research, 25(9), 1059-1067. Tung, S. C., Lin, T. F., Tseng, I. C., & Lin, H. M. (2006). Identification of 2-MIB and geosmin producers in Feng-Shen reservoir in south Taiwan. Water Science & Technology: Water Supply, 6(2), 55-61. Umetrics. (2012). Introducion to multivariate data analysis User Guide to Simca-P+ (pp. 13-25). Sweden: Umetrics. Uwins, H. K. (2011). Triggers for taste and odour events: A study of microbial production of geosmin and 2-methylisoborneol. (Doctor of Philosophy Ph.D), Griffith University, Griffith University, Australia. Uwins, H. K., Teasdale, P., & Stratton, H. (2007). A case study investigating the occurrence of geosmin and 2-methylisoborneol (MIB) in the surface waters of the Hinze Dam, Gold Coast, Australia. Water Science and Technology, 55(5), 231- 238. van Keulen, G., Alderson, J., White, J., & Sawers, R. G. (2005a). Nitrate respiration in the actinomycete Streptomyces coelicolor. Biochemical Society Transactions, 33(1), 210-212. van Keulen, G., Alderson, J., White, J., & Sawers, R. G. (2007). The obligate aerobe actinomycete Streptomyces coelicolor A3(2) Environmental Microbiology, 9(12), 3143-3149. van Keulen, G., Hopwood, D. A., Dijkhuizen, L., & Sawers, R. G. (2005b). Gas vesicles in actinomycetes: old buoys in novel habitats? Trends in Microbiology, 13(8), 350-354. van Keulen, G., Jonkers, H. M., Claessen, D., Dijkhuizen, L., & Wosten, H. A. B. (2003). Differentiation and anaerobiosis in standing liquid cultures of Streptomyces coelicolor. Journal of Bacteriology, 185(4), 1455-1458. van Keulen, G., Siebring, J., & Dijkhuizen, L. (2011). Central carbon metabolic pathways in Streptomyces In P. Dyson (Ed.), Streptomyces: molecular biology and biotechnology (pp. 105-124). Northfolk, UK: Caster Academic Press. van Wezel, G. P., & McDowall, K. J. (2011). The regulation of the secondary metabolism of Streptomyces: new links and experimental advances Natural Product Reports, 28(7), 1311-1333. 307
Ventura, M., Canchaya, C., Tauch, A., Chandra, G., Fitzgerald, G. F., Chater, K. F., & van Sinderen, D. (2007). Genomics of actinobacteria: Tracing the evolutionary history of an ancient phylum. Microbiology and Molecular Biology Reviews, 71(3), 495-548. Vilalta, E., Guasch, H., Munoz, I., Romani, A., Valero, F., Rodriguez, J. J., . . . Sabater, S. (2004). Nuisance odours produced by benthic cyanobacteria in a Mediterranean river. Water Science and Technology, 49(9), 25-31. Vobis, G. (1997). Morphology of actinomycetes. In S. Miyadoh (Ed.), Atlas of actinomycetes (pp. 180-191). Japan: The Society for Actinomycetes Japan/Asakura Publishing Co., Ltd. Waksman, S. A. (1967). The actinomycetes: a summary of current knowledge. New York: The Ronald Press Company. Wang, C. L., Wang, Z. F., Qiao, X., Li, Z. J., Li, F. J., Chen, M. H., . . . Cui, H. Y. (2013a). Antifungal activity of volatile organic compounds from Streptomyces alboflavus TD-1. Fems Microbiology Letters, 341(1), 45-51. Wang, C. M., & Cane, D. E. (2008). Biochemistry and molecular genetics of the biosynthesis of the earthy odorant methylisoborneol in Streptomyces coelicolor. Journal of the American Chemical Society, 130(28), 8908-8909. Wang, Z., Wang, C., Li, F., Li, Z., Chen, M., Wang, Y., . . . Zhang, H. (2013b). Fumigant activity of volatiles from Streptomyces alboflavus TD-1 against Fusarium moniliforme Sheldon. Journal of Microbiology, 51(4), 477–483. Wang, Z., Xu, Y., Wang, J., & Renhui, L. (2011). Genes associated with 2- methylisoborneol biosynthesis in cyanobacteria: isolation, characterisation, and expression in response to light. Plos One, 6(4), e18665. Watson, S. B. (2003). Cyanobacterial and eukaryotic algal odour compounds: signals or by-products? A review of their biological activity. Phycologia, 42(4), 332-350. Watson, S. B., Brownlee, B., Satchwill, T., & Hargesheimer, E. E. (2000). Quantitative analysis of trace levels of geosmin and MIB in source and drinking water using headspace SPME. Water Research, 34(10), 2818-2828. Watson, S. B., Charlton, M., Rao, Y. R., Howell, T., Ridal, J., Brownlee, B., . . . Millard, S. (2007). Off flavours in large waterbodies: physics, chemistry and biology in synchrony. Water Science & Technology, 55(5), 1-8. Watson, S. B., & Ridal, J. (2004). Periphyton: a primary source of widespread and severe taste and odour. Water Science and Technology, 49(9), 33-39. Weete, J. D., Blevins, W. T., Wilt, G. R., & Durham, D. (1977). Chemical, biological and environmental factors responsible for the earthy odour in the Auburn City water supply. Alabama Agricultural Experiment Station Bulletin, 490, 46pp. Williams, S. T., & Davies, F. L. (1965). Use of antibiotics for selective isolation and enumeration of actinomycetes in soil. Journal of General Microbiology, 38(2), 251-261. Williams, S. T., Lanning, S., & Wellington, E. M. H. (1984). Ecology of actinomycetes. In M. Goodfellow, M. Mordarski & S. T. Williams (Eds.), The biology of the actinomycetes (pp. 481-528). London: Academic Press. Willoughby, L. G. (1969). A study of the aquatic actinomycetes of Blelham Tarn. Hydrobiologia, 34(3-4), 465-483. Willoughby, L. G. (1971). Observations on some aquatic actinomycetes of streams and rivers. Freshwater Biology, 1(1), 23-27. Willoughby, L. G. (1974). The activity of Streptomyces phase-virus in fresh water Hydrobiologia, 49(3), 215-228.
308
Willoughby, L. G. (1976). Freshwater actinomycetes. In E. B. G. Jones (Ed.), Recent advances in aquatic mycology (pp. 673-698). London: Elek Science. Willoughby, L. G., Smith, S. M., & Bradshaw, R. M. (1972). Actinomycete virus in freshwater. Freshwater Biology, 2(1), 19-26. Wilson, K. H., Blitchington, R. B., & Greene, R. C. (1990). Amplification of bacterial 16S ribosomal DNA with polymerase chain reaction. Journal of Clinical Microbiology, 28(9), 1942-1946. Wnorowski, A. U. (1992). Tastes and odours in the aquatic environment: A review. Water Sa, 18(3), 2O3-214. Wohl, D. L., & McArthur, J. V. (1998). Actinomycete-flora associated with submersed freshwater macrophytes. Fems Microbiology Ecology, 26(2), 135-140. Wold, S., Eriksson, L., & Kettaneh, N. (2010). PLS in data mining and data integration. In V. Esposito Vinzi, W. W. Chin, J. Henseler & H. Wang (Eds.), Handbook of partial least squares (pp. 327-358). Berlin: Springer. Wood, S., Williams, S. T., & White, W. R. (1983a). Microbes as a source of earthy flavours in potable water: A review International Biodeterioration & Biodegradation, 48(1-4), 26-40. Wood, S., Williams, S. T., & White, W. R. (1985). Potential sites of geosmin production by streptomycetes in and around reservoirs. Journal of Applied Bacteriology, 58(3), 319-326. Wood, S., Williams, S. T., White, W. R., & Jones, F. (1983b). Factors influencing geosmin production by Streptomyces and their relevance to the occurence of earthy taints in reservoirs. Water Science and Technology, 15, 191-198. Wu, J. T., Ma, P. I., & Chou, T. L. (1991). Variation of geosmin content in Anabaena cells and its relation to nitrogen-utilization. Archives of Microbiology, 157(1), 66- 69. Xie, Y. Q., He, J., Huang, J., Zhang, J. B., & Yu, Z. (2007). Determination of 2- methylisoborneol and geosmin produced by Streptomyces sp and Anabaena PCC7120. Journal of Agricultural and Food Chemistry, 55(17), 6823-6828. Yagi, O., Sugiura, N., & Sudo, R. (1987). Chemical and physical factors in the production of mustyodor by Streptomyces spp isolated from Lake Kasumigaura. Agricultural and Biological Chemistry, 51(8), 2081-2088. Yague, P., Rodriguez-Garcia, A., Lopez-Garcia, M. T., Martin, J. F., Rioseras, B., Sanchez, J., & Manteca, A. (2013). Transcriptomic analysis of Streptomyces coelicolor differentiation in solid sporulating cultures: First compartmentalized and second multinucleated mycelia have different and distinctive transcriptomes. Plos One, 8(3). Yamamoto, Y., Tanaka, K., & Komori, N. (1994). Volatile compounds excreted by myxobacteria isolated from lake water and sediments. Japanese Journal of Limnology, 55(4), 241-245. Ye, J., Coulouris, G., Zaretskaya, I., Cutcutache, I., Rozen, S., & Madden, T. L. (2012). Primer-BLAST: a tool to design target-specific primers for polymerase chain reaction. BMC Bioinformatics, 13(1), 134. Zacheus, O. M., Lehtola, M. J., Korhonen, L. K., & Martikainen, P. J. (2001). Soft deposits, the key site for microbial growth in drinking water distribution networks. Water Research, 35(7), 1757-1765. Zaitlin, B., & Watson, S. B. (2006). Actinomycetes in relation to taste and odour in drinking water: Myths, tenets and truths. Water Research, 40(9), 1741-1753.
309
Zaitlin, B., Watson, S. B., Dixon, J., & Steel, D. (2003a). Actinomycetes in the Elbow River Basin, Alberta, Canada. Water Quality Research Journal of Canada, 38(1), 115-125. Zaitlin, B., Watson, S. B., Ridal, J., Satchwill, T., & Parkinson, D. (2003b). Actinomycetes in Lake Ontario: Habitats and geosmin and MIB production. Journal American Water Works Association, 95(2), 113-118. Zhang, T., Li, L., Song, L. R., & Chen, W. (2009). Effects of temperature and light on the growth and geosmin production of Lyngbya kuetzingii (Cyanophyta). Journal of Applied Phycology, 21(3), 279-285. Zhang, T., Li, L., Zuo, Y. X., Zhou, Q., & Song, L. R. (2010). Biological origins and annual variations of earthy-musty off-flavours in the Xionghe Reservoir in China. Journal of Water Supply Research and Technology-Aqua, 59(4), 243-254. Zhi, X. Y., Li, W. J., & Stackebrandt, E. (2009). An update of the structure and 16S rRNA gene sequence-based definition of higher ranks of the class Actinobacteria, with the proposal of two new suborders and four new families and emended descriptions of the existing higher taxa. International Journal of Systematic and Evolutionary Microbiology, 59(3), 589-608. Zuo, Y., Li, L., Wu, Z., & Song, L. (2009a). Isolation, identification and odour-producing abilities of geosmin/2-MIB in actinomycetes from sediments in Lake Lotus, China Journal of Water Supply Research and Technology-Aqua, 58(8), 552-561. Zuo, Y. X., Li, L., Wu, Z. X., & Song, L. R. (2009b). Isolation, identification and odour- producing abilities of geosmin/2-MIB in actinomycetes from sediments in Lake Lotus, China. Journal of Water Supply Research and Technology-Aqua, 58(8), 552-561. Zuo, Y. X., Li, L., Zhang, T., Zheng, L. L., Dai, G. Y., Liu, L. M., & Song, L. R. (2010). Contribution of Streptomyces in sediment to earthy odour in the overlying water in Xionghe Reservoir, China. Water Research, 44(20), 6085-6094.
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APPENDICES
APPENDIX A: Partial least squares (PLS) background information
PLS creates a multivariate regression model by projecting independent (X) and dependent
(Y) variables into a new space using a principal components (latent variable) approach to model the covariance structure between two matrices (X and Y) and maximise the correlation between X and Y. The response block is represented by the Y scores, U, while the predictive block is described by X scores, T. PLS maximises the covariance between
U and T by decomposing Y and X as shown by the equations below:
X = TWT+ EX and
Y = UCT + EY such that the scores in X (T) and the scores of the yet unexplained part of Y (U) have maximum covariance. W and C are X and Y PLS loadings (weights) respectively. EX and EY are the X and Y residuals respectively. The decomposition models of X and Y and the expression relating these models through regression constitute the linear PLS regression model. The extracted X-scores are used to predict Y-scores which in turn are used to construct predictions for the responses (Singh et al., 2007).
The weight vectors of each model dimension (component) express how the X-variables are combined to form T and the Y variables to form U. The data are modelled as a set of factors in X and Y and their relationships. Loadings are the contribution of each of the variables to the PLS factor. The PLS procedure is described in detail by Geladi and
Kowalksi (1986).
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APPENDIX B: Comparison between cell lysis methods for DNA extraction
Table B.1 DNA concentrations measured from Streptomyces spp. spore and vegetative cell suspensions and a garden soil sample using either the cellular disruption protocol of the Power Soil DNA Isolation Kit® or the developed enzymatic-chemical-heat lysis technique. The values represent the mean ± standard error based on three replications. Crude DNA resulting from both lysis procedures were purified using the Power Soil DNA Isolation Kit® purification protocol. The DNA purity ratios for each are shown.
Cell lysis DNA Species DNA yield DNA purity method Source (ng/µL) (A260/A280) Spores S. antibioticus 47.80 ±.0.80 1.82±0.03 Power Soil DNA S. coelicolor Isolation Kit® A3(2) 20.35 ± 1.63 1.74±0.12 (mechanical- Vegetative S. antibioticus 4.65±0.15 1.74±0.19 chemical-heat). cells S. coelicolor A3(2) 22.13±2.36 1.68±0.09 Garden Soil 14.73±1.27 1.50±0.13 Spores S. antibioticus 1.65±0.55 1.34 ±0.48 Enzymatic- S. coelicolor chemical-heat A3(2) 4.73±2.36 1.57±0.16 Vegetative S. antibioticus 3.25 ±0.45 1.60 ±0.10 cells S. coelicolor A3(2) 16.73 ±0.60 1.75 ±0.04 Garden Soil 3.57±0.25 1.58±0.15
The values of the A260/A280 ratio indicated that the extracted DNA was in general acceptably pure, although the ratios were lower than generally accepted for the soil DNA extracts.
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APPENDIX C: Melting curve analysis for qPCR assays
Figure C.1 melting curve analysis of PCR products using Streptomyces DNA after qPCR using the 16S rRNA Streptomyces-specific primer pair. The Tm value of the single peak was identified to be 84°C
Figure C.2 melting curve analysis of PCR products using Streptomyces DNA after qPCR using the 16S rRNA actinobacteria-specific primer pair. The Tm value of the single peak was identified to be 86.5°C.
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APPENDIX D: Colony forming units and qPCR-determined abundance of Streptomyces and actinobacteria in environmental samples
Table D.1 colony forming units (CFU) per gram or liter of Streptomyces obtained using selective isolation agar in Grahamstown Reservoir during the wet and dry sampling period. Values represent the mean ± SE (n=3). ND = not detected. Starch casein agar Actinomycete isolation agar Sample ID Wet Dry Wet (February) Dry (February) (November) (November) Soil R2 853± 0 91,408± 0 5,973± 0 25,137±2,500 Sediment R2 3,007±430 7,752±1,723 7,302±3,500 16,796±500 Soil R6 3,534± 0 3,810± 952 31,360± 500 571± 0 Sediment R6 6,791±3,396 1,918± 0 10,798± 6,500 ND Soil W 4,837±440 11,719± 0 8,355±1,500 1,465± 500 Sediment W 8,513± 448 5,665±730 7,168± 0 1,789± 0
Marginal H20 R2 15,500± 1,500 3,000± 1,000 10,000 ± 3,000 8,000± 4,000
Marginal H20 R6 3, 000 ± 0 ND 1,000 ± 160 2,000 ± 0
Marginal H20 W 11,000± 1,000 ND 3,000± 1,000 ND Macrophyte R2 1,700± 300 ND 1,650± 350 ND Torpedo Grass R6 ND ND 600± 0 ND Macrophyte W ND ND ND ND
Surface H20 R2 8,000± 2,000 4,000±1,414 4,000 ± 2,000 3,000± 0
Bottom H20 R2 670 ± 30 4,000 ± 0 180 ± 60 ND Bottom sediment R2 152± 76 2,147±907 284 ± 205 1,247± 0
Surface H20 R6 3,000± 0 6,667± 0 733± 20 ND
Bottom H20 R6 30 ± 10 3,000± 1,000 20 ± 10 3,000± 1,000 Bottom sediment R6 385± 43 485±244 184± 85 171± 100
Surface H20 R12 4,000± 0 ND 390± 70 ND
Bottom H20 R12 3,000± 1,000 ND 57,500±500 ND Bottom sediment R12 93± 50 61± 11 160 ± 75 43 ± 0
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Table D.2 Colony forming units (CFU) per gram or liter of Streptomyces obtained using selective isolation agar in Chichester Reservoir during the wet and dry sampling period. Values represent the mean ± SE (n=3). ND = not detected. Starch casein agar Actinomycete isolation agar Sample ID Wet (April) Dry (October) Wet (April) Dry (October) Soil (south) 72,046±4,803 74,928± 28,818± 0 96,061± 0 11,527 Sediment (south) 9,662±966 53,140±4,831 63,124±18,652 11,594± 0 Soil (north) 39,914±3,004 10,300±3,433 31,502±0 22,318±1,717 Sediment (north) 1,719±524 23,214±8,929 4,018±446 8,929 ± 0 Debris (south) 17,686±478 267,686±38,24 21,989±1,912 53,537± 0 1 Debris (north) ND 3,413± 0 2,560±853 30,717± 0 Macrophyte 1 (south) ND ND 105±5 ND Macrophyte 2 (south) 527±300 ND 100± 0 ND Algae (south) 500± 0 ND 400 ± 0 ND
Marginal H20 (south) 0 ± 0 11,500±6,500 2,000± 0 5,000±2,000
Marginal H20 (north) 1,000± 0 11,500±1,500 0 ± 0 9,500±500
Surface H20 (south) 2,000± 0 1,000± 0 1000 ± 0 3,000± 0
Bottom H20 (south) 2,000± 0 0 ± 0 1,000± 0 2000 ± 0 Bottom sediment (south) 13,592± 0 1,942± 0 11,650± 0 518± 0
Surface H20 (north) ND ND 140± 0 1,000± 0
Bottom H20 (north) 1,080± 0 1,000± 0 ND 1,000± 0 Bottom sediment (north) 1,202±168 1,096±0 2,740±685 2,603±0
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Table D.3 Grahamstown Reservoir dry sampling period. Total cells refer to qPCR results determined using DNA extracted from samples using the vigorous PowerSoil lysis technique. Vegetative cells refers to qPCR results for DNA obtained using the enzymatic- chemical lysis approach. Values represent the mean ± SE (n=6). ND = not detected Streptomyces-16SrRNA Actinobacteria- 16SrRNA Sample ID Total cells Vegetative Cells Total cells Soil (R2) 2,034,284±71,564 1,223,129±256,553 73,453,017±9,118,479 Sediment (R2) 172,909±47,772 171,429±13,867 93,843,325±7,590,374 Soil (R6) 147,401±12,027 13,283±1,344 37,248,115±3,507,206 Sediment (R6) 94,695±8,358 51,610±22,767 28,719,011±3,086,358 Soil (W) 86,027±7,667 23,430±3,813 30,157,304±3,327,091 Sediment (W) 116,197±14,721 18,308±4,685 16,102,021±1,196,984 Macrophyte (R2) 1,790±266 1,787±286 3,281,135±343,922 Torpedo grass (R6) 17,478±909 13,911±1,397 1,471,387±167,772 Torpedo grass (W) 11,856±639 5,402±855 1,212,795±365,307 Macrophyte (W) 2,360±741 ND 497,379±161,725 Bottom sediment 15,007±3,331 2,574±275 (R2) 5,972,879±564,240 Bottom sediment 7,848±1,551 205±82 (R6) 13,299,069±2,342,466 Bottom sediment 126,080±12,815 24,474±2,139 (R12) 3,708,198±630,794 Marginal H20 (R2) 9,255±1,183 3,134±1,158 199,223,758±14,144,100 Marginal H20 (R6) 2,286±212 1,289±139 165,529,958±5,999,119 Marginal H20 (W) 1,829±161 233±50 36,500,427±2,154,891 Surface H20 (R2) 11,311±1,282 10,602±928 144,395,168±44,873,259 Bottom H20 (R2) 43,641±3,142 26,027±1,327 216,552,575±10,313,121 Surface H20 (R6) 29,469±1,117 14,008±757 175,649,117±27,119,592 Bottom H20 (R6) 5,739±805 5,596±421 192,177,408±21,408,812 Surface H20 (R12) 2,699±219 1,171±144 75,016,630±28,006,006 Bottom H20 (R12) 6,472±340 2,661±259 133,356,161±17,182,214
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Table D.4 Grahamstown Reservoir wet sampling period. Total cells refer to qPCR results determined using DNA extracted from samples using the vigorous PowerSoil lysis technique. Vegetative cells refers to qPCR results for DNA obtained using the enzymatic- chemical lysis approach. Values represent the mean ± SE (n=6). ND = not detected Streptomyces-16SrRNA Actinobacteria- 16SrRNA Sample ID Total cells Vegetative Cells Total cells Soil (R2) 485,205±178,103 27,881±1,690 58,462,922±37,984,777 Sediment (R2) 137,463±19,878 37,651±7,757 43,219,302±13,429,988 Soil (R6) 78,105±8,218 13,079±1,388 20,380,994±4,933,564 Sediment (R6) 195,159±27,483 7,985±1,943 30,434,890±9,890,192 Soil (W) 118,255±51,490 341±37 17,868,028±9,808,423 Sediment (W) 436,917±89,641 16±2 13,388,057±4,174,918 Macrophyte (R2) 3,949±367 ND 2,588,849±264,242 Torpedo grass (R6) 10,077±1,835 43±27 3,153,984±499,737 Torpedo grass (W) 201,771±28,463 347±135 10,671,373±865,016 Macrophyte (W) 3,868±461 283±63 577,507±191,134 Bottom sediment (R2) 54,602±2,971 10,273±764 24,565,999±4,444,719 Bottom sediment (R6) 8,547±2,800 391±172 11,209,766±3,626,164 Bottom sediment (R12) 52,803±18,671 286±2 35,029,884±9,312,711 Marginal H20 (R2) 52,335±6,783 1,342±322 21,447,622±4,488,539 Marginal H20 (R6) 673,561±47,195 1,955±115 19,843,458±7,698,176 Marginal H20 (W) 185,644±9,259 1,628±142 3,360,750±496,028 Surface H20 (R2) 134,419±6,175 1,581±165 20,480,436±4,694,549 Bottom H20 (R2) 121,118±8,496 11,868±709 11,008,700±921,768 Surface H20 (R6) 572,971±57,676 865±133 7,394,503±2,137,070 Bottom H20 (R6) 256,129±21,648 265±35 17,290,594±1,307,434 Surface H20 (R12) 87,066±11,080 2,173±135 8,697,830±1,065,941 Bottom H20 (R12) 846,079±47,017 624±81 14,144,054±1,152,952
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Table D.5 Chichester Reservoir dry sampling period. Total cells refer to qPCR results determined using DNA extracted from samples using the vigorous PowerSoil lysis technique. Vegetative cells refers to qPCR results for DNA obtained using the enzymatic- chemical lysis approach. Values represent the mean ± SE (n=6). ND = not detected Streptomyces-16SrRNA Actinobacteria- 16SrRNA Sample ID Total cells Vegetative Cells Total cells Soil (south) 792,900±136,504 237,481±7,785 7,752,403±1,749,019 Sediment (south) 2,439,088±422,129 824,039±21,235 35,574,061±4,552,174 Soil (north) 10,944,850±2,461,276 9,205,261±1,042,585 83,670,196±13,893,927 Sediment (north) 446,960±180,181 109,153±4,389 8,163,720±687,086 Debris (south) 991,878±47,348 783,141±19,544 32,365,656±1,653,488 Debris (north) 503,208±152,802 383,600±17,470 11,759,099±501,072 Macrophyte 1 10,723±1,098 1,137±131 129,564,900±6,077,038 (south) Macrophyte 2 16,497±404 ND 152,159,063±3,570,222 (south) Algae (south) 138,896±37,145 34,546±1,820 14,013,377±596,841 Bottom sediment 130,738±37,746 2,714±407 5,958,144±172,540 (south) Bottom sediment 48,769±14,159 2,386±377 1,912,040±62,324 (north) Marginal H20 52,004±2,150 40,5992,051± 6,278,287±175,359 (south) Marginal H20 64,262±18,564 44,124±3,048 12,426,359±1,667,189 (north) Surface H20 5,362±2,473 3,392260± 91,824,283±4,669,384 (south) Bottom H20 2,675±759 2,166±136 136,198,484±4,573,772 (south) Surface H20 3,635±1,214 888±124 164,486,652±2,678,424 (north) Bottom H20 3,336±989 1,529±135 138,313,306±9,632,133 (north)
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Table D.6 Chichester Reservoir wet sampling period. Total cells refer to qPCR results determined using DNA extracted from samples using the vigorous PowerSoil lysis technique. Vegetative cells refers to qPCR results for DNA obtained using the enzymatic- chemical lysis approach. Values represent the mean ± SE (n=6). Streptomyces-16SrRNA Actinobacteria- 16SrRNA Sample ID Total cells Vegetative Cells Total cells Soil (south) 306,806±19,746 118,805±24,902 28,272,124±3,696,912 Sediment (south) 131,293±8,372 37,725±3,748 28,689,914±3,486,239 Soil (north) 39,285±1,849 30,568±5,039 3,262,975±554,248 Sediment (north) 18,965±4,952 2,520±1,077 10,876,578±1,501,458 Debris (south) 179,035±29,829 71,771±11,676 29,499,804±3,426,822 Debris (north) 14,535±2,926 3,458±222 9,325,800±607,934 Macrophyte 1 (south) 14,525±862 9,671±574 126,742,852±3,855,381 Macrophyte 2 (south) 10,033±994 1,344±85 28,251,422±2,721,482 Algae (south) 36,051±7,167 2,565±361 138,864±6,620 Bottom sediment 84,744±5,791 22,638±1,479 3,408,755±89,834 (south) Bottom sediment 75,635±4,551 6,552±480 3,521,496±51,317 (north) Marginal H20 (south) 291,160±67,348 291,711±22,112 4,607,425±114,083 Marginal H20 (north) 270,921±62,889 191,206±19,862 9,266,768±66,996 Surface H20 (south) 105,394±18,280 92,394±4,687 45,909,894±616,914 Bottom H20 (south) 63,753±11,182 76,434±10,969 70,050,212±8,845,741 Surface H20 (north) 273,534±88,219 98,221±3,623 43,495,219±3,557,871 Bottom H20 (north) 68,757±5,355 76,434±10,969 110,415,384±8,126,061
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Actinobacteria abundance in environmental samples
Primers targeting taxonomically broader 16S rRNA sequences unique to the class actinobacteria were also applied in qPCR reactions using environmental DNA (obtained using the complete PowerSoil® DNA isolation kit procedure only) from the study presented in Chapter 4. The details of the primer pair are provided in Table D.7. The cycling conditions of the qPCR reactions were identical to the Streptomyces-specific primers as described in Chapter 4 (section 4.2.3) with the exception of a lower annealing temperature (57°C) and longer extension time (28s) due the longer amplicon being generated (274 bp). The specificity of the qPCR assay for detecting the target gene, was verified by conventional PCR using DNA isolated from S. coelicolor A3(2) (ATCC-471) and S. antibioticus (STR0). Figure D.1 shows the amplification products separated by gel electrophoresis. DNA isolated from cultures of S. coelicolor A3(2) and S. antibioticus
(STRO) were used as standards for the qPCR assay to establish calibration curves as described in section Chapter 4 (section 4.2.3) for the Streptomyces-specific primer pair.
The calibration curves are shown in Figure D.2.
Table D.7. Sequences of primer pair used in qPCR protocol for detection and quantification of actinobacteria-specific 16S rRNA. The amplified fragment size (base pairs, bp) is indicated. Primer ID Primer sequences (5’ → 3’) Amplicon Target gene Reference size Com2XF AAACTCAAAGGAATTGACGG ≈274 Actinobacteria Schafer et Ac1186R CTTCCTCCGAGTTGACCC 16S rRNA al. (2010)
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Figure D.1 Agarose gel electrophoresis of PCR products obtained by amplification from DNA isolated from two Streptomyces spp. using the two primer pairs targeting actinobacteria-specific 16S rRNA. Lane 1 contains S. antibioticus (STR0) DNA, lane 2 contains S. coelicolor A3(2) DNA and lane 3 is a negative control amplified with the actinobacteria -specific (270 bp) 16S rRNA targeting primer pair. 40 Streptomyces isolate S. coelicolor A3(2)
35 y = -3.766x + 42.809 y = -3.674x + 42.001
) R² = 0.998 R² = 0.995 T
30
25
20
Cycle theshold value (C value theshold Cycle 15
10 1 2 3 4 5 6 7 8 Log number of spores (cells) 10 Figure D.2 Standard curve for the qPCR assay using actinobacteria-specific 16S rRNA targeting primers. The linear relationship between the threshold cycle (CT) and the number of Streptomyces spores (cells) was obtained by serially diluting spore DNA extracts and performing qPCR. The standard error based on three replicates is indicated. 321
Comparisons of the Streptomyces and actinobacteria qPCR results (shown in Figure D.3) revealed that Streptomyces in some samples comprised a significant portion of the actinobacterial population (up to 13%), whilst representing a minority genus in the majority of samples (<0.1% of total abundance). Streptomyces comprised a greater proportion of the total actinobacterial population in Chichester marginal samples during the dry period. The proportion of Streptomyces in bottom sediment samples remained relatively constant in both reservoirs. During wet conditions when the density of
Streptomyces was higher in water samples, their numbers relative to the total population of actinobacteria was correspondingly greater.
322
Figure D.3 The proportion of Streptomyces comprising to the total actinobacterial abundance based on a comparison of qPCR data using primers targeting Streptomyces and actinobacteria 16S rRNA sequences respectively in Chichester (top) and Grahamstown (bottom) Reservoir samples for both sampling events.
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APPENDIX E: Hunter Water Corporation data for sampling dates
Table E.1 selected parameters measured by HWC in Grahamstown Reservoir for the sampling periods. Values represent the mean ± SD (average values for the month before sampling event) unless otherwise indicated in italics. Site R6 Wet conditions Dry conditions Water level (m AHD) on date of sampling 156.21 154.13 Air temp (°C) 21.1±2.2 17.3±4.5 Water temperature (°C) surface 23.82±1.2 20.12±1.50 Water temperature (°C) bottom 23.92±1.13 20.12±1.13 Rainfall (mm) monthly total 307 34 pH 7.3±0.4 7.4±0.04 Chlorophyll ɑ (µg/L) 5.4±1.6 4.3±1.2 Pheophytin (µg/L) 2.3±1.4 1.5±1.5 Dissolved Oxygen (mg/L) surface 9.8±0.40 11±0.44 Dissolved Oxygen (mg/L) bottom 9.5±0.33 10.8±0.38 Electrical Conductivity (µS/cm) 215.8±2.3 221±2.9 Suspended solids (mg/L) 3.7±1.5 3.5±0.7 E coli (cfu/100mL) 3.33±1.1 2±0 Total coliform (cfu/100mL) 6379±4665.5 144±90.5 Turbidity (NTU) 4.9±2.1 6.3±5 Silica (mg/L) 0.34±0.21 0.18±0.01 Total phosphorous (mg/L) 0.02±0.002 0.01±0.007 Total nitrogen (mg/L) 0.56±0.27 0.25±0 Total organic carbon (mg/L) 7.9±0.05 7.5±0.3 Phytoplankton (cells/100mL) 600,165±539,440 88,719±50,910 Cyanobacteria (cells/100mL) 592,439±542,472 84,204±50,613 Anabaena (cells/100mL) 450±0 20±0 Geosmin (ng/L) raw water intake 2.79±2.27 2.23±0.79 2-MIB (ng/L) raw water intake 0.91±0.16 1.12±0.23
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Table E.2 selected parameters measured by HWC in Chichester Reservoir for the sampling periods. Values represent the mean ± SD (average values for the month before sampling event) unless otherwise indicated in italics. Site R6 Wet conditions Dry conditions Water level (m AHD) on date of sampling 156.21 154.13 Water temperature (°C) surface 19.5±0.5 15.7±0.9 Water temperature (°C) bottom 19.4±0.2 15.7±1.0 Rainfall (mm) monthly total 295 13 pH 6.90±0.01 6.94±0.53 Chlorophyll_a (µg/L) 5.80±6.69 6.19±1.44 Pheophytin (µg/L) -2.00±6.43 0.36±1.06 Dissolved Oxygen (mg/L) surface 7.8±0.2 8.9±0.2 Dissolved Oxygen (mg/L) bottom 6.7±1.1 9.1±1.1 Electrical Conductivity (µS/cm) 70.00±0.63 72.67±2.62 Suspended solids (mg/L) 4.8±1.94 1.50±1.1 E coli (cfu/100mL) 6.4±4.1 0±0 Total coliform (cfu/100mL) 1494.20±1057.80 194.00±94.75 Heterotrophic plate count (cfu/mL) 506.00±299.70 59.33±10.21 Turbidity (NTU) 8.19±2.50 1.71±0.21 Silica (mg/L) 15.24±0.45 13.43±0.53 Total phosphorous (mg/L) 0.03±0.00 0.02±0.00 Total nitrogen (mg/L) 0.26±0.19 1.14±0.84 Total organic carbon (mg/L) 3.00±0.25 2.10±0.08 Phytoplankton (cells/100mL) 32,275±44,459 16,023±6,004 Cyanobacteria (cells/100mL) 30,910±44,974 7,673±5,029 Anabaena (cells/100mL) ND ND Geosmin (ng/L) raw water intake 0.3±0.1 1.4±0.3 2-MIB (ng/L) raw water intake 0.1±0.1 0.1±0.2
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APPENDIX F: Additional results for Chapter 6 (multifactorial experiments)
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Experimental matrices and results for Multifactorial Experiments (Chapter 6)
Experiment 1: The effects of carbon, nitrogen and phosphorous concentration on geosmin and 2-MIB production by Streptomyces spp.
Table F.1 Experimental matrix and results for assessing the influence of factors (carbon, phosphorous and nitrogen concentration) on the production of geosmin, 2-MIB and biomass by S. coelicolor A3(2). The differentiation value for each of the nine treatments is also indicated. Values represent the mean±SD. Experimental Factors Responses Experiment ID Carbon Nitrogen Phosphorous 2-MIB Geosmin Biomass Differentiation (mM) (µM) (µM) (ng/mg) (ng/mg) (mg) value (0-5) 1.1 416.6 3.6 1.6 2.07±0.76cd 5.72±0.79d 4.68±0.20bc 3 1.2 416.6 1,784.8 1.6 1.11±0.47cd 5.69±0.41d 5.56±0.76bc 3 1.3 416.6 3.6 807.2 1.00±0.19cd 1.80±0.38e 6.24±0.33ab 2 1.4 416.6 1,784.8 807.2 0.60±0.34d 1.29±0.32e 7.71±0.76a 0 1.5 208.5 894.2 404.4 1.80±0.74cd 2.03±0.33e 4.68±0.69c 0 1.6 0.24 3.6 1.6 11.13±2.5a 21.80±2.23a 0.96±0.05d 5 1.7 0.24 1,784.8 1.6 5.82±2.44b 12.46±0.35b 1.76±0.22d 4 1.8 0.24 3.6 807.2 4.00±0.95bc 10.58±1.70bc 1.52±0.59d 4 1.9 0.24 1,784.8 807.2 4.26±0.87bc 9.10±2.65c 2.01±0.69d 4 Values that do not contain the same letter superscript are significantly different (p<0.05).
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Table F.2 Experimental matrix and results for assessing the influence of factors (carbon, phosphorous and nitrogen concentration) on the production of geosmin, 2-MIB and biomass by S. antibioticus. The differentiation value for each of the nine treatments is also indicated. Values represent the mean±SD. Experimental Factors Responses Experiment Carbon Nitrogen Phosphorous 2-MIB Geosmin Biomass (mg) Differentiation ID (mM) (µM) (µM) (ng/mg) (ng/mg) value (0-5) 1.1 416.6 3.6 1.6 16.45±2.32a 11.31±1.26a 4.82±0.27bc 5 1.2 416.6 1,784.8 1.6 2.94±0.31cd 3.86±0.35b 5.21±0.26b 3 1.3 416.6 3.6 807.2 4.13±1.31c 4.44±0.71b 5.70±0.46b 4 1.4 416.6 1,784.8 807.2 0.38±0.20d 0.56±0.10c 7.41±0.30a 0 1.5 208.5 894.2 404.4 0.84±0.28d 1.25±0.42c 4.00±0.55c 0 1.6 0.24 3.6 1.6 11.60±1.03b 13.15±1.90a 0.74±0.09e 5 1.7 0.24 1,784.8 1.6 3.05±0.72c 3.30±1.65bc 1.83±0.26d 3 1.8 0.24 3.6 807.2 2.51±0.82cd 4.12±1.36b 1.52±0.17de 4 1.9 0.24 1,784.8 807.2 2.78±0.79cd 3.94±0.80b 1.61±0.03de 3 Values that do not contain the same letter superscript are significantly different (p<0.05).
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Experiment 2: The effects of temperature, pH and NaCl concentration on geosmin and 2-MIB production by Streptomyces spp.
Table F.3 Experimental matrix and results for assessing the influence of factors (temperature, NaCl concentration and pH) on the production of geosmin, 2-MIB and biomass by S. coelicolor A3(2). The differentiation value for each of the nine treatments is also indicated. Values represent the mean±SD.
Experimental Factors Responses
Experiment Temperature NaCl pH 2-MIB Geosmin Biomass Differentiation ID (°C) (%) (ng/mg) (ng/mg) (mg) value (0-5) 2.1 30 0 6 1.61±0.58c 11.17±1.60c 2.24±0.48d 5 2.2 30 2 6 0.25±0.07c 0.90±0.42c 7.96±0.60a 1 2.3 30 0 9 4.71±1.86b 28.34±8.41b 2.44±0.13d 5 2.4 30 2 9 0.42±0.14c 0.85±0.28c 6.88±0.10ab 0 2.5 27 1 7.5 1.45±0.85c 3.81±0.68c 3.40±0.41c 3 2.6 24 0 6 7.74±2.46a 23.22±1.74b 1.34±0.10d 5 2.7 24 2 6 0.39±0.23c 0.78±0.18c 7.79±0.73a 0 2.8 24 0 9 10.40±0.84a 40.66±11.50a 1.43±0.34d 5 2.9 24 2 9 0.42±0.08c 1.44±0.26c 6.48±0.09b 0 Values that do not contain the same letter superscript are significantly different (p<0.05).
329
Table F.4 Experimental matrix and results for assessing the influence of factors (temperature, NaCl concentration and pH) on the production of geosmin, 2-MIB and biomass by S. antibioticus. The differentiation value for each of the nine treatments is also indicated. Values represent the mean±SD. Experimental Factors Responses Experiment Temperature NaCl pH 2-MIB Geosmin Biomass Differentiation ID (°C) (%) (ng/mg) (ng/mg) (mg) value (0-5) 2.1 30 0 6 9.30±3.43b 21.61±2.68d 1.56±0.46d 5 2.2 30 2 6 0.37±0.14c 2.37±0.61d 7.47±0.35a 2 2.3 30 0 9 38.71±4.08a 57.70±1.96c 0.85±0.08d 5 2.4 30 2 9 0.78±0.31c 3.70±1.06d 6.39±0.51a 2 2.5 27 1 7.5 11.14±2.56b 17.84±4.00d 3.28±0.65c 4 2.6 24 0 6 13.34±2.27b 91.29±24.05b 0.66±0.19d 5 2.7 24 2 6 1.43±0.61c 6.76±2.24d 6.25±0.64ab 4 2.8 24 0 9 36.34±2.48a 192.53±17.99a 0.68±0.05d 5 2.9 24 2 9 0.77±0.03c 7.43±1.01d 5.02±0.05b 2 Values that do not contain the same letter superscript are significantly different (p<0.05).
330
Experiment 3: The effects of calcium, magnesium and potassium concentration on geosmin and 2-MIB production by Streptomyces spp.
Table F.5 Experimental matrix and results for assessing the influence of factors (calcium, potassium and magnesium concentration) on the production of geosmin, 2-MIB and biomass by S. coelicolor A3(2). The differentiation value for each of the nine treatments is also indicated. Values represent the mean±SD. Experimental Factors Responses Experiment Calcium Magnesium Potassium 2-MIB Geosmin (ng/mg) Biomass Differentiation ID (µM) (µM) (µM) (ng/mg) (mg) value (0-5) 33 3.1 1000 1 1 33.42±3.87bc 98.46±13.69bc 1.21±0.11ab 5 37 3.2 1000 1000 1 30.29±5.28bc 65.24±10.11c 1.16±0.16ab 5 34 3.3 1000 1 1000 58.75±8.20a 180.92±25.54a 0.94±0.14b 5 38 3.4 1000 1000 1000 33.12±2.72bc 110.69±13.90bc 1.15±0.00ab 5 30 3.5 500.1 500.1 500.1 39.10±6.66b 115.85±19.38b 1.07±0.12ab 5 31 3.6 0.25 1 1 20.60±0.64c 83.62±1.08bc 1.15±0.00ab 5 35 3.7 0.25 1000 1 25.04±6.05c 80.56±17.43bc 1.15±0.00ab 5 32 3.8 0.25 1 1000 23.83±0.80c 94.33±16.46bc 1.28±0.11a 5 36 3.9 0.25 1000 1000 23.45±0.99c 90.78±7.30bc 1.21±0.11ab 5 Values that do not contain the same letter superscript are significantly different (p<0.05).
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Table F.6 Experimental matrix and results for assessing the influence of factors (calcium, potassium and magnesium concentration) on the production of geosmin, 2-MIB and biomass by S. antibioticus. The differentiation value for each of the nine treatments is also indicated. Values represent the mean±SD. Experimental Factors Responses Experiment Calcium Magnesium Potassium 2-MIB Geosmin Biomass Differentiation ID (µM) (µM) (µM) (ng/mg) (ng/mg) (mg) value (0-5) 33 3.1 1000 1 1 11.44±3.03bc 59.60±3.55b 1.04±0.04ab 5 37 3.2 1000 1000 1 16.03±5.24ab 61.34±4.05b 1.22±0.06ab 5 34 3.3 1000 1 1000 25.12±8.51a 111.46±15.16a 0.64±0.00c 5 38 3.4 1000 1000 1000 13.07±2.63ab 66.29±3.21b 1.30±0.04a 5 30 3.5 500.1 500.1 500.1 14.23±6.19ab 66.29±24.25b 1.12±0.21ab 5 31 3.6 0.25 1 1 9.30±1.15bc 48.76±6.44b 1.01±0.13ab 5 35 3.7 0.25 1000 1 9.43±2.70bc 41.57±2.43b 1.27±0.06ab 5 32 3.8 0.25 1 1000 10.84±1.28bc 51.76±4.89b 0.94±0.07bc 5 36 3.9 0.25 1000 1000 12.57±2.27ab 49.38±1.44b 1.11±0.04ab 5 Values that do not contain the same letter superscript are significantly different (p<0.05).
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Experiment 4: The effects of iron, zinc, copper and manganese concentration on geosmin and 2-MIB production by Streptomyces spp. Table F.7 Experimental matrix and results for assessing the influence of micronutrient factors (iron, zinc, copper and manganese concentration) on the production of geosmin, 2-MIB and biomass by S. coelicolor A3(2). The differentiation value for each of the 17 treatments is also indicated. Values represent the mean±SD. Experimental Factors Responses Experiment Iron Zinc Copper Manganese 2-MIB Geosmin Biomass Differentiation ID (µM) (µM) (µM) (µM) (ng/mg) (ng/mg) (mg) value (0-5) 4.1 25 25 25 25 6.90±1.44ef 7.70±3.81f 1.20±0.00abc 0 4.2 25 25 25 0.25 11.08±1.94def 7.92±2.80f 1.32±0.12ab 0 4.3 25 25 0.25 25 13.91±1.52cdef 54.32±5.00de 1.25±0.09ab 5 4.4 25 25 0.25 0.25 22.32±11.32bcd 77.57±13.30cde 1.08±0.08bc 4 4.5 25 0.25 25 25 5.91±0.99ef 10.03±1.89f 1.37±0.04ab 0 4.6 25 0.25 25 0.25 5.16±1.78ef 7.38±0.86f 1.61±0.07a 0 4.7 25 0.25 0.25 25 22.64±10.29bcd 80.79±11.18bcd 1.28±0.00ab 4 4.8 25 0.25 0.25 0.25 43.98±11.05a 135.26±36.67a 0.98±0.25bc 5 4.9 125.1 125.1 125.1 125.1 8.07±1.91ef 45.75±10.19e 1.23±0.32ab 3 4.10 0.25 25 25 25 4.03±1.23f 9.39±1.21f 1.28±0.00ab 0 4.11 0.25 25 25 0.25 4.39±1.10f 11.66±1.18f 1.10±0.00bc 0 4.12 0.25 25 0.25 25 18.92±0.61bcde 81.77±10.85bcd 1.43±0.11ab 4 4.13 0.25 25 0.25 0.25 14.23±0.89cdef 66.95±7.17cde 1.34±0.03ab 4 4.14 0.25 0.25 25 25 9.93±3.36def 13.33±3.67f 0.75±0.00c 0 4.15 0.25 0.25 25 0.25 4.01±0.78f 9.17±3.09f 1.11±0.20bc 0 4.16 0.25 0.25 0.25 25 26.82±1.00bc 97.94±1.46bc 1.25±0.22ab 3 4.17 0.25 0.25 0.25 0.25 29.26±4.14b 114.90±24.91ab 1.08±0.07bc 5 Values that do not contain the same letter superscript are significantly different (p<0.05).
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Table F.8 Experimental matrix and results for assessing the influence of micronutrient factors (iron, zinc, copper and manganese concentration) on the production of geosmin, 2-MIB and biomass by S. antibioticus. The differentiation value for each of the 17 treatments is also indicated. Values represent the mean±SD. Experimental Factors Responses Experiment Iron Zinc Copper Manganese 2-MIB Geosmin Biomass (mg) Differentiation ID (µM) (µM) (µM) (µM) (ng/mg) (ng/mg) value (0-5) 4.1 25 25 25 25 4.40±1.40b 7.65±2.33d 1.39±0.23a 1 4.2 25 25 25 0.25 3.63±1.85ab 7.17±1.40d 1.60±0.12a 1 4.3 25 25 0.25 25 5.35±1.12ab 21.18±7.61bcd 1.50±0.00abc 4 4.4 25 25 0.25 0.25 8.90±5.77a 18.36±6.14cd 1.03±0.11ef 5 4.5 25 0.25 25 25 3.32±0.76ab 8.35±2.80d 1.63±0.05a 4 4.6 25 0.25 25 0.25 3.89±0.77ab 11.43±0.64cd 1.61±0.09a 3 4.7 25 0.25 0.25 25 8.22±1.42ab 34.95±14.55ab 0.99±0.20ef 5 4.8 25 0.25 0.25 0.25 8.94±2.00a 37.53±1.78a 0.59±0.10g 5 4.9 125.1 125.1 125.1 125.1 5.20±1.65ab 22.63±3.22bc 1.16±0.03def 4 4.10 0.25 25 25 25 2.06±1.09ab 5.75±1.70d 1.60±0.01abcd 1 4.11 0.25 25 25 0.25 4.79±1.93ab 7.03±1.46d 1.54±0.10ab 1 4.12 0.25 25 0.25 25 7.38±2.59ab 26.72±7.97abc 0.95±0.09f 5 4.13 0.25 25 0.25 0.25 5.37±2.17ab 14.39±3.08cd 1.26±0.02bcde 5 4.14 0.25 0.25 25 25 3.80±1.20ab 16.89±0.89cd 1.10±0.08def 2 4.15 0.25 0.25 25 0.25 6.09±1.66ab 12.59±1.48cd 1.26±0.00bcdef 2 4.16 0.25 0.25 0.25 25 6.15±0.65ab 27.05±7.64abc 1.20±0.18cdef 5 4.17 0.25 0.25 0.25 0.25 7.25±2.87ab 26.62±3.50abc 1.23±0.06cdef 5 Values that do not contain the same letter superscript are significantly different (p<0.05
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Validity of the MLR models for Chapter 6
The MLR models for each of the experiments were evaluated for their validity (regression model significance and fitting power) with analysis of variance (ANOVA). This is an evaluation of how well the MLR model fits the experimental data. In analysis of variance
(ANOVA) the total variation or the sum of squares (SS) of the response is partitioned into two components: one part due to the regression model (SS regression) and a part due to the residuals (SS residuals), the variation that cannot be modelled. The total variance (SS) of the response corrected for the mean can be expressed as:
SS = SS regression + SS residuals
In the regression model significance test, a good model results if the SS regression is high and if the SS residual is low. That is, the model fits the experimental data well. The sizes of these two variances are compared by an F-test (ratio of MS regression divided by MS residuals). MS is obtained by dividing the SS by the respective degree of freedom (DF). If the probability value
(p) <0.05, the variance explained by the model (MS regression) is significantly larger than the unexplained variance (MS residuals) and the regression model can be considered significant.
Thus MLR is based on finding the regression model which minimises the residual sum of squares (SS residuals) of the response variables (i.e. the amount of variance in the response that cannot be explained by the model). Given that there was replication in the experiments, the residuals component (SS residuals) was further divided into sum of square lack of fit (SS lof) and sum of squares pure experimental error (SS pe). Ideally both values are small and of similar size. The model error is tested with an F-test (ratio of MS model error divided by MS replicate error). If the resulting p>0.05, the model has small error and good fitting power and the model equation is said to not present a lack of fit. 335
To evaluate the fit of the MLR models for each of the experiments, values of explained variation R2 and predicted variation Q2 provide excellent guidance. R2 is the fraction of the total variation in the response explained in the model corrected for the mean. It is a measure of how well the raw data fits the model and consists of two parts; one from the regression model (SS regression) and another due to the residual (SS residuals). It is calculated as follows:
2 R = (SS-SS residual)/SS
Small residuals will render a higher degree of explained variation (R2). This is the classical quantity used to evaluate goodness of fit. The predicted variation Q2 is the fraction of the total variation of the response that can be predicted in the model and is calculated as:
Q2 = (SS – PRESS)/SS
PRESS is the prediction residual sum of squares and is determined through cross-validation.
Small deviations between the actual residuals and predicted residuals render low PRESS values and high values of predicted variation (Q2). Thus Q2 estimates the predictive power of the MLR model.
In addition to examining the fit quality, significance and precision of a MLR model, ANOVA also allows determination of the effects of variables and interactions between variables. All variables and interactions between them in the MLR model are characterised by a regression coefficient and corresponding p-value determined through ANOVA which indicates the significance of their influence on the response. The scaled and centred coefficients of the model determined through ANOVA indicate the extent to which experimental variables and interactions between them influence the response variable.
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Experiment 1
When evaluating the validity of the models for geosmin, 2-MIB and biomass production by
S. coelicolor A3(2) and S. antibioticus with ANOVA (Tables F.9 and F.10 respectively), results showed that the F-values exceeded the F- critical values and p<0.05, indicating the regression models were statistically significant at the 95% confidence level in the range studied. The lack of fit however was significant (p<0.05) for all models relative to the pure error indicating a lack of fitting power as the SD LOF was of similar value to the SD pe multiplied by the square root of critical F (12.111). It should be noted here that extremely high reproducibility (close to 1.0) can lead to negative values for model validity. This occurs when the replicated error (pure error) is very small and not representative compared to the total variance of the response. However, this is not a real lack of fit, but an artefact due to the extremely low standard deviation of the replicated points. This behaviour was observed for the majority of models presented. For S .coelicolor A3(2), the coefficient of determination
(R2) for geosmin was 0.79, meaning that 79% of the variation in geosmin can be explained by the fitted model and the response variation percentage predicted by the MLR model according to cross validation (Q2) was 0.65. In the case of 2-MIB, R2 and Q2 were 0.73 and
0.62 and for biomass, 0.91 and 0.87 respectively. Significant models were also developed for
S. antibioticus with a R2 and Q2 values of 0.70 and 0.57, 0.80 and 0.72 and 0.96 and 0.93 respectively for geosmin, 2-MIB and biomass. Values of R2 and Q2 acceptable for biological data are reported to be >0.7 and >0.4 respectively, indicating that the estimated models for each response fitted the experimental data adequately. Furthermore an appropriate model is evident if the differences between these two values does not exceed 0.2-0.3 which was apparent in the above models (Lundstedt et al., 1998).
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Table F.9 ANOVA table for the MLR models developed for geosmin, 2-MIB and biomass production by S. coelicolor A3(2) in Experiment 1.
Response Source of variation DF SS MS (variance) Fvalue Fcritical p SD Geosmin Total Corrected 32 318.03 9.94 3.15 Regression 6 249.58 41.60 15.80 2.47 0.000 6.45 Residual 26 68.45 2.63 1.62 Lack of fit (model 2 34.38 17.19 12.11 3.40 0.000 4.15 error) Pure error (replicate 24 34.07 1.42 1.19 error) 2-MIB Total Corrected 32 1240.60 38.77 6.23 Regression 6 902.84 150.47 11.58 2.47 0.000 12.27 Residual 26 337.76 12.99 3.60 Lack of fit (model 2 304.76 152.38 110.82 3.40 0.000 12.34 error) Pure error (replicate 24 33.00 1.38 1.17 error) Biomass Total Corrected 32 150.25 4.70 2.17 Regression 6 136.75 22.80 43.90 2.47 0.000 4.77 Residual 26 13.50 0.52 0.72 Lack of fit (model 2 5.27 2.637 7.68 3.40 0.003 1.62 error) Pure error (replicate 24 8.237 0.347 0.59 error) DF: degrees of freedom, SS: sum of squares, MS: mean square, SD: standard deviation.
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Table F.10 ANOVA data for the MLR models developed for geosmin, 2-MIB and biomass production by S. antibioticus in Experiment 1.
Response Source of variation DF SS MS F-test Fcritical p SD (variance) Geosmin Total Corrected 32 535.69 16.74 4.091 Regression 6 375.01 62.50 10.11 2.47 0.000 7.91 Residual 26 160.68 6.18 2.49 Lack of fit (model 2 137.18 68.59 70.04 3.40 0.000 8.289 error) Pure error (replicate 24 23.50 0.98 0.99 error) 2-MIB Total Corrected 32 803.77 25.12 5.01 Regression 6 641.70 106.95 17.16 2.47 0.000 10.34 Residual 26 162.07 6.23 2.50 Lack of fit (model 2 141.16 70.58 81.01 3.40 0.000 8.40 error) Pure error (replicate 24 20.91 0.87 0.93 error) Biomass Total Corrected 32 131.50 4.11 2.03 Regression 6 125.97 21.00 98.65 2.47 0.000 4.58 Residual 26 5.53 0.21 0.46 Lack of fit (model 2 2.64 1.32 10.98 3.40 0.000 1.15 error) Pure error (replicate 24 2.89 0.12 0.35 error) DF: degrees of freedom, SS: sum of squares, MS: mean square, SD: standard deviation.
Experiment 2
The ANOVA results indicating the validity of the MLR models developed for geosmin, 2-
MIB and biomass production by S. coelicolor A3(2) and S. antibioticus (Tables F.11 and
F.12 respectively) revealed the regression models were statistically significant at the 95% confidence level, with the F-values exceeding the F- critical values for all models. While the lack of fit was significant (p<0.05) for the models, as indicated previously, this can be attributed to the low replicate error (i.e. very small SD of replicates). The exceptions were the MLR models developed for biomass and 2-MIB production by S. antibioticus, where the lack of fit was not significant (p>0.05) in which case larger replicate errors (SD of replicates) 339
are evident. For S. coelicolor A3(2), the relationships between the treatment factors and
response variables using MLR provided significant models as indicated by the R2 and Q2
values (0.83 and 0.73, 0.88 and 0.81 and 0.93 and 0.89 respectively for geosmin, 2-MIB and
biomass). Satisfactory results indicating a high goodness of fit and goodness of prediction
were also obtained for S. antibioticus, with R2 and Q2 values of 0.90 and 0.85, 0.96 and 0.94
and 0.97 and 0.95 for geosmin, 2-MIB and biomass response variables respectively.
Table F.11 ANOVA results for the MLR models developed for geosmin, 2-MIB and biomass production by S. coelicolor A3(2) in Experiment 2.
Response Source of variation DF SS MS Fvalue Fcritical p SD (variance) Geosmin Total Corrected 32 6113.94 191.06 13.82 Regression 6 5050.28 841.71 20.57 2.47 0.000 29.01 Residual 26 1063.67 40.91 6.40 Lack of fit (model 2 637.32 318.66 17.94 3.40 0.000 17.85 error) Pure error (replicate 24 426.35 17.76 4.22 error) 2-MIB Total Corrected 32 374.14 11.69 3.42 Regression 6 328.07 54.68 30.86 2.47 0.000 7.39 Residual 26 46.07 1.77 1.33 Lack of fit (model 2 16.28 8.14 6.56 3.40 0.005 2.86 error) Pure error (replicate 24 29.79 1.24 1.11 error) Biomass Total Corrected 32 196.00 6.13 2.48 Regression 6 181.42 30.24 53.88 2.47 0.000 5.50 Residual 26 14.59 0.56 0.75 Lack of fit (model 2 10.88 5.43 35.02 3.40 0.000 2.33 error) Pure error (replicate 24 3.72 0.16 0.39 error) DF: degrees of freedom, SS: sum of squares, MS: mean square, SD: standard deviation.
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Table F.12 ANOVA results for the MLR models developed for geosmin, 2-MIB and biomass production by S. antibioticus Experiment 2.
Response Source of variation DF SS MS Fvalue Fcritical p SD (variance) Geosmin Total Corrected 32 101755 3179.86 56.39 Regression 6 91214.2 15202.4 37.50 2.47 0.000 123.30 Residual 26 10541.2 405.43 20.14 Lack of fit (model 2 8325.57 4162.79 45.10 3.40 0.000 64.52 error) Pure error (replicate 24 2215.63 92.32 9.61 error) 2-MIB Total Corrected 32 5649.5 176.55 13.29 Regression 6 5421.22 903.54 102.91 2.47 0.000 30.06 Residual 26 228.28 8.78 2.96 Lack of fit (model 2 47.28 23.64 3.13 3.40 0.062 4.86 error) Pure error (replicate 24 181.00 7.54 2.75 error) Biomass Total Corrected 32 187.96 5.87 2.42 Regression 6 181.90 30.32 130.03 2.47 0.000 5.51 Residual 26 6.06 0.23 0.48 Lack of fit (model 2 0.99 0.50 2.35 3.40 0.117 0.70 error) Pure error (replicate 24 5.07 0.21 0.46 error) DF: degrees of freedom, SS: sum of squares, MS: mean square, SD: standard deviation.
Experiment 3
Tables F.13 and F.14 show the tabulated output obtained by performing ANOVAs for all response variables for S. coelicolor A3(2) and S. antibioticus respectively. The statistics indicated that the fit of the regression models were significant (F-value >F-critical and p<0.05) with the exception of biomass for S. coelicolor A3(2) in which case the MLR model developed did not fit the experimental data adequately (p>0.05). While the lack of fit was significant (p<0.05) for the models, as indicated previously, this can be attributed to the low replicate error (i.e. very small SD of replicates). The exceptions were the MLR models developed for geosmin and 2-MIB production by S. antibioticus, where the lack of fit was 341
not significant (p>0.05) in which case larger replicate errors (SD of replicates) are evident.
For S. coelicolor A3(2), the MLR models developed for geosmin and 2-MIB had R2 and Q2 values of 0.72 and 0.59 and 0.76 and 0.63 respectively, indicating the models fitted the experimental data adequately. The low values obtained for biomass (R2 = 0.316, Q2 = -0.086) indicated poor goodness of fit and predictability of the model developed for this response.
For S. antibioticus, the MLR developed for 2-MIB also lacked fit and predictive power (R2
= 0.40, Q2 = 0.06), whereas explained variation and predicted variation were satisfactory for geosmin (R2 = 0.60, Q2 = 0.47) and biomass (R2 = 0.63, Q2 = 0.49).
Table F.13 ANOVA results for the MLR models developed for geosmin, 2-MIB and biomass production by S. coelicolor A3(2) in Experiment 3.
Response Source of variation DF SS MS F-test Fcritical p SD (variance) Geosmin Total Corrected 32 33584.2 1049.51 32.40 Regression 6 25401 4233.49 13.45 2.47 0.000 65.07 Residual 26 8183.22 314.74 17.74 Lack of fit (model 2 2026.73 1013.36 3.95 3.40 0.033 31.83 error) Pure error (replicate 24 6156.49 256.52 16.02 error) 2-MIB Total Corrected 32 4145.16 129.54 11.38 Regression 6 2981.40 496.90 11.10 2.47 0.000 22.29 Residual 26 1163.76 44.76 6.69 Lack of fit (model 2 540.81 270.41 10.42 3.40 0.001 16.44 error) Pure error (replicate 24 622.94 25.96 5.09 error) Biomass Total Corrected 32 0.518 0.02 0.13 Regression 6 0.164 0.03 1.20 2.47 0.102 0.17 Residual 26 0.35 0.01 0.12 Lack of fit (model 2 0.09 0.05 4.35 3.40 0.024 0.22 error) Pure error (replicate 24 0.26 0.01 0.10 error) DF: degrees of freedom, SS: sum of squares, MS: mean square, SD: standard deviation.
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Table F.14 ANOVA results for the MLR models developed for geosmin, 2-MIB and biomass production by S. antibioticus in Experiment 3.
Response Source of variation DF SS MS F-test Fcritical p SD (variance) Geosmin Total Corrected 32 21509.4 672.17 25.93 Regression 6 12911.4 2151.89 6.51 2.47 0.000 46.39 Residual 26 8598.08 330.70 18.19 Lack of fit (model 2 1681.84 840.92 2.92 3.40 0.073 29.00 error) Pure error (replicate 24 6916.24 288.18 16.98 error) 2-MIB Total Corrected 32 1582.8 49.46 7.03 Regression 6 632.54 105.42 2.88 2.47 0.027 10.27 Residual 26 950.26 36.55 6.05 Lack of fit (model 2 185.28 92.64 2.91 3.40 0.074 9.63 error) Pure error (replicate 24 764.98 31.88 5.651 error) Biomass Total Corrected 32 1.34 0.04 0.21 Regression 6 0.84 0.14 7.40 2.47 0.000 0.38 Residual 26 0.50 0.02 0.14 Lack of fit (model 2 0.14 0.07 4.48 3.40 0.022 0.26 error) Pure error (replicate 24 0.36 0.02 0.12 error) DF: degrees of freedom, SS: sum of squares, MS: mean square, SD: standard deviation.
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Experiment 4
The ANOVA results presented in Tables F.15 and F.16 (S. coelicolor A3(2) and S. antibioticus respectively) indicate the validity of the regression models developed for each response variable, being significant at the 95% confidence level (F-value>F-critical and p<0.05). As indicated previously, the significant lack of fit (p<0.05) obtained for the majority of models can be attributed to the small replicate errors obtained. The goodness of fit (R2) and goodness of predictability (Q2) values obtained for the models developed for S. coelicolor A3(2) were satisfactory including respective values of 0.92 and 0.87 for geosmin,
0.78 and 0.66 for 2-MIB and 0.55 and 0.41 for biomass. R2 and Q2 values obtained for S. antibioticus models were acceptable for geosmin and biomass (0.76 and 0.62 and 0.55 and
0.41 respectively) but lower values obtained for 2-MIB (0.48 and 0.15) indicate poor fitting and predicting power of the MLR model.
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Table F.15 ANOVA results for the MLR models developed for geosmin, 2-MIB and biomass production by S. coelicolor A3(2) in Experiment 4.
Response Source of variation DF SS MS F-test Fcritical p SD (variance)
Geosmin Total Corrected 56 95327.4 1702.28 41.26 Regression 10 87546.2 8754.62 51.76 2.04 0.000 93.57 Residual 46 7781.17 169.16 13.01 Lack of fit (model error) 6 2231.56 371.93 2.68 2.34 0.028 19.29 Pure error (replicate 40 5549.6 138.74 11.78 error) 2-MIB Total Corrected 56 7064.72 126.16 11.23 Regression 10 5484.28 548.43 15.96 2.04 0.000 23.42 Residual 46 1580.44 34.36 5.86 Lack of fit (model error) 6 757.71 126.29 6.14 2.34 0.000 11.24 Pure error (replicate 40 822.73 20.57 4.54 error) Biomass Total Corrected 56 2.71 0.05 0.22 Regression 10 1.48 0.15 5.54 2.04 0.000 0.39 Residual 46 1.23 0.03 0.16 Lack of fit (model error) 6 0.32 0.05 2.31 2.34 0.052 0.23 Pure error (replicate 40 0.91 0.02 0.15 error)
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Table F.16 ANOVA results for the MLR models developed for geosmin, 2-MIB and biomass production by S. antibioticus in Experiment 4.
Response Source of variation DF SS MS F-test Fcritical p SD (variance)
Geosmin Total Corrected 56 5951.87 106.28 10.31 Regression 10 4505.42 450.54 14.33 2.04 0.0000 21.23 Residual 46 1446.46 31.44 5.61 Lack of fit (model error) 6 440.11 73.35 2.92 2.34 0.019 8.57 Pure error (replicate 40 1006.35 25.16 5.02 error) 2-MIB Total Corrected 56 363.64 6.49 2.55 Regression 10 172.94 17.29 4.17 2.04 0.0000 4.16 Residual 46 190.69 4.15 2.04 Lack of fit (model error) 6 24.42 4.07 0.98 2.34 0.452 2.02 Pure error (replicate 40 166.27 4.16 2.04 error) Biomass Total Corrected 56 4.51 0.08 0.28 Regression 10 3.21 0.32 11.37 2.04 0.000 0.57 Residual 46 1.30 0.03 0.17 Lack of fit (model error) 6 0.90 0.15 15.23 2.34 0.000 0.39 Pure error (replicate 40 0.40 0.01 0.10 error)
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Relationship between stage of morphological differentiation and T&O metabolite production in multifactorial experiments
Biomass from each of the cultures grown under the various treatments in the experiments were examined microscopically to assess the extent of differentiation. The scatter plots presented below (Figures F.1 to F.6) show the positive relationship that exists between the degree of morphological differentiation and the yields of geosmin and 2-MIB produced for each of the experiments conducted for both Streptomyces spp. In the case of Experiment 3, for all treatments, the cells were highly differentiated and heavily sporulating and were assigned with a differentiation value of 5. Hence no regression analyses of the relationship between the extent of differentiation and production of the secondary metabolites was conducted.
2-MIB Geosmin 25 y = 0.73e0.43x y = 1.297e0.5146x 20 R² = 0.66 R² = 0.892
15
10
Concentration(ng/mg) 5
0 0 1 2 3 4 5 Differentiation value Figure F.1 Relationship between geosmin and 2-MIB production by S. coelicolor A3(2) and differentiation value in Experiment 1 (effects of carbon, nitrogen and phosphorous concentration).
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18 2-MIB Geosmin 16 14 R² = 0.8789 R² = 0.9124 12 10 8 6
4 Concentration(ng/mg) 2 0 0 1 2 3 4 5 Differentiation value Figure F.2 Relationship between geosmin and 2-MIB production by S. antibioticus and differentiation value in Experiment 1 (effects of carbon, nitrogen and phosphorous concentration).
45 2-MIB Geosmin 40 R² = 0.9195 35 R² = 0.822 30 25 20 15 10
5 Concentration(ng/mg) 0 0 1 2 3 4 5 Differentiation value Figure F.3 Relationship between geosmin and 2-MIB production yields by S. coelicolor A3(2) and differentiation value in Experiment 2 (effects of temperature, pH and NaCl concentration).
348
250 2-MIB Geosmin 200 R² = 0.8279 R² = 0.7304
150
100
50 Concentration(ng/mg) 0 0 1 2 3 4 5 Differentiation value Figure F.4 Relationship between geosmin and 2-MIB production yields by S. antibioticus and differentiation value in Experiment 2 (effects of temperature, pH and NaCl concentration).
2-MIB Geosmin 160 140 R² = 0.715 R² = 0.920 120 100 80 60 40
Concentration(ng/mg) 20 0 0 1 2 3 4 5 Differentiation value
Figure F.5 Relationship between geosmin and 2-MIB production by S. coelicolor A3(2) and differentiation value in Experiment 4 (effects of micronutrient concentration).
349
40 35 2-MIB Geosmin 30 R² = 0.5084 R² = 0.6922 25 20 15 10
Concentration(ng/mg) 5 0 0 1 2 3 4 5 Differentiation value
Figure F.6 Relationship between geosmin and 2-MIB production by S. antibioticus and differentiation value in Experiment 4 (effects of micronutrient concentration).
Geladi, P., & Kowalski, B. R. (1986). Partial least-squares regression: a tutorial. Anlytica Chimica Acta, 185, 1-17. Singh, K. P., Malik, A., Basant, N., & Saxena, P. (2007). Multi-way partial least squares modeling of water quality data. Analytica Chimica Acta, 584(2), 385-396.
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