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UBC Theses and Dissertations

Assessment of microbial cell viability in municipal sludge after microwave and ultrasound and subsequent… Cella, Monica Angela 2015

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ASSESSMENT OF MICROBIAL CELL VIABILITY IN MUNICIPAL SLUDGE AFTER MICROWAVE AND ULTRASOUND AND SUBSEQUENT IMPACTS ON ANAEROBIC DIGESTION  by  Monica Angela Cella  B.Sc., University of British Columbia, 2012  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF APPLIED SCIENCE  in  THE COLLEGE OF GRADUATE STUDIES  (Civil Engineering)  THE UNIVERSITY OF BRITISH COLUMBIA  (Okanagan)  September 2015   © Monica Angela Cella, 2015  ii Abstract Anaerobic digestion (AD) is an effective method of waste sludge stabilization with lower costs and energy footprints; recently, researchers have begun to optimize the enhancement of AD by pretreating sludge prior to AD. In this study, two sludge pretreatment methods, ultrasonication (US) and microwave irradiation (MW) were compared to determine the relative amounts of microbial inactivation within pretreated secondary sludge, and how this cellular destruction translates to AD processes. The intensities chosen for MW-pretreatment were based on previous studies using sludge from the wastewater treatment plant in Kelowna, BC, Canada. The intensity range chosen for MW possessed specific energy inputs of 2.17, 2.62, 4.89, and 6.48 kJ/g total solids (TS). For comparison purposes, US intensities were calculated to be 2.37, 4.74, 6.73, 23.09, and 27.71 kJ/g TS. Using a novel approach, the extent of cellular destruction caused at these intensities was measured using microbial viability fluorophore assays: 1) a molecular assay to measure live and dead cells and 2) a fluorescein diacetate assay to measure relative metabolic activity of sludge micro-organisms. From the results of the viability assays, it was determined that MW had the greatest effect on cells, having several times greater cell death and inactivation than both US-pretreated and non-pretreated sludge, even at lower specific energy values. Additionally, a MW and US intensity with similar specific energy inputs (2.62 kJ/gTS and 2.37 kJ/gTS, respectively) were applied to feed sludge of bench-scale digesters to compare effects on AD over three consecutive sludge retention times (SRTs), at 20 d, 14 d, and 7 d. The MW-fed digester had the highest overall methane production (248.2 L CH4/kg volatile solids), greatest pathogen removal (73.4% and 69.8% less fecal coliforms over control during the 14 d and 7 d SRTs, respectively), and greatest solids removal (44.2% TS reduction). Interestingly, fecal coliform concentrations in the digester fed US-pretreated sludge increased over the control for both the 14 d (31%) and 7 d (39.6%) SRT. All digesters possessed positive net energy production over the three chosen SRTs.  iii Preface The results of this thesis was presented as a technical podium presentation and as a peer-reviewed conference paper at the Water Environment Federation (WEF)/ International Water Association (IWA) Residuals and Biosolids Conference: The Next Generation of Science, Technology, and Management in Washington DC, USA (June 7th – 10th 2015). The results of this thesis have also been submitted to Applied Microbiology and Biotechnology, as a journal paper, and is currently under review.   iv Table of Contents Abstract ............................................................................................................................. ii Preface .............................................................................................................................. iii List of Tables ................................................................................................................... vii List of Figures .................................................................................................................. ix List of Abbreviations ..................................................................................................... xvi Acknowledgements ...................................................................................................... xviii Dedication ....................................................................................................................... xix Chapter 1 | Introduction ................................................................................................... 1 1.1. Background and motivation for research ................................................................................. 1 1.2. Research goals .......................................................................................................................... 2 1.2.1. Application of microbial viability assays to sludge samples .............................................. 2 1.2.2. Running of three bench-scale, semi-continuously fed, AD systems .................................. 3 1.3. Hypotheses ............................................................................................................................... 4 1.4. Research scope ......................................................................................................................... 4 1.5. Novelty of research .................................................................................................................. 5 1.6. Thesis organization .................................................................................................................. 5 Chapter 2 | Literature Review ......................................................................................... 7 2.1. Characteristics of wastewater ................................................................................................... 7 2.2. General outline of the wastewater treatment process ............................................................... 8 2.3. Aerobic, anoxic, and anaerobic conditions used to treat wastewater ..................................... 10 2.4. Characteristics of waste sludges ............................................................................................. 15 2.5. Processes for stabilizing waste sludges .................................................................................. 17 2.6. Alkaline stabilization of sludge .............................................................................................. 18 2.7. Composting of sludge ............................................................................................................ 19 2.8. Aerobic digestion for sludge stabilization .............................................................................. 21 2.9. AD for sludge stabilization .................................................................................................... 22 2.10. Microbiology and biochemical pathways of AD ................................................................. 23 2.11. Important performance parameters for AD operation .......................................................... 26  v 2.12. Pretreatment of sludge prior to AD ...................................................................................... 30 2.13. Brief review of physical pretreatment methods in AD ......................................................... 32 2.13.1. Thermal pretreatments .................................................................................................... 32 2.13.2. Comparison of CH and MW pretreatments .................................................................... 33 2.13.3. Process of MW pretreatment .......................................................................................... 34 2.13.4. Mechanical pretreatments ............................................................................................... 36 2.13.5. Process of US pretreatment ............................................................................................ 37 2.14. Current applications of molecular assays ............................................................................. 39 2.14.1. S16:SO (live:dead) assay ................................................................................................ 42 2.14.2. FDA assay....................................................................................................................... 43 2.15. Summary .............................................................................................................................. 44 Chapter 3 | Methodology ................................................................................................ 45 3.1. Sludge sample collection ........................................................................................................ 45 3.2. Procedure for MW pretreatment of sludge ............................................................................. 45 3.3. Procedure for US pretreatment of sludge ............................................................................... 47 3.4. Selection of parameters to normalize pretreatment effects on sludge .................................... 48 3.5. Methodology for microbial viability characterization ............................................................ 49 3.6. Selection and preparation of tWAS pretreatment intensities and controls ............................. 49 3.7. Selection of microbial viability compounds for microbial characterization .......................... 52 3.8. Preliminary study of selected viability assays in wastewater samples ................................... 52 3.9. Assay optimization ................................................................................................................. 55 3.10. Microplate reader and fluorescence measurements.............................................................. 57 3.11. Methodology for AD digester performance comparison ..................................................... 58 3.12. Determination of pretreatment intensity for MW-AD digester feed .................................... 58 3.13. Determination of pretreatment intensity for US-AD digester feed ...................................... 60 3.14. Acclimatization of AD digester inoculum ........................................................................... 60 3.15. Set-up of bench-scale AD digesters ..................................................................................... 61 3.16. Start-up and operation of AD digesters ................................................................................ 62 3.17. Analytical measurements of biogas volume and composition ............................................. 63 3.18. Measurements of TS and VS ................................................................................................ 65 3.19. Fecal coliform detection ....................................................................................................... 66 3.20. Analytical measurements for tCOD and sCOD ................................................................... 67 3.21. Measurements of VFA concentrations ................................................................................. 68  vi 3.22. Alkalinity and pH measurements ......................................................................................... 69 3.23. Ammonia measurements ...................................................................................................... 69 3.24. Measurement of sludge dewaterability ................................................................................ 69 3.25. Net energy production .......................................................................................................... 70 3.26. Statistical analysis for microbial characterization ................................................................ 71 Chapter 4 | Results and Discussion ............................................................................... 72 4.1. Microbial characterization using S16:SO assay ..................................................................... 72 4.2. Microbial characterization using FDA assay ......................................................................... 75 4.3. Production of CH4 in AD digesters ........................................................................................ 77 4.4. VS and TS removals in AD digesters ..................................................................................... 79 4.5. Removal of tCOD in AD digesters ........................................................................................ 81 4.6. Alkalinity and pH ................................................................................................................... 82 4.7. Ammonia concentrations ....................................................................................................... 84 4.8. Dewaterability of AD digester effluents ................................................................................ 85 4.9. Fecal coliform detection in AD digester effluents ................................................................. 87 4.10. Net energy production from AD digesters ........................................................................... 89 Chapter 5 | Conclusion ................................................................................................... 92 5.1. Summary ................................................................................................................................ 92 5.2. Overall conclusion ................................................................................................................. 95 5.3. Recommendations for future work ......................................................................................... 96 Bibliography .................................................................................................................... 97 Appendices……………………………………………………………………………………115 Appendix A | Preliminary results for S16:SO and FDA assays .................................... 115 Appendix B | Calibration curves for measurements of AD data ................................... 116 Appendix C | Tukey Kramer method for post hoc analyses of assay data .................... 118 Appendix D | Statistical analyses for microbial characterization data .......................... 119 Appendix E | Correlation analysis for microbial characterization data ......................... 123  vii  List of Tables Table 2.1. End products of aerobic, anoxic, and anaerobic decomposition by heterotrophic micro-organisms in conventional wastewaters, in addition to new biomass (adapted from Pelczar and Reid, 1958; Davis and Cornwell, 2008). ............... 12 Table 2.2. Relative degree of attenuation of pathogen concentrations, putrefaction, and odour potential in sludge with various stabilization methods (adapted from Tchobanoglous et al., 2003). ............................................................................ 18 Table 2.3. Common bulking agents added to sludge to assist in stabilization during aerobic composting, and their respective characteristics and benefits (adapted from WEF, 1998; Tchobanoglous et al., 2003). ....................................................... 21 Table 3.1. Conditions of microwave irradiated (MW) and ultrasonicated (US) pretreatments chosen for microbial characterization. The live (CL) and dead (CD) controls were used for assay comparisons, and therefore their respective conditions are not shown. MW80 and US1060 were the only pretreated sludges used as feed for AD. ......................................................................................... 54 Table 3.2. Characterization of both non-pretreated control (CON), ultrasonicated (US1060), and microwave irradiated (MW80) mixed sludge feeds for AD digesters. ....... 62 Table 3.3. The energy needed to heat the digester sludges to mesophilic (35°C) temperatures (QH) was calculated using equation (3.4), and added to the energy required to pretreat the tWAS samples in order to determine the total input energy administered to the sludge added per digester. .................................... 71 Table D.1. ANOVA results and summary for S16:SO assay. Table includes the sum of squares (SS), degrees of freedom (df), mean of squares (MS), F value (F), P value, and the F critical value (F crit). ........................................................... 119 Table D.2. Tukey Kramer post-hoc analysis of S16:SO assay results for live control (CL), dead control (CD), microwave (MW) irradiated, and ultrasonicated (US) tWAS  viii samples at designated temperatures and intensities. The data are significantly different if the absolute difference (AD) between the two compared groups (Group 1 and Group 2) > calculated critical range (CR). Results show “true” for a statistically significant difference between the two groups under comparison, and “false” for differences between results that are not statistically significant. ........................................................................................................................ 119 Table D.3. ANOVA results and summary for FDA enzymatic assay. Table includes the sum of squares (SS), degrees of freedom (df), mean of squares (MS), F value (F), P value, and the F critical value (F crit). ................................................. 121 Table D.4. Tukey Kramer post-hoc analysis of FDA assay results for live control (CL), dead control (CD), microwave (MW) irradiated, and ultrasonicated (US) tWAS samples at designated temperatures and intensities. The data are significantly different if the absolute difference (AD) between the two compared groups (Group 1 and Group 2) > calculated critical range (CR). Results show “true” for a statistically significant difference between the two groups under comparison, and “false” for differences between results that are not statistically significant. ........................................................................................................................ 121 Table E.1. Result of correlation analysis indicating a positive direct linear correlation between the two microbial characterization assays, S16:SO and FDA assays. The sample size (n) is n = 11. ........................................................................ 124   ix List of Figures Figure 2.1. Flow of a generalized WWTP with primary and secondary treatment. Included is the flow of sludge through to the AD digester, and the final products, biogas and nutrient-rich biosolids (fertilizers). PS is primary sludge, MLSS is mixed liquor suspended solids, and WAS is waste activated sludge. ......................... 10 Figure 2.2. Diagram of a modified Bardenpho design, a system providing biological nutrient removal (BNR) of nitrogen and phosphorus at the WWTP in Kelowna, BC, Canada (adapted from Tchobanoglous et al., 2003). ................................ 13 Figure 2.3. Stages of composting for sludge isolated from wastewater in terms of time and temperature (adapted from Epstein, 1997). Vertical axis begins at ambient temperature of sludge. ...................................................................................... 20 Figure 2.4. Diagram showing the flow of substrate through the process of AD: hydrolysis, acidogenesis, acetogenesis, and methanogenesis (adapted from Appels et al., 2008). ............................................................................................................... 25 Figure 3.1. Set-up of MW irradiation of tWAS samples. The 12 pressure-sealed containers to hold tWAS samples are contained in a holding unit (top). The Microwave Lab station (Ethos EZ, 2450 MHz, 0-1200 W; maximum temperature of 300°C and maximum pressure of 35 bars) with an ATC-400-CE temperature probe is shown, with the holding unit containing the 12 pressure-sealed containers locked in place (bottom). ............................................................................................. 46 Figure 3.2. Set-up for US pretreatment of tWAS samples. The Fisher Scientific Model 500 Ultrasonic Dismembrator (with a maximum output of 400 W and frequency of 20 kHz), in combination with a Branson Model 102 Converter (101-135-066R) for Digital Sonifiers (with disruptor horn) is also shown with a beaker containing tWAS prior to US treatment (left). The disruptor horn is inserted into a beaker containing tWAS sample, which is held in a cold water bath containing crushed  x ice and plastic ice packs to minimize thermal effects during sonication (right). .......................................................................................................................... 47 Figure 3.3. The ¾ inch Branson Ultrasonic Disruptor Horn (stepped, tapped; 611-005-021) used for the selected US intensities for tWAS pretreatment. .................. 48 Figure 3.4. Microplate reader used for measuring fluorescence holding a 96-well black Nunc™ MicroWell™ optical-bottom microplate (top). A close-up of the MicroWell™ microplate (bottom) used for fluorescence measurements using the microplate reader. ....................................................................................... 59 Figure 3.5. The three (US1060, MW80, and CON) AD digesters used in this experiment and their set-up. ....................................................................................................... 64 Figure 3.6. Experimental outline of AD experiment for microwave irradiated tWAS feed at 80°C (MW80), ultrasonicated tWAS feed at 60% amplitude for 10 minutes (US1060), and non-pretreated (CON) tWAS feed. All feeds were mixed with primary sludge (PS) prior to being fed to the respective digester acclimatized to the designated pretreatment intensities. Also shown are the measurements performed with the biogas and biosolids (effluents) produced during the AD process over the 20 d, 14 d, and 7 d SRTs. ...................................................... 64 Figure 4.1. Ratios of live to dead cells (S16:SO) within sludge samples at various MW and US pretreatment intensities, CL, and CD (bar graph) based on level of fluorescence emitted (as arbitrary fluorescence units, AFU). Specific energy inputs of pretreatment intensities administered to tWAS samples are shown as kJ/g TS of sludge (scatter graph). Specific energy for CL is zero and was not considered for CD. CD: Dead Control (n = 8), CL: Live Control (n = 10), MW60: MW at 60°C (n = 4), MW80: MW at 80°C (n = 8), MW120: MW at 120°C (n = 6), MW160: MW at 160°C (n = 4), US1060: US for 10 min at 60% amplitude (n = 6), US2060: US for 20 min at 60% amplitude (n = 4), US2280: US for 22 min at  xi 80% amplitude (n = 4), US50100: US for 50 min at 100% amplitude (n = 4), US60100: US for 60 min at 100% amplitude (n = 4). ......................................... 73 Figure 4.2. FDA assay results as a function of specific energy input of pretreatment. Relative fluorescence (as arbitrary fluorescence units, AFU) emitted by metabolically active cells within sludge samples at various MW and US pretreatment intensities, CD, and CL (bar graph). Specific energy inputs of pretreatment intensities administered to the sludge samples are shown as kJ/g TS of sludge (scatter graph). Specific energy for CL is zero and was not considered for CD. CD: Dead Control (n = 7), CL: Live Control (n = 6), MW60: MW at 60°C (n = 6), MW80: MW at 80°C (n = 6), MW120: MW at 120°C (n = 6), MW160: MW at 160°C (n = 6), US1060: US for 10 min at 60% amplitude (n = 9), US2060: US for 20 min at 60% amplitude (n = 6), US2280: US for 22 min at 80% amplitude (n = 6), US50100: US for 50 min at 100% amplitude (n = 6), US60100: US for 60 min at 100% amplitude (n = 6). ......................................... 75 Figure 4.3. Specific daily biogas production (mL/d/L of digester) for microwave irradiated at 80°C (MW80), ultrasonicated for 10 minutes at 60% amplitude (US1060), and non-pretreated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs obtained at steady state…………………………………………………………………...77 Figure 4.4. Average daily specific CH4 production for microwave irradiated at 80°C (MW80), ultrasonicated for 10 minutes at 60% amplitude (US1060), and non-pretreated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent  average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of samples (n) = 64 for MW80, n = 62 US1060, and n = 65 for CON. At 14 d SRT, n = 50 for MW80, n = 51 US1060, and n = 49 for CON. At 7 d SRT, n = 50 for MW80, n = 48 US1060, and n = 51 for CON.. ...................................... 78 Figure 4.5. Total solids (TS) % removal for microwave irradiated at 80°C (MW80), ultrasonicated for 10 minutes at 60% amplitude (US1060), and non-pretreated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady  xii state. Data represent  average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of data points (n) = 76 for MW80, US1060, and CON. At 14 d SRT, n = 67 for MW80, US1060, and CON. At 7 d SRT, n = 59 for MW80, US1060, and CON. .............. 80 Figure 4.6. Volatile solids (VS) % removal for microwave irradiated at 80°C (MW80), ultrasonicated for 10 minutes at 60% amplitude (US1060), and non-pretreated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent  average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of  data points (n) = 76 for MW80, US1060, and CON. At 14 d SRT, n = 67 for MW80, US1060, and CON. At 7 d SRT, n = 59 for MW80, US1060, and CON. .............. 80 Figure 4.7. The % organics removal as depicted by total chemical oxygen demand (tCOD) of the AD digester effluents for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent average and error bars represent the standard deviations of data collected during each SRT over the steady state.  At 20 d SRT, number of data points (n) = 18 for MW80, US1060, and CON. At 14 d SRT, n = 14 for MW80, US1060, and CON. At 7 d SRT, n = 10 for MW80, US1060, and CON. .............. 81 Figure 4.8. Alkalinity (mg/L as CaCO3) of the AD digester effluents for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of data points (n) = 6 for MW80, US1060, and CON. At 14 d SRT, n = 4 for MW80, US1060, and CON. At 7 d SRT, n = 4 for MW80, US1060, and CON. ......................................................................................................... 83 Figure 4.9. The pH of the AD digester effluents for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control  xiii (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of data points (n) = 35 for MW80, US1060, and CON. At 14 d SRT, n = 15 for MW80, US1060, and CON. At 7 d SRT, n = 11 for MW80, US1060, and CON. .............. 83 Figure 4.10. The ammonia concentrations measured for the AD digester effluents for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of data points (n) = 8 for MW80, US1060, and CON. At 14 d SRT, n = 6 for MW80, US1060, and CON. At 7 d SRT, n = 4 for MW80, US1060, and CON. ................................................................................. 84 Figure 4.11. Dewaterability of digester effluents for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs, as seconds per total solids (TS % by weight) achieved at steady state during the last two weeks of measurements. At 20 d SRT, number of data points (n) = 11 for MW80, US1060, and CON. At 14 d SRT, n = 9 Data represent  average and error bars represent the standard deviations of data collected.for MW80, US1060, and CON. At 7 d SRT, n = 9 for MW80, US1060, and CON. ......................................................... 85 Figure 4.12. Concentrations of fecal coliforms detected in effluents for microwave irradiated at 80°C (MW80), ultrasonicated for 10 minutes at 60% amplitude (US1060), and non-pretreated control (CON) digesters, as well as for the non-treated feed sludge, over the 14 d and 7 d SRTs achieved at steady state. Data represent  average and error bars represent the standard deviations of data collected during the last two weeks of measurement. At 14 d SRT, number of data points (n) = 9 for MW80, US1060, and CON. At 7 d SRT, n = 6 for MW80, US1060, and CON. The limits for both Class A and Class B biosolids (BC OMRR,  xiv 2008) are displayed to show soil ammendment applicability. Fecal coliform levels were measured as colony forming units (CFU) per gram of dry sample solids. ............................................................................................................... 87 Figure 4.13. The net energy production (kJ/g VSfed) of the AD digesters produced over the 20 d, 14 d, and 7 d SRTs for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control (CON) digesters achieved at steady state. Data represent  average and error bars represent the standard deviations of data collected during steady state. At 20 d SRT, number of data points (n) = 64 for MW80, n = 62 US1060, and n = 65 for CON. At 14 d SRT, n = 50 for MW80, n = 51 US1060, and n = 49 for CON. At 7 d SRT, n = 50 for MW80, n = 48 US1060, and n = 51 for CON. ........................ 90 Figure A.1. Preliminary results in terms of live:dead cell ratios for S16:SO assay (n = 4) showing stepwise dilutions of  % live cells (CL) to % dead cells (CD). ....... 115 Figure A.2. Preliminary results for FDA assay (n = 6) showing fluorescence measurements (as arbitrary fluorescence units, AFU) of both live control (CL) and control dead (CD) tWAS samples over a 3 hour time period. Where the equation for live control is y = 47969x + 0.752, and the equation for dead control is y = 1111x + 148.45. ............................................................................................................ 115 Figure B.1. Calibration curve for biogas measurement using a manometer (at STP: 0°C, 1 atm), showing the height of the displaced volume within the manometer converted into volume of gas measured (equation is y = 5.0671x – 1E-05). . 116 Figure B.2. Calibration curve for chemical oxygen demand (COD), measured at 600 nm using a spectrophotometer (equation is y = 2359.5x – 0.9308). .................... 117 Figure B.3. Calibration curve for ammonia concentration (mg/L) measured against mV reading of samples (equation is y = -21.49ln(x) – 66.757). ........................... 117 Figure E.1. Graph of correlation analysis of the S16:SO (live:dead) cell ratios compared to the relative fluorescence of the FDA results. A positive correlation is  xv indicated by the increasing relative fluorescence to increasing live:dead cell ratio. Sample size (n) is n = 11. ...................................................................... 123     xvi List of Abbreviations AD Anaerobic digestion ANOVA Analysis of variance AOB Ammonia-oxidizing bacteria BNR Biological nutrient removal BOD Biochemical oxygen demand BOD5 5 day Biochemical oxygen demand CaO Quicklime Ca(OH)2 Slaked lime CD Dead control CFDA Carboxyfluorescein diacetate CFU Colony forming units CH Conventional Heating CH4 Methane CL Live control COD Chemical oxygen demand CON Control CO2 Carbon dioxide CST Capillary suction time DMSO Dimethyl sulfoxide DO Dissolved oxygen EPS Extracellular polymeric substances F:M Food to micro-organism ratio FDA Fluorescein diacetate GC Gas chromatograph MLSS Mixed liquor suspended solids MW Microwave irradiation MW60 Microwave irradiation at 60°C MW80 Microwave irradiation at 80°C MW120 Microwave irradiation at 120°C MW160 Microwave irradiation at 160°C N2 Nitrogen gas NH3 Ammonia NO2- Nitrite NO3- Nitrate NOB Nitrite-oxidizing bacteria OLR Organic loading rate PAO Phosphorus accumulating organisms  xvii PI Propidium iodide PO43- Phosphate PS Primary sludge RAS Return activated sludge S9 Syto® 9 Green S16 Syto® 16 Green S16:SO Syto® 16 Green to Sytox Orange ratio in a sample sCOD Soluble chemical oxygen demand SO Sytox® Orange SRT Sludge retention time tCOD Total chemical oxygen demand TS Total solids tWAS Thickened waste activated sludge US Ultrasonication US1060 Sonication for 10 minutes at 60% amplitude US2060 Sonication for 20 minutes at 60% amplitude US2280 Sonication for 22 minutes at 80% amplitude US50100 Sonication for 50 minutes at 100% amplitude US60100 Sonication for 60 minutes at 100% amplitude VBNC Viable but non-culturable (micro-organisms) VFA Volatile fatty acids VS Volatile solids WAS Waste activated sludge WWTP Wastewater treatment plant              xviii Acknowledgements First and foremost, I would like to express my deepest gratitude to my supervisor Dr. Cigdem Eskicioglu for her support and guidance, and for having faith in a lone microbiologist given free rein in her engineering laboratory. I would also like to thank the members of my committee, Dr. Rehan Sadiq and Dr. Sumi Siddiqua, for their time and expertise over the past few years.  My enduring thanks to Dr. Deniz Akgul for her mentorship in the UBC Bioreactor Technology Group laboratory, and without whose help I would still be up the creek without a paddle.   I would like to thank Dr. Kevin Hanna for introducing me to environmental management practices in BC, and for including me in his research. I’d also like to thank my fellow students and staff in the UBC Bioreactor Technology Group for their help and training.  I thank the City of Kelowna and BC Ministry of Environment for supporting this project, as well as for the sponsorships of the Natural Science and Engineering Council of Canada (NSERC) Strategic Project Grant (#396519-10) and the financial support of the Scientific and Technological Research Council of Turkey (TUBITAK).   I would like to express a special thanks to my parents, Claudio and Judith Cella, for their unending support and patience. I’d also like to thank my brother Matthew Cella for always reminding me to kick back and relax once in a while. Lastly, I would like to thank Dr. Ali Ahmadi for his unfaltering encouragement throughout the years.    xix  Dedication  This work is dedicated with love to those who came before me.  Angela Cella Sergio Cella  Gertrude Elizabeth Dufresne Leopold Dufresne     1  Chapter 1 | Introduction 1.1. Background and motivation for research As the world’s population continuously increases, solutions must be found to manage the large amounts of waste that is produced. In particular, the disposal of untreated wastewater is a global burden and the mishandling of which can lead to environmental damage, contamination of drinking water sources, unhygienic living conditions, and disease (Khursheed and Kazmi, 2011). Previous methods of waste sludge or simply “sludge” (generated during municipal wastewater treatment) disposal have included incineration, landfilling, and pumping at great depths in large bodies of water, as well as composting. The majority of these methods are either no longer sustainable, contribute greatly to greenhouse gas emissions, or are overwhelmed by the enormous volumes of sludge that must be disposed of daily.  It is no longer acceptable to simply dispose of municipal waste sludge, but to instead develop methods of using this waste as a source of viable energy and products in order to meet increasing demands on the world’s resources (Chynoweth et al., 2001). Although there are many versions of achieving this goal, an ideal solution to remedy this problem is to further stabilize the residual sludge generated by wastewater treatment plants (WWTPs), rather than simply sending it to a landfill; landfilling is no longer an acceptable disposal route for most of the organic waste streams. This may be performed by engineering biochemical processes that can convert organic materials found in sludges into useable products. One such method is the use of anaerobic digestion (AD). AD is a biochemical process that ultimately converts organics (i.e. fats, sugars, and proteins found in sludge) into methane (CH4) gas (a source of electrical and heat energy) as well as nutrient-rich fertilizers that may be used as soil amendments or conditioners for a variety of agricultural needs. Additionally, AD is favoured due to its ability to stabilize high-strength organic waste, reduce greenhouse gas emissions, and divert organic waste from landfills (Botheju and Bakke, 2011). To date, AD has been applied to numerous full-scale digester systems at WWTPs and also as farm-based ADs, and there is now a shift to optimizing this  2  biological process.  Developments over the past few years have indicated that the pretreatment (disintegration) of sludges containing complex organics into simpler compounds prior to AD has reduced the time needed for AD, as well as increasing CH4 production in smaller AD digesters. Sludge pretreatment via mechanical (i.e. ultrasonication, stirred ball mills, high pressure homogenization), thermal (i.e. microwave irradiation, conventional heating), chemical (i.e. ozonation, acidic and alkaline degradation), and biological (i.e. enzymatic degradation) methods have led to improvements in AD (Kim et al., 2003; Carrère et al., 2010; Ge et al., 2010; Salsabil et al., 2010). The majority of this past and ongoing research has focused on conventional performance parameters of advanced AD systems, such as biogas production, solids removal, dewaterability, and pathogen removal; however, recent advancements in the understanding of the mechanisms behind engineered biological processes, such as pretreatments in AD systems, have helped to optimize conventional systems, as well as develop new ones (Narihiro and Sekiguchi, 2007). 1.2. Research goals The goal of this thesis was to understand the extent of cellular lysis or disruption in micro-organisms caused by mechanical and thermal pretreatments of thickened waste activated sludge (tWAS) generated by treatment plants prior to AD, and how this cellular destruction translates to enhancements in AD processes. This goal was achieved by implementing two experimental studies, which are described in sections 1.2.1 and 1.2.2. 1.2.1. Application of microbial viability assays to sludge samples In the first part of the study, two microbial viability assays were applied to a wide range of pretreated (thermal and mechanical) sludges in order to characterize the effects of the pretreatments on sludge micro-organisms. The two assays chosen included: a) Syto 16® Green (S16) and Sytox® Orange (SO) assay to discern the integrity of cellular membranes in the sludge samples, and   3  b) Fluorescein diacetate (FDA) assay to determine the relative enzymatic activity of microbial cells within the sludge samples. Both assays were conducted using a fluorescence detecting, high throughput microplate reader. The range of pretreatments included five different ultrasonication (US) intensities as a mechanical pretreatment, and four temperatures reached via microwave irradiation (MW) as a thermal pretreatment; these respective ranges were chosen based on similar specific energy inputs to the tWAS. A detailed explanation of the microbial assay methodology and selection of the pretreatment ranges are described in section 3.5. 1.2.2. Running of three bench-scale, semi-continuously fed, AD systems The second part of this study involved the running of three bench-scale, semi-continuously fed AD systems. Each digester was fed one of the following:  a) non-pretreated tWAS (control),  b) US-pretreated tWAS (mechanical pretreatment), and  c) MW-pretreated tWAS (thermal pretreatment).  The effects of pretreatments on AD efficiency were compared and measured for each of the three digesters in terms of the following: a) total (TS) and volatile (VS) solids removal, b) total chemical oxygen demand (tCOD) removal,  c) biogas production, d) CH4 composition of biogas produced, e) alkalinity and pH f) ammonia g) dewaterability of digested effluent sludge, h) fecal coliform densities of digested effluent sludge to be land applied, and  4  i) net energy production (input energy for pretreatment – output energy via CH4 generation = net energy). A detailed explanation of the complete methodology for the three AD digesters is described in chapter three. 1.3. Hypotheses The hypotheses of this research are as follows: 1. It was expected that both AD digesters fed pretreated tWAS would produce a greater amount of CH4 gas and have a greater fecal coliform destruction than the control (non-pretreated) digester.  2. The level of enzymatic activity and cellular integrity of microbial cells within the pretreated samples would be much lower than those found in the non-treated control samples.  3. Greater cellular disruption expected in microbially characterized (i.e. assayed) pretreated tWAS samples would correspond to greater overall AD efficiency within the two digesters fed pretreated tWAS.  4. Based on previous literature (see chapter two), it was also hypothesized that MW-pretreated tWAS would provide better substrate for AD processes than US-pretreated tWAS feed, due to innate abilities of both thermal and athermal (i.e. dipolar rotation of molecules leading to bond breakage) MW effects, and that this will be reflected in both the AD efficiency and microbial characterization of MW-pretreated tWAS samples. 1.4. Research scope The research scope of this thesis, in order to validate the hypotheses stated, is as follows: a) To successfully apply microbial viability assays to understand the connection between cellular disruption, expulsion of intracellular substrate into the liquid fraction of sludge samples, and AD efficiency.   5  b) To understand and compare the effects of a thermal (MW) and a mechanical (US) pretreatment on tWAS, each with similar specific energy input values (2.62 and 2.37 kJ/ g TS, respectively), in semi-continuously fed, bench-scale AD systems at sludge retention times (SRTs) of 20 d, 14 d, and 7 d.  c) To determine if additional factors, besides the amount of specific energy applied, potentially contribute to cellular disruption within pretreated tWAS samples. d) To observe the optimum levels of specific energy inputs to tWAS samples (i.e. the minimum amount of energy needed to obtain maximum cell disruption, according to the applied cell assays, using a specific pretreatment) before the treatment process becomes cost-inhibitive. 1.5. Novelty of research Although there is previous literature comparing pretreatment of sludge feed prior to AD, this is the first time that two different pretreatment types (a thermal and a mechanical pretreatment) are compared based on the specific energy inputs applied to the sludge feed, and compared in terms of biogas production, CH4 composition, tCOD % removals, fecal coliform destruction, TS and VS % removals, dewaterability, and net energy production over the course of three different SRTs. Moreover, to the best of my knowledge, it is the first time that two different microbial characterization assays were successfully applied to a range of pretreated sludge samples, again based on specific energy inputs of both thermal and mechanical pretreatments (including the intensities used for the AD digesters), and measured using a high throughput microplate reader.  1.6. Thesis organization This thesis is organized into five chapters. The first chapter provides a brief background to the study, and also discusses the scope, hypotheses, and novelty of the research. The second chapter entails the literature review, which is written to give the reader a broader insight to wastewater treatment, sludge treatment and disposal, applications of AD  6  technology, and pretreatment processes. This chapter will also include information on the microbiology of wastewater, as well as previous and current applications of microbial viability assays and fluorophores. The third chapter discusses the materials and methodology used in this research. The results and discussion of the research are provided in chapter four. Finally, chapter five contains the final conclusions, research contributions, and recommendations for future studies.      7  Chapter 2 | Literature Review This chapter will provide an in-depth discussion in regards to the general process of wastewater treatment, the basic microbiology of wastewater, AD processes, current pretreatments of waste sludge prior to AD, and existing applications of fluorophores in molecular assays, among other topics. 2.1. Characteristics of wastewater The composition of wastewater is dependent on the communities that a WWTP services. In general, wastewater can be categorized into two main groups: municipal (domestic) wastewater and industrial wastewater. Industrial wastewaters are generated as a result of water used for cooling, production, or cleaning purposes in textile industries, pulp and paper mills, slaughterhouse and dairy units, and distilleries, among others. This type of wastewater can contain extremely high organic contents, and toxic compounds such as high metal concentrations and phenol (Rajeshwari et al., 2000). Municipal wastewater refers to the domestic water wasted from urban and suburban areas of a municipality, such as from household toilets and sinks, grey water, and storm waters. For the purpose of this study, this literature review will focus on the characteristics and treatment of municipal wastewater from residential areas. The most common solid components of municipal wastewater include suspended and dissolved organic materials. These materials include sugars, lipids, and proteins; in addition to these compounds, inorganic particulates and minerals (i.e. nitrogen, sulphur, and phosphorus) are also present. Moreover, microbial cells are also existent in the wastewater, and can utilize the organic compounds in wastewater for energy and reproduction. Additionally, these microbial cells secrete extracellular polymeric substances (EPS) through their own metabolic processes or during autolysis. EPS compounds are composed of aggregated polymers. These polymers include uronic acids, lipids, nucleic acids, proteins, humic acids, and polysaccharides, among other compounds, and greatly contribute to the overall composition of wastewater sludge. Therefore, it is the  8  primary goal of wastewater process engineers to design treatment processes that optimize the removal of these organics in order to minimize residual sludge as well as odours caused by putrescible organic material (Corbitt, 1990). As treated wastewater is often discharged back to sources of water supplies (i.e. lakes, reservoirs), the sustainability of water sources is dependent on the efficiency of wastewater management (Carey and Migliaccio, 2009). The ultimate goal of WWTPs is to separate the solids from the liquid fraction of wastewater; the liquid fraction (i.e. treated water) is treated to near-palatable levels and returned to a lake or water reservoir. However, the isolation, treatment, and disposal of the residual excess sludges, the byproduct of physical, chemical and biological treatment of wastewater, can be difficult and cost-intensive. This study is centered on these excess sludges and corresponding issues. 2.2. General outline of the wastewater treatment process WWTPs are necessary to improve the quality of wastewater released to surface or groundwater supplies in order to prevent toxic and eutrophic events. Additionally, over the past fifty years, many countries have strived to reduce the overall volume of excess sludges isolated from wastewater by carefully monitoring and upgrading WWTPs (Tchobanoglous et al., 2003; Carey and Migliaccio, 2009). There are several components to conventional treatment of municipal wastewater:  1) Primary treatment, 2) Secondary treatment, and, in some cases, 3) Tertiary treatment. Wastewater which first enters the WWTP treatment stream will go through primary treatment. During this primary treatment, unit operations such as screening, grit removal, and primary sedimentation remove both coarse materials (e.g. rags, garbage, silt) and finer settleable particles. The settleable organics removed during this process are collectively termed as primary sludge (PS). The organics that remain in the wastewater after primary  9  treatment are then subjugated to secondary treatment, which consists of oxidizing both suspended and soluble organic matters by a variety of micro-organisms. This secondary treatment is also termed biological treatment, and consists of an activated sludge treatment process in which the wastewater organics are converted into carbon dioxide (CO2), water, and active biomass (Rocher et al., 1999). This process occurs in an aerated tank. The heterogeneous culture of micro-organisms that oxidize these organic materials during secondary treatment are generally termed as mixed liquor suspended solids (MLSS). Sludges removed from this process are termed waste activated sludge (WAS) as they consist of both EPS and the MLSS. In most WWTPs, a certain amount of the WAS is returned to the aeration tank in order to maintain the MLSS concentration. However, a large portion of the MLSS must be wasted along with the WAS. Essentially, this is due to a drastic spike in microbial reproduction during the aeration process prior to secondary settling. By using oxygen (O2) as a high-affinity electron acceptor with a strong reduction potential difference, aerobic treatment of wastewater allows for more energy to be produced by bacteria within the aeration tank, thus allowing for higher rates of biomass reproduction (Lovley and Goodwin, 1988). Unfortunately, the use of aerobic biological treatment of domestic wastewater leads to high biomass growth that must be disposed of in addition to the inorganic solids removed from the wastewater. Following secondary treatment, tertiary treatment is implemented in WWTPs when a higher quality of wastewater is required before discharge to a receiving body of water. Tertiary treatment may include a variety of processes, including nutrient (i.e. nitrogen and phosphorous) removal, ultra-violet light disinfection, chlorine addition, granular filtration, or carbon adsorption, among others, and is specific to the type of wastewater being treated (Droste, 1997). The PS or WAS removed from the treatment process is often dewatered or thickened, and digested in order to stabilize these sludges for handling, transport, and final disposal. All WWTPs possess primary treatment, and the majority also include secondary treatment in their processing of wastewater. However, not all municipal wastewaters necessitate tertiary treatment in order to meet the nutrient and organic pollutant discharge requirements by law within a country. A generalized schematic of municipal wastewater  10  treatment is shown in Figure 2.1.  Figure 2.1. Flow of a generalized WWTP with primary and secondary treatment. Included is the flow of sludge through to the AD digester, and the final products, biogas and nutrient-rich biosolids (fertilizers). PS is primary sludge, MLSS is mixed liquor suspended solids, and WAS is waste activated sludge. 2.3. Aerobic, anoxic, and anaerobic conditions used to treat wastewater The manipulation of environmental conditions is a crucial aspect of wastewater treatment design. Stabilization of organic pollutants in wastewater has traditionally been achieved through biological means under aerobic, anoxic, or anaerobic conditions, as well as combinations of these through WWTPs, as described in Figure 2.1 (Kim and Hao, 1990). The definition of aerobic, anoxic, and anaerobic conditions are based on the availability of terminal electron acceptors that may be utilized by micro-organisms during metabolic processes. Electron acceptors are compounds that are able to be reduced by micro-  11  organisms in order to produce energy via the oxidation of an electron donor. Wastewaters under aerobic conditions possess free molecular O2 that is available as a terminal electron acceptor for respiration or in metabolic oxygenation. Under aerobic conditions, heterotrophic micro-organisms metabolize the organic materials present in wastewater via oxidization. In this process, the free O2 is used to oxidize some of the carbon present in organic materials to CO2; any residual carbon is then available to be used as substrate to produce new biomass (NRC, 1993). Moreover, during this process, O2 is reduced, resulting in the production of water molecules. Subsequently, the process of biodegradation under aerobic conditions results in the final production of CO2, water, some minerals, energy, and new microbial biomass from the oxidized organics (Tchobanoglous and Schroeder, 1985). Aerobic conditions have been described to occur at dissolved O2 (DO) concentrations of 2 mg/L or greater (Hai et al., 2011). Compared to both anaerobic and anoxic conditions, aerobic decomposition results in much greater production of biomass, due to the large amount of energy released during aerobic oxidation, as previously explained. Therefore, although aerobic conditions are favoured for treatment of wastewaters with dilute concentrations of organics (possessing a 5 day biochemical oxygen demand, or BOD5, of less than 500 mg/L), other methods of stabilization are preferred for high-strength wastewaters (i.e. possessing a BOD5 greater than 1000 mg/L) due to energy cost and difficulties in proper aeration and the large volume of biological excess sludges produced. However, aerated lagoons have been used in the treatment of smaller volumes of wastewater containing a BOD5 of 3000 mg/L or more (Davis and Cornwell, 2008).  In wastewater treatment, the term “anoxic” is used to describe environmental conditions that possess none or extremely low concentrations of DO, which has been previously described as less than 0.5 mg DO/L (Xing et al., 2006). Within wastewaters, anoxic conditions differ from anaerobic conditions as only nitrate (NO3-) and nitrite (NO2-) are utilized as electron acceptors, resulting in the production of gaseous nitrogen during biological denitrification. In anaerobic conditions, one of the most common electron acceptors, O2, is absent. Subsequently, micro-organisms present in these conditions must  12  utilize other forms of electron acceptors, such as sulphate or CO2. The representative end products of aerobic, anoxic, and anaerobic decomposition within wastewaters are detailed in Table 2.1. Table 2.1. End products of aerobic, anoxic, and anaerobic decomposition by heterotrophic micro-organisms in conventional wastewaters, in addition to new biomass (adapted from Pelczar and Reid, 1958; Davis and Cornwell, 2008). Substrates Aerobic and anoxic decomposition Anaerobic decomposition Organic nitrogen-containing compounds (i.e. proteins)  Amino acids  Ammonia  nitrites  nitrates  Nitrates  nitrogen gas*  Alcohols  CO2 + H2O  Organic acids  Amino acids  Ammonia  Hydrogen sulphide  CH4  CO2  Alcohols  Organic acids Carbohydrates (i.e. glucose)  Alcohols  CO2 + H2O  Fatty acids  CO2  Fatty acids  CH4 Fats and related substances  Fatty acids + glycerol  Alcohols  CO2 + H2O  Lower fatty acids  Fatty acids + glycerol  CO2  Alcohols  Lower fatty acids  CH4 *Only under anoxic conditions  When applied to wastewater treatment, aerobic and anaerobic conditions are often favoured to stabilize carbonaceous organic matter. Anoxic conditions are most commonly used in combination with aerobic and/or anaerobic conditions in order to reduce nutrient (i.e. nitrogen and phosphorous) concentrations during biological nutrient removal (BNR) processes within tertiary wastewater treatments. An example of a BNR process includes the modified Bardenpho design, which has been implemented at many full-scale WWTPs with both carbon and nutrient discharge limits, including the WWTP in Kelowna, BC,  13  Canada. In this process, carbon, phosphorus and nitrogen concentrations in the wastewater effluent are decreased via biological systems made up of a combination of manipulated environmental conditions. The modified Bardenpho design is described in Figure 2.2.   Figure 2.2. Diagram of a modified Bardenpho design, a system providing biological nutrient removal (BNR) of nitrogen and phosphorus at the WWTP in Kelowna, BC, Canada (adapted from Tchobanoglous et al., 2003). In the first stage of the modified Bardenpho design, an anaerobic tank is provided to produce volatile fatty acids (VFAs) as substrate in the form of BOD for phosphorus accumulating organisms (PAOs). The uptake of BOD by PAOs requires energy, and in purely anaerobic conditions (i.e. no DO to act as an electron acceptor), the PAOs obtain the required energy by the cleaving of phosphate (PO43-) molecules from their own intracellular PO43- storage granules. PO43- then moves across the cell membrane of PAOs, and into the MLSS matrix. During this first stage, there is an overall decrease in BOD and an increase in PO43- within the anaerobic tank (Ross, 2013). Moreover, by maintaining a shorter SRT in the anaerobic tank, CH4 production is minimized. In the second stage, the influent moves into an anoxic tank, in which some additional BOD removal occurs, as well as biological denitrification. During this process, nitrogen gas (N2) is produced along with excess biomass and CO2 by the biodegradation of organic materials and the use of NO3- as  14  the terminal electron acceptor. As the majority of these heterotrophic bacteria are able to use both O2 and NO3- as an electron acceptor, the presence of O2 in the second tank will inhibit denitrification; this is caused by preferential selection of electron acceptors with higher redox potential (i.e. O2) by these micro-organisms. Moreover, alkalinity is formed in this stage. In the third stage, the wastewater flows to an aerobic tank. In this stage, PAOs absorb PO43- molecules from the MLSS and again store the PO43- in intracellular granules using energy obtained from the BOD uptake in the initial anaerobic tank (via DO as the electron acceptor). Poor phosphorus uptake in the aerobic tank can be caused by diluted levels of BOD (i.e. VFAs) in the anaerobic tank. In addition to PO43- uptake, nitrification occurs in the aerobic tank, causing the production of additional nitrogen precursors for additional denitrification later in the BNR process. Nitrification is a two-step biochemical process performed by two types of bacteria, ammonia-oxidizing bacteria (AOBs) and nitrite-oxidizing bacteria (NOBs), as shown below.  1) NH4+ +1.5O2  NO2- + 2H+ + H2O, (via AOBs) 2) NO2- + 0.5O2 NO3-, (via NOBs) Nitrosomonas spp. (i.e. Nitrosomonas europaea) and Nitrosospira spp. (i.e. Nitrosospira tenuis) are common AOBs found in wastewater treatment systems; it has been reported that the activity of these nitrifying species is dependent on the concentration of ammonia (NH3) present in their surrounding environment, with Nitrosomonas being more active in high NH3 concentrations, and Nitrosospira more dominant in low NH3 concentrations (Aoi et al., 2000). NOBs commonly present in WWTPs include Nitrococcus spp. (i.e. Nitrococcus mobilis), Nitrobacter spp. (i.e. Nitrobacter winogradskyi), and Nitrospira spp. (i.e. Nitrospira marina) among others. Siripong and Rittmann (2007) have reported that cooler temperatures (i.e. during winter months) of influent industrial wastewaters to full-scale water reclamation plants can result in greater diversity of NOB species, in particular the Nitrospira spp., and that seasonal temperature fluctuations have a greater impact on NOB diversity than other parameters that were analyzed during the study, such as SRT. Additionally, there is some BOD removal in the aerobic tank. Nitrifying bacteria do not  15  use organic carbon as a growth basis, but instead use the alkalinity present in the tank as their electron donating compounds. Thus, enormous decreases in alkalinity can be seen in the aerobic tank, which is offset by the production of alkalinity during the initial anoxic condition. However, if the alkalinity of the aerobic tank decreases to less than 50 mg/L as CaCO3, the pH will significantly decrease. The optimum pH for nitrification is 7 – 8.5, and a pH below 6 will cause inhibition of the AOBs and NOBs in the tank. Also, a portion of the MLSS containing high concentrations of NO3- from the aerobic tank is often recycled back to the initial anoxic tank in order to assist in denitrification. In the fourth tank, a second anoxic condition exists, and residual NO3- is converted to N2 via denitrification. The electron donors used in this step is provided by the system biomass progressing through endogenous decay, which may also be supplemented with carbon addition in the form of methanol. The electron acceptor used in this step is again NO3- (Droste, 1997). Lastly, the fifth tank contains a final aerobic condition, in which N2 is stripped from solution. These aerobic conditions also prevent the use of PO43- as an electron acceptor by PAOs during respiration, and sufficient DO concentrations are necessary to prevent PO43- release into the clarifier following BNR (Ross, 2013). The overall process contains an SRT of 10 – 20 d, which allows for the increased oxidation of the wastewater organics. After the BNR system, the effluent is sent to a final clarifier to remove any remaining settleable solids; some of the WAS is often returned to the beginning of the BNR system, and is called return activated sludge (RAS). The treated wastewater is sent for further disinfection and subsequent release to a receiving body of water. BNR systems are modifications of the most common activated sludge processes (as described in Figure 2.1) for tertiary (nutrient) treatment, and are examples of the importance of understanding the micro-organisms and biochemical processes within engineered biological systems. 2.4. Characteristics of waste sludges In addition to wastewater treatment, WWTPs must stabilize waste sludge generated as a result of wastewater treatment in an environmentally sustainable way. The innate characteristics of sludges isolated from various wastewaters are dependent on which treatment level the sludge is removed from the wastewater treatment flow, as well as the  16  wastewater itself, and are named accordingly. As described previously, PS is removed from the treatment train in the first (primary) stage of the WWTP. Due to the high water content of PS (> 95%), PS is often thickened (i.e. water is removed) prior to handling. This type of sludge is high in organics readily available for biodegradation (Kropp and Ditchtl, 2001). Additionally, sludge stabilization is particularly important for PS, as PS has a much higher concentration of pathogens and putrescible organic materials than WAS. Due to a combination of these and other characteristics (i.e. less EPS and colloidal matter), PS has been reported as having an overall better biodegradability and dewatering than WAS (Turovskiy and Mathai, 2006).  Secondary sludge, or WAS, is removed from the WWTP after secondary treatment processes. During the process of aeration, residual organics, inorganics, minerals, EPS, and MLSS aggregate together to form WAS “flocs”. Nielsen (2002) reported the relative % composition of the organic materials typically found in WAS flocs, including cell biomass (10 – 20%), fibrous organic material (10 – 30%), and EPS (40 – 60%). It was also shown that the EPS in WAS contained 40 – 60% protein, 20 – 30% humic acids, 10 – 20% polysaccharides, 2 – 5% nucleic acids, and 5 – 10% of additional compounds such as lipids (Nielsen, 2002; Park and Helm, 2008; Park et al., 2008). Floc properties (such as size, shape, density, and strength) contribute to the characteristics seen in sludges, such as flocculation, settling ability, membrane fouling, and dewaterability (Nielsen et al., 2012). In particular, flocculation and accumulation of microbial cells within flocs are usually caused by low nutrient conditions within the environment; as cells accumulate together, the products released by one group of cells are used as substrate by another cell cluster. The proximity of these cells facilitates an efficient exploitation of food and waste distributions (McLoughlin, 1994; Olofsson et al., 1998). Therefore, sludge settling, a crucial characteristic in the design of primary and secondary treatments, depends on the food to micro-organism (F:M) ratio and the age of the sludge. Poor sludge settling may also be caused by fluctuations in pH, temperature, lack of nutrients, or the presence of compounds toxic to micro-organisms such as heavy metals. The EPS secreted by micro-organisms found in WAS also contribute to the flocculation of sludge particles, often  17  trapping water in these flocs and making dewatering and handling much more difficult. These sludges also possess high concentrations of nutrients, such as nitrogen and phosphorus, which are important parameters to consider in wastewater treatment design as these nutrients can cause eutrophication of receiving body waters (Kropp and Ditchtl, 2001). 2.5. Processes for stabilizing waste sludges As greater amounts of wastewater solids must be continuously isolated and treated by WWTPs, there is a need to find methods of safely disposing the excess sludges. Conventional methods, such as incineration of sludge and landfilling, often possess large carbon footprints. Currently, there is a shift to applying biochemical processes to these sludges in order to produce useful resources, in addition to reducing the large carbon footprint of conventional disposal methods (Barber, 2009), as detailed in the following sections. As discussed in section 2.3, various environmental conditions are manipulated to achieve removal of organics and nutrients from wastewater. Similar methods are also employed to stabilize the residual sludge (i.e. WAS and PS removed from the WWTP) generated during wastewater treatment. The purpose of sludge stabilization is to achieve the reduction of pathogens, eliminate offensive odours, reduce the overall sludge volume, minimize the potential of the sludge to putrefy, and, in some cases, improve sludge dewaterability (WEF, 1998; Tchobanoglous et al., 2003). Moreover, sludge stabilization can result in the production of nutrient-rich biosolids, which may be used as soil fertilizers or conditioners. There are many variations of sludge stabilization, the four most common being alkaline stabilization, composting, aerobic digestion, and AD. The ability of these methods to attenuate the potential of sludge to putrefy, produce odours, and contain pathogens are summarized in Table 2.2. Once sludge has been stabilized, it is often able to be used for beneficial purposes and is thusly called “biosolids” (WEF, 1998). The following sections further discuss the benefits and disadvantages of these stabilization methods. Although sludge stabilization is not implemented at all WWTPs, these methods have been  18  successfully used at WWTPs treating various volumes of wastewater (Tchobanoglous et al., 2003). Table 2.2. Relative degree of attenuation of pathogen concentrations, putrefaction, and odour potential in sludge with various stabilization methods (adapted from Tchobanoglous et al., 2003).  Degree of attenuation Stabilization method Pathogens Putrefaction Odour potential Alkaline stabilization Good Fair Fair Anaerobic digestion Fair Good Good Aerobic digestion Fair Good Good Composting Fair Good Poor to fair Composting (thermophilic) Excellent Good Poor to fair  2.6. Alkaline stabilization of sludge During alkaline stabilization, chemical addition (most commonly in the form of lime) is administered to the sludge in order to raise the pH > 12. The high pH halts the biological activity of micro-organisms within the untreated sludge, which subsequently prevents putrefaction, vector attraction, and production of odours. In addition to the increase in pH, if the chemical addition includes high concentrations of quicklime (CaO), the CaO will react with both CO2 and water in exothermic reactions, releasing approximately 64 kJ/g mole and 180 kJ/g mole, respectively (US EPA, 1983). These reactions can lead to large increases in sludge temperature, which have been reported at greater than 50°C in some cases (Roediger, 1987). The typical lime dosages, as slaked lime or Ca(OH)2, required to stabilize PS, WAS, and septic sludge are 60 – 70 g Ca(OH)2/ kg TS, 210 – 430 g Ca(OH)2/ kg TS, and 90 – 510 g Ca(OH)2/ kg TS, respectively (WEF, 1995). As the % TS concentration of the sludge increases, the lime dose also increases. The decay of pH over time is a significant problem during alkaline stabilization, as lime addition is not able to  19  destroy the organic substrate used by pathogens as pH levels falter and decrease. Therefore, recommended design objectives outlined by governing bodies must be followed closely to prevent reactivation of micro-organisms caused by decreasing pH. The general minimal criterion for lime stabilization is to maintain the pH at > 12 for a period of at least 2 h, and to provide additional alkalinity within the treated sludge to prevent the pH from falling to less than 11 for several days (WEF, 1995). An additional disadvantage of alkaline stabilization of sludge is the substantial increase in product mass (to be disposed of) caused by adding lime to sludge bulk. 2.7. Composting of sludge Composting has been practiced by people since the first agriculture-based settlements. It is a cost-effective method of sludge stabilization that results in the decomposition and stabilization of the organic fraction of sludge by a succession of micro-organisms. Most commonly, composting of sludge is performed in mostly aerobic conditions, although there are anaerobic conditions that occur at the deepest layers of sludge held in composting containers. The use of aerobic conditions allows for the acceleration of the composting time needed for organic decomposition, causes greater exothermic reactions that result in higher temperature changes within sludge, and reduces the odours produced (Tchobanoglous et al., 2003).  There are three stages of composting. In the first mesophilic stage, acid-producing bacteria and fungi begin to break down the organic fraction, causing exothermic reactions that result in the increase of sludge temperature from ambient to 40°C. In the second stage, thermophilic micro-organisms (fungi, actinomycetes, and bacteria) replace the mesophilic micro-organisms as the temperatures steadily rise from 40°C to around 70°C. During this period of pasteurization, maximum biodegradation of the organic fraction occurs; it has been reported that composting can result in a 20 – 30% VS decrease in organic solids in sludge. In order to kill pathogens, weeds, and seeds entrapped within the sludge, the thermophilic stage must be maintained for a minimum period of 24 h. In the final stage, microbial activity begins to decrease and the sludge temperature cools. Humic acid and pH  20  stabilization as well as additional water evaporation occur at this cooling stage. The time needed for the progression of these three stages of composting is several weeks. However, the composted biosolids must then be held for a final maturation stage, during which microbial populations decline and the residual organics finish stabilizing. This maturation (curing) stage can take several weeks to a year to complete (Vergara, 2012). The outline of sludge composting is shown in Figure 2.3. The resulting product (compost) is often used as fertilizers, mulch, soil conditioners, or as peat substitutes for a variety of agricultural needs.   Figure 2.3. Stages of composting for sludge isolated from wastewater in terms of time and temperature (adapted from Epstein, 1997). Vertical axis begins at ambient temperature of sludge.  In some cases, amendments or bulking agents are added to sludge to assist in composting and odour mitigation. An amendment is an organic material added to reduce the bulk weight of the sludge, decrease moisture content, and increase air voids within the sludge to improve aeration during composting. Common amendment materials include straw, rice hulls, recycled compost, and sawdust. A bulking agent is either an organic or inorganic material that provides structural support to the bulk sludge, as well as increasing porosity to improve aeration. Examples of bulking agents and their benefits are described in Table High-rate composting (maximum organic degradation) Thermophilic (°C) Mesophilic (°C) Maturation (curing) Mature and stable compost Time (days)  Temperature (°C) Cooling 40 70 20 1 365 5  21  2.3. During the thermophilic stage of composting, actinomycetes and fungi often break down cellulose content found within amendments or bulking agents. Following the thermophilic and cooling stages, recovery of a bulking agent is possible (WEF, 1998). Table 2.3. Common bulking agents added to sludge to assist in stabilization during aerobic composting, and their respective characteristics and benefits (adapted from WEF, 1998; Tchobanoglous et al., 2003). Bulking agent Characteristics Wood chips  May require purchase  High recovery rate via screening  Provides supplemental carbon source  Chipped brush  Possibly available as a waste material  Low recovery rate by screening  Provides supplemental carbon source  Leaves and yard waste  Must be shredded  Wide range of moisture content  Readily available source of carbon  Relatively low porosity  Non-recoverable  Shredded tires  Often mixed with other bulking agents  Supplemental carbon is not available  Nearly 100% recoverable  May contain metals  Ground waste lumber  Possibly available as waste material  A poor source of supplemental carbon 2.8. Aerobic digestion for sludge stabilization Aerobic digestion of sludge has been implemented at a range of WWTPs treating wastewater flows of 0.2 – 2 m3/s (WEF, 1998). The process of aerobic digestion of sludge isolated from wastewater is similar to the activated sludge process of influent wastewater in WWTP (refer to Figure 2.1). In the aerobic digestion of sludge, the electron donor (i.e.  22  organic compounds) is purposefully kept limited. As the micro-organisms deplete the level of available organics in the sludge, they enter an endogenous respiration phase in which energy is procured by self-digesting organics from surrounding cell protoplasm. During this phase, cellular compounds are oxidized to CO2, water, and NH3. Moreover, due to the long SRTs required for aerobic digestion, the NH3 produced by oxidation of both intracellular and extracellular (i.e. EPS) compounds is oxidized by nitrifying bacteria, producing NO3- (Droste, 1997). This nitrification process can cause a decrease in alkalinity and a subsequent decrease in pH in the digester tank if not maintained. Decreases in pH below 5.5 may cause disturbances in the system. If DO concentrations are kept below 1 mg/L, nitrification will not occur. An alternative solution in maintaining pH above 5.5 is to alternate periods of aeration and non-aerated mixing; during non-aerated mixing, the tanks enter an anoxic stage, in which NO3- produced during the aeration phase will be converted into N2 via denitrification; denitrification also replaces around 50% of the alkalinity consumed during nitrification, thus helping to maintain the tank pH. The benefits of aerobically digesting sludge includes the minimization of odours, low BOD concentrations in supernatant of dewatered digester effluent (recycled back to WWTP), low capital costs, and relatively easy operation (Tchobanoglous et al., 2003). However, the high cost of aeration is a disadvantage to this system, and recovery of power costs cannot be offset by the production and capture of CH4-rich gas, such as in AD. 2.9. AD for sludge stabilization AD is the biodegradation of organic compounds within sludge by various micro-organisms in the absence of O2. It is commonly used to stabilize excess solids generated during wastewater treatment, and has an increased popularity due to both economic and environmental benefits. AD is commonly performed in tanks called digesters in order to maintain an anaerobic environment and prevent gas leakage. The general products of AD include stabilized organic compounds and biogas; biogas is composed of CH4, CO2, N2, and trace amounts of water vapor and sulfur compounds such as hydrogen sulfide (H2S) (Rasi et al., 2011). Within AD digesters, CH4 and CO2 gases often make up between 55 – 65% and 35 – 44% of the total biogas volume produced, respectively, although higher %  23  compositions have been described (Jönsson et al., 2003; Rasi et al., 2011). Benefits of sludge stabilization via AD include the overall reduction of sludge volume, deactivation of various pathogens (Sahlström, 2003), production of energy in the form of CH4 gas (Carrère et al., 2010), reduction of odour production from biodegradation of putrescible compounds (Smet et al., 1999), and the production of nutrient-rich fertilizers from excess sludge (Appels et al., 2008). AD also reduces the need for landfill space, and offsets the carbon footprint of waste disposal via the production of biogas and nutrient-rich agricultural fertilizers (Barber, 2009). AD also negates the need for energy-intensive methods of disposal that also contribute to greenhouse gas emissions, such as sludge incineration. Limitations of AD include high capital costs, low process efficiencies, and the requirement of a high degree of technical expertise needed to maintain full scale AD digesters (Forster-Carneiro et al., 2012). Nevertheless, the use of AD is the favoured mechanism of PS and WAS stabilization. Due to these various benefits, particularly the ability of AD to produce energy in the form of CH4 gas, AD was chosen as the sludge stabilization method for this research. Therefore, the process of AD is described more thoroughly in the following sections. 2.10.  Microbiology and biochemical pathways of AD AD of wastewater treatment sludge occurs in four phases, and, for simplicity, occurs in a stepwise fashion as follows: 1) hydrolysis,  2) acidogenesis,  3) acetogenesis, and  4) methanogenesis.  The first step of AD, hydrolysis, is the solubilization of EPS compounds into simpler materials that can be more easily degraded by microbes (Shimizu et al., 1993). During this  24  process, large polymers, such as high density lipids, polysaccharides, humic acids, lignin, cellulose, proteins, and nucleic acids are biodegraded into their respective lower molecular weight components, such as glucose, glycerol, fatty acids, and amino acids. This catalysis is carried out by extracellular enzymes (e.g. cellulases, lipases, and proteases) that are secreted from the hydrolytic bacteria present within the AD tank. The monomers produced during this step are immediately available for the acidogenic bacteria in the second step of AD. In the second step, acidogenesis, acid-forming fermentative bacteria further hydrolyze the organic monomers of the sludge, converting the sugars, amino acids, and fatty acids into volatile fatty acids, including acetic, butyric, and propionic acids, as well as alcohols, acetate, CO2 and H2. Acetate is the major product of sugar fermentation by these acidogenic bacteria. In the third step, acetogenesis, acetogenic (i.e. H2 and acetate producing) bacteria convert the products during acidogenesis (i.e. fatty acids) into acetate, H2, and CO2, the precursors of methanogenesis. This group of bacteria that performs acetogenesis include species such as Syntrobacter wolinii and Syntrophomonas wolfei (McInernay et al., 1981). The final step of AD is methanogenesis. In this step, the products of acetogenesis are converted into CH4 and CO2 by obligate anaerobic archaea, or methanogens (McInernay and Bryant, 1981). There are two groups of methanogens commonly found in AD digesters:  1) hydrogenotrophic methanogens that convert H2 and CO2 into CH4 and water CO2 + 4H2  CH4 + 2H2O 2) acetotrophic methanogens that convert acetate to CO2 and CH4 CH3COOH  CH4 + CO2 It has been reported that two-thirds (nearly 70%) of the CH4 gas is produced during AD is produced by acetotrophic methanogens in a process categorized as acetoclastic methanogenesis, and one-third of the CH4 gas produced by the hydrogenotrophic methanogens during hydrogenoclastic methanogenesis (Mackie and Bryant, 1981; Demirel and Scherer, 2008). Common genera of methanogens found within AD digesters  25  include Methanosaeta and Methanosarcina, among others (Tabatabaei et al., 2009). However, shifts between dominant methanogens have been reported due to a variety of parameter changes, such as pH and temperature changes or variations in sludge feed within AD digesters (Demirel and Scherer, 2008). A basic outline of the process of AD is shown in Figure 2.4.  Figure 2.4. Diagram showing the flow of substrate through the process of AD: hydrolysis, acidogenesis, acetogenesis, and methanogenesis (adapted from Appels et al., 2008). In regards to temperature, one of the most important governing parameters of methanogenesis, AD can be performed at both mesophilic and thermophilic temperatures. Mesophilic micro-organisms are more tolerant to changes in environmental conditions than thermophilic micro-organisms (Droste, 1997). Therefore, thermophilic digesters are less often applied in full-scale systems, although the benefits of applying thermophilic temperatures include faster rates of hydrolysis and greater pathogen destruction (Gavala et al., 2003). Additionally, AD may be performed at psychrophilic temperatures (< 20°C), and has even been performed for the startup of batch cultures and accumulation systems  26  using inocula adapted to temperatures as low as 5°C and 10°C (Zeeman et al., 1988; Wellinger and Sutter, 1988). However, the rate of organic hydrolysis is reduced as temperatures are decreased, and AD performed at low temperatures require SRTs approximately twice as long as those required for mesophilic temperatures (Kashyap et al., 2003), which translates into much larger digester volume requirements. There are several factors that reduce the efficiency of AD. Residual organics, EPS, and MLSS amassed into compact matrices within sludge flocs during secondary wastewater treatment are highly recalcitrant to AD processes (Urbain et al., 1993). These flocs are so condensed that they reduce the ability of hydrolytic bacteria in the AD digester to fully utilize substrate within the sludge, thus preventing conversion of organics to CH4-precursors and ultimately reducing the overall biogas production and solids reduction efficiency. Therefore, it is unsurprising that the rate-limiting step of AD is that of hydrolysis (Eastman and Ferguson, 1981; Rocher et al., 1999), which can cause detrimental impact to AD systems. This hydrolytic step is relatively slow as biodegradation of cellulolytic wastes (e.g. lignin) and other difficult to degrade wastes (e.g. n-paraffins) within sludges can be a time-consuming process during AD (Polprasert, 1989). Chyi and Dague (1994) have also reported that larger particle sizes require a longer time for hydrolysis. 2.11. Important performance parameters for AD operation Various operational parameters must be constantly measured in order to monitor the stability of AD digesters throughout an SRT. The digester SRT is one of the most important parameters to measure during AD, as it is the average time that sludge stays within the digester. If the digester SRT is too short, then not only will there be a decrease in overall production of CH4 gas, but at a certain point there will be a net loss of methanogens from the bacterial population within the digester. As methanogens have a relatively long doubling time, the overall growth of methanogenic bacteria would not be able to compensate for this removal as these micro-organisms are washed out of the digester with the effluent solids. It has been stated that the minimal time needed to properly perform  27  methanogenesis and to prevent bacterial washout is between 5 days (Droste, 1997) and 10 days (Apples et al., 2008). Lawrence and McCarty (1970) have reported washout of methanogenic microbes at minimum SRTs of 7.8 d, 5.9 d, and 3.2 d, at digestion temperatures of 20°C, 25°C and 35°C, respectively. Previous research using pilot-scale AD digesters have utilized SRTs anywhere from as low as 3, 5, and 9 d (Nges and Liu, 2010), up to 15, 20, 27, 40, and 75 d (de la Rubia et al., 2006). It should be noted that pilot-scale reactors run at low SRTs may result in failure due to a high organic loading rate (OLR), but are used to observe the threshold for process imbalance and drive the reactors to extend their capacity for organic degradation (Nges and Liu, 2010). For this research, the three SRTs chosen were 20 d, 14 d, and 7 d, in order to compare the efficiency of pretreatment, as well as to prevent partial or even complete washout of the methanogens. Each SRT is commonly run for between 3 – 20 times the selected SRT in order to achieve biochemical stability of the AD process as well as to maintain successful operation of the digesters; previous studies have indicated that pretreatment of sludge feed will allow for the decrease of SRT while maintaining high levels of CH4 production (Droste, 1997).  The temperatures chosen for this research corresponded to the mesophilic range of 20 – 40°C, although AD processes can also occur in thermophilic temperature ranges of 45 – 60°C. Thermophilic AD has many benefits over mesophilic AD, including a greater destruction of pathogens and a doubling of CH4 gas production rate for every increase of 10°C (Droste, 1997). However, thermophilic temperatures require a greater energy input, and are therefore less cost efficient. Moreover, higher temperatures increase the metabolism of substrate fermentation and acid production by micro-organisms within digesters. This ultimately increases the VFA production, which can result in drastic decreases in the pH; this can cause digester failure if not closely monitored.  A stable pH indicates stability of the overall AD process, and a neutral pH is often preferred when running AD systems. A pH below 6.8 will begin to inhibit methanogenic activity (Droste, 1997). AD processes are very sensitive to pH and inhibitory substances (such as  28  NH3, H2S), so it is therefore important to measure the pH of the digester effluent constantly. As the concentration of VFAs directly relates to the pH of the digester system, drastic changes in pH can indicate shifts in the biochemical processes occurring in the digester. An additional factor that can affect the pH of the digester system is the level of CO2 (usually between 30 – 35%) in the gas phase of the AD system. Due to these high concentrations of CO2, alkalinity is needed to be in the range of 3000 – 5000 mg/L to buffer the pH of the system. Natural alkalinity of the system is produced by the breakdown of protein and amino acids to produce NH3. NH3 reacts with the CO2 and H2O to form NH4(HCO3), which is a form of alkalinity that buffers the AD system (Droste, 1997). Thus AD performed at thermophilic temperatures may require addition of alkalinity due to increases of VFA or CO2 production. The organic removal efficiency (i.e. ability of digester systems to reduce the overall volume of degradable solids within sludge, therefore making sludge transport and final disposal less difficult and more cost effective) of AD is commonly measured using tCOD or VS measurements. Due to the low concentrations of soluble organic compounds found in sludge, the maximum amount of VS or tCOD that may be removed by AD is between 30 – 50% of the TS (Chiu et al., 1997). Moreover, without pretreating the sludge prior to AD, this biological degradation requires a long fermentation period, and extremely large digestion tanks (Yeow and Peng, 2012). Mehdizadeh et al. (2013) have reported that the TS and VS removal performances of semi-continuous digesters fed MW-pretreated mixed sludge at 80oC decreased as the SRT was shortened; however, relative to the control digesters, the removal values improved. At the SRT of 20 d, the digester fed MW-pretreated sludge at 80°C had similar TS, VS, and tCOD removals as the control digester. At SRTs of 10 d and 5 d, the digester fed MW-pretreated sludge had VS removals of around 30% and 125%, as well as TS removals of around 40% and 190% relative improvement over the control, respectively. Moreover, at 10 d and 5 d SRTs, the tCOD removal improved between 2 – 31% and 54 – 132% over the control, respectively. These results indicate that as SRTs are shortened, the relative improvement of solids removal generally decreases; however, the sludge removal improvements were still greater for the  29  MW-pretreated digester than the control digester. The large relative improvements of the digester fed MW-pretreated sludge at 80°C during the shorter SRTs is due to the failure of the control digesters and inability to stabilize at an OLR of 5.40 ± 0.28 g VS/L of digester/d (Mehdizadeh et al., 2013). These VS and TS removal results were also supported by reports by Eskicioglu et al. (2007a) and Toreci et al. (2009) at SRTs of 20 d and 10 d.  Dewatering of sludge is a physical unit operation that reduces the moisture content of excess or digested sludges. This process allows for the reduction of the overall volume of sludge (for better handling, transport, and final disposal purposes), reduction of costs associated with final disposal, increase of the calorific value prior to sludge incineration, lowers the requirement for bulking agents or amendments needed for sludge composting, reduces odours produced from moist putrescible solids, and to reduce leachate produced during landfilling (Droste, 1997). It is well established that both aerobic digestion and AD cause significant decreases in the dewaterability of sludges (Bruss et al., 1993; Murthy and Novak, 1999; Novak et al., 2003). Pretreatment of sludge prior to AD has shown improvements in dewatering properties of the digested sludge, compared to AD systems fed non-pretreated sludge (Yeow and Peng, 2012). Biosolids are strictly regulated throughout Canada in terms of production, storage, sale, and application. In British Columbia, Canada, biosolids must achieve either Class A or Class B distinction in order to be used as soil amendment. To achieve these distinctions, biosolids must achieve strict regulations in terms of trace heavy metal concentrations, pathogen concentrations, and vector attraction. One of the most important parameters in achieving these class distinctions is the level of pathogenic micro-organisms found within the biosolids. Although there are many thousands of various pathogenic micro-organisms found within excess sludges, a select group of organisms are monitored in order to reduce time and costs of monitoring microbial populations within complex environmental samples. Fecal coliforms are universally accepted as indicator organisms of biosolids contamination, and are traditionally defined as thermo-tolerant micro-organisms that ferment lactose at 44.5°C in media containing bile salts (WHO, 2008). Common examples of fecal coliforms include Escherichia coli (E. coli), Klebsiella oxytoca, and Klebsiella  30  pneumonia, all of which can cause dehydration and diarrhea, septicemia, and even bacterial pneumonia, which can be fatal in extreme cases. Lime stabilization and composting are common sludge stabilization methods that are able to achieve Class A biosolids or Class A compost distinction in terms of fecal coliform densities. Conventional AD is also able to achieve Class B biosolids distinction at mesophilic temperatures from treatment times trialed up to 31 d (Rojas Oropeza et al., 2001; Méndez-Contreras et al., 2009). However, a disadvantage of mesophilic AD is its inability to achieve Class A biosolids, even at very long retention times, and the need to be couple it with additional processes (such as heat drying or pasteurization) to achieve higher distinction (WEF, 2004). It has been reported that thermophilic AD can achieve greater pathogen reduction at lower SRTs, compared to mesophilic AD (Popova and Bolotina, 1963; Iranpour et al., 2006). Class A biosolids distinction has been achieved at thermophilic temperatures using single-stage AD at SRTs from between 20 – 25 d (Willis and Schafer, 2006), and at 28 d (Méndez-Contreras et al., 2009). Class A biosolids have also been achieved following an aerobic digestion step as pretreatment (WEF, 2004), or by temperature phased anaerobic digestion (TPAD) (US EPA, 2006). It has been reported that as the SRT is decreased during AD, the extent of pathogen reduction is also decreased. However, this effect is diminished at higher (i.e. thermophilic) AD temperatures (Rojas Oropeza et al., 2001; Coelho et al., 2011).  The TPAD process is a variation of high temperature digestion in which thermophilic AD is followed by mesophilic AD. By following thermophilic AD with mesophilic AD, TPAD takes advantage of using thermophilic temperatures to thoroughly inactivate pathogens without the high concentrations of VFAs in the biosolids, which often contribute to foul odours (WEF, 2004). TPAD systems generally have a combined total SRT of greater than 20 d (Willis and Schafer, 2006). 2.12. Pretreatment of sludge prior to AD  To circumvent the rate-limiting nature of the first hydrolysis step of AD, it is crucial to optimize the availability of sludge organics in order to enhance the rate and yield of AD products. Not only does this include the hydrolysis of sludge flocs and their constituent  31  organics, but also includes the destruction of micro-organisms present in the sludge.  To achieve this objective, there are proposals to first pretreat WAS prior to AD using chemical, mechanical, thermal, enzymatic, and physical pretreatments, or a combination of these methods. The main goal of sludge pretreatment is to alter chemical and physical properties of the floc structure to accelerate AD and improve dewatering of residual biosolids. Moreover, these pretreatments are also able to fracture microbial cell membranes and thus release intracellular compounds into the liquid phase, providing more substrate for acid-forming bacteria, and ultimately enhancing the AD process (Mata-Alvarez et al., 2000). Essentially, the majority of pretreatments are applied to WAS and not PS, as WAS possesses a greater fraction of EPS and microbial cells that are resistant to biodegradation (Wahidunnabi and Eskicioglu, 2014). Moreover, PS directly removed from the primary settler during wastewater treatment retains a higher degradability than that of WAS, and Ge et al. (2010) have suggested that pretreatment methods administered to sludges removed after primary treatment may be ineffective or superfluous. Although pretreatment of PS is generally deemed to be unnecessary, due to the high concentrations of biodegradable organics and high moisture content within PS, it is often mixed with WAS (pretreated or not) to increase the available substrate for hydrolytic bacteria in AD digesters. However, the age and storage temperature of PS directly relates to its ability to be digested, which translates to the amount of biogas that is able to be produced during AD; the greater the age or storage temperature of PS sludge, the poorer its biodegradability becomes over time (Bhattacharya, 1981). In summary, if the rate-limiting factor of the hydrolysis stage can be bypassed via WAS pretreatment, then this would not only result in improved degradability of sludge for AD, but would also substantially decrease the total time and costs required for digestion while simultaneously increasing biogas production and fertilizer quality (Carrère et al., 2010). Moreover, the production of biogas often offsets the cost of most pretreatments used to break apart sludge flocs and lyse microbial cells. Neyens and Baeyens (2003) have also reported that various AD pretreatment methods can result in greater dewaterability of sludge, and thus further reduce costs and handling issues.  32  2.13. Brief review of physical pretreatment methods in AD Pretreatment technologies are used predominantly to break up floc structure via dissociation of complex sludge aggregates in order to increase the surface area of sludge particles for the hydrolysis step of AD. Additionally, these pretreatments are used to lyse the cellular membranes of the huge amounts of biomass that grow within the aeration tank during initial (main stream) wastewater treatment to increase the amount of degradable substrate for hydrolytic bacteria to utilize. The extent of these effects varies for each pretreatment.  There is an increased need to analyze and evaluate the various pretreatment options in terms of mechanism, costs, and efficiency (Forster-Carneiro et al., 2012). The major classes of sludge pretreatments include physical (i.e. thermal, mechanical), chemical, and biological methods. For the purpose of this study, the major physical pretreatments will be reviewed, with a focus on conventional heating (CH), MW, and US pretreatment technologies. 2.13.1. Thermal pretreatments Heating of WAS is one of the most popular pretreatment methods for AD. Thermal pretreatments were first applied to sludge to improve solids dewatering (Haug et al., 1978). In addition to better dewatering, further benefits of thermal pretreatments include increased solids removal and biogas production, decreased pathogen concentrations and reduction in odours (Bougrier et al., 2005). There are several mechanisms of heating that may be applied to sludges, with MW and CH being the most prevalent. Thermal pretreatments with temperatures ranging from 60°C to 270°C have been applied to both bench-scale and pilot-scale AD digesters. The most common temperatures administered to sludges are in the range of 60 – 180°C (Ferrer et al., 2008). It has been reported that applying thermal pretreatments above 180°C could promote an inhibitory effect on AD processes due to the production of recalcitrant soluble organics or toxic intermediates, most especially the formation of recalcitrant melanoidins during Maillard reactions at temperatures of 170°C or higher. Moreover, temperatures greater than 200°C can release excess nitrogen  33  (ammonia) and phosphorous into the liquid fraction of treated sludge samples, which can also inhibit AD (Wilson and Novak, 2009). 2.13.2. Comparison of CH and MW pretreatments Two of the most important methods of thermal pretreatment are CH and MW irradiation. CH is the transfer of heat to a sample, usually using an electric oven. When CH is applied to a sludge sample, the heat flow initiates from the surface of the sample. The rate of heating is dependent on the temperature differential and thermal properties of the sample. It is often performed at high temperatures under pressure for a short period of time. The chemical and physical changes that are caused during CH are independent of reaction time and are heavily dependent on the temperature of the CH (Valo et al., 2004), although there are other studies disagreeing with this previous research and which have reported that both exposure time to heat source and pretreatment temperature are important (Koupaie and Eskicioglu, 2015). Although this technique has traditionally been the standard method of heating, the CH pretreatment method can require longer heating times and create undesirable temperature gradients within a sample due to non-uniform heating. A rising trend is the use of MW technology for thermal solids treatment. MW is a relatively new pretreatment that can be applied at various frequencies prior to AD. It is a more practical method of heating due to its superiority over CH; MW systems are able to heat both selectively and rapidly and also possess instantaneous on/off control, thus allowing for these systems to accelerate or suppress specific reaction rates and increase energy efficiency as compared to CH systems (Tyagi and Lo, 2013). Moreover, MW-irradiated cells have shown greater damage than cells that were heated via CH at similar temperatures (Hong et al., 2004). Yet Eskicioglu et al. (2006) compared MW and CH pretreatments in which CH resulted in greater solubilization of soluble chemical oxygen demand (sCOD) and sugars. It is suggested that this result was due to the fact that CH requires a longer heating period than MW to maintain the target temperatures within a sludge sample; moreover, it was concluded that present-day technology is unable to accurately compare CH methods to MW pretreatment. Very recent research by Koupaie and Eskicioglu (2015)  34  revealed that, at identical heating rates as well as both above and below boiling point pretreatment temperatures reached by both CH and MW pretreatment, these two methods of pretreatment achieved identical particulate organic solubilization and AD performance assessed by biogas production. These results imply that the decision making for choosing one thermal pretreatment over another should be based on the energy input requirement in order to achieve effective heating of waste sludge. 2.13.3. Process of MW pretreatment MW irradiation is a pretreatment option which has been shown to increase sludge dewaterability, inactivate pathogenic organisms, and increase the efficiency of AD, both as a lone treatment and in combination with other methods, such as in the presence of specific chemicals (Wojciechowska, 2005). As explained by Yu et al. (2010), the thermal effects of MW are indicative of the generation of heat due to the absorption of MW energy, via the liquid or organic matter within a sample, by either constant or induced polarization. The MW energy applied to the sample is converted into heat from the internal resistance of rotation. Increased generation and distribution of heat within samples caused by MW irradiation is due to the reorientation and frictional movements of water molecules caused by the varying electromagnetic field within the sludge sample. MW also allows for uniform and rapid heating of a sludge sample which may easily be controlled by the operator, and includes all electromagnetic radiation possessing frequencies between 300 MHz and 300 GHz. MW is most commonly used in table-top ovens for the re-heating of food in kitchens, and the majority of MW ovens are designed to emit MW frequencies of 2.45 GHz due to a suitable penetration depth provided at this frequency to heat samples (Mehdizadeh, 2012).  In addition to the thermal effects of MW, previous research has indicated the presence of potential athermal effects during MW pretreatments. Athermal effects are caused by the splitting of hydrogen bonds and the alteration of the hydration zone due to polarized fractions of macromolecules becoming aligned with the poles of the electromagnetic field (Dreyfuss and Chipley, 1980; Hong et al., 2006; Eskicioglu et al., 2007b). These athermal  35  effects are independent of temperature changes within the sample being pretreated. The benefits of applying MW to sludge feed prior to AD include a reduction in processing time, ease of control, greater pathogen destruction, improvement in sample solubilization, increased CH4 gas production, and a superior digested product (Jones et al., 2002; Eskicioglu et al., 2009). MW pretreatment is also able to release greater amounts of intracellular components into the liquid phase of sludge samples by its ability to cause greater degradation to microbial cell membranes; Woo et al. (2000) observed increased measurements of proteins and nucleic acids within the liquid portion of MW-pretreated sludge samples, which reportedly indicates the cellular membranes of sludge micro-organisms were damaged as MW temperatures were increased. These intracellular materials then become more available for AD processes. It has been shown that certain athermal effects may cause denaturation of molecules, possibly causing the destruction of microbial membranes and cell processes within sludges (Fleming, 1961). A study by Pino-Jelcic et al. (2006) reported that the irradiation of sludge samples resulted in lower fecal coliform and Salmonella spp. levels, greater VS destruction, and achieved greater WAS floc damage compared to CH heating at similar temperatures. A study by Sato et al. (1996) compared both CH and MW pretreatment of E. coli (K-12 IFO 3301) at 35°C, 45°C, 47°C, and 50°C, and it was found that there was a non-thermal killing effect by MW pretreatment over 45°C and that bacterial death rates could be accelerated by increasing the MW power, yet no cell destruction was possible at temperatures below 35°C and MW intensity of 35 W. The majority of past studies involving WAS pretreatment before AD have often focused on pretreatment effects on the entire sludge floc, yet there is a current shift to understand exactly what occurs when individual microbial cells are pretreated. This research may help to optimize cellular degradation based on species-specific pretreatment. Woo et al. (2000) have shown that when select species of bacteria (E. coli and Bacillus subtilis or B. subtilis) are treated with MW, although there is a significant inactivation of cells, many of the cells are not lysed. It was shown that for both bacterial species, the total number of cells within the culture after irradiation was not equal to the number of viable cells. A scanning electron  36  microscope was used to compare both treated and non-treated E. coli and B. subtilis cells, and MW-pretreated E. coli cells appeared to have cracked and wrinkled surfaces, while their non-treated counterparts possessed smooth surfaces. In contrast, neither of the B. subtilis treatment groups had any cracking on their surfaces. This research indicates that although MW irradiation caused inactivation of both bacterial species, it by no means attained this inactivation purely through lysis of cells (Woo et al., 2000). 2.13.4. Mechanical pretreatments Mechanical pretreatments may be categorized into two general categories, ultrasonic and high shear pretreatments. High shear methods can include stirred ball mills, high pressure homogenization, collision plates, and mechanical jet processes; all of these methods are performed at very high pressures in order to disintegrate sludge samples. Mechanical pretreatment causes size reduction of substrate by rupturing microbial cell walls and by increasing accessibility to these internalized substrates to hydrolytic bacteria within AD digesters. US pretreatment promotes the solubilization of particulate materials within the liquid phase of WAS. The most commonly used processes are mechanical jets and high pressure homogenizers. Mechanical jets use pressure to lyse biomass via high-speed propulsion and collision. High pressure homogenizers disrupt cell walls and cause the release of cytoplasmic elements into the WAS liquid phase via changes in pressure gradients, turbulence, shear stress, and extensional shearing (Kleinig and Middelberg, 1998; Zhang et al., 2012). Demonstration-scale applications of high pressure sludge pretreatment systems have been incorporated at several WWTPs in North America using pressure ranging from 0 – 14,500 psi (Chilliwack, BC, Canada, Joint Water Pollution Control Plant, LA County, USA, and Wastewater Reclamation Facility, Des Moines, Iowa, USA), and previous bench-scale AD digesters have been fed pretreated mixed sludge pretreated with high pressure homogenization within a range of 0 – 12, 000 psi and chemical dose of 0.009 g NaOH/g TS (Wahidunnabi and Eskicioglu, 2014). For the purpose of this study, this review will focus on US pretreatment of sludge.   37  2.13.5. Process of US pretreatment When used as a sludge pretreatment, US acts as a mechanical disruption to bacterial cell membranes and sludge flocs, and is capable of high degrees of cell destruction. Ultrasonic sound waves possess a frequency over 20 kHz. Generally, the frequencies required for physical and chemical changes within sludge samples are performed in the range of 20 – 100 kHz (Pilli et al., 2011).  US systems are generally comprised of a power source (generator), a convertor, and a disruptor horn (also called a detachable probe). At the convertor, electrical energy is transformed into mechanical energy by the excitation of piezoelectric crystals within the convertor; these crystals move in a longitudinal direction, ultimately causing a motion through the disruptor horn. This motion causes the tip to move up and down, and the distance of one movement is the amplitude of the US system. The selected horn used for US-pretreatment has a maximum amplitude, and is adjustable. The % amplitude refers to the relative distance traveled by the horn, with 100% amplitude being the maximum distance able to be achieved. This distance is specific to the size (i.e. diameter) of the probe, and is usually measured in μm. Amplitude and intensity also have a direct relationship: as the amplitude is increased, the intensity of sonication increases as well. The power required to sonicate samples is not entirely dependent on the amplitude selected, but is affected by additional factors, such as the viscosity of a sample. For example, the power required to sonicate (i.e. the electrical energy required to move the horn up and down) will be much higher for more viscous samples, such as sludge with high TS concentrations (as compared to water), due to the greater load on the disruptor horn. During US pretreatment, ultrasonic intensity diminishes axially and radially from the horn, which can create dead zones (i.e. areas with reduced cavitation and mixing, as well as diminished particle collisions) within a sample being pretreated by US. As dead zones reduce the efficiency of US-treatment, these zones may be reduced by sonicating samples in deep and narrow containers, as well as by choosing an optimal horn size for the required sludge sample volume (Santos et al., 2009). When applied to sludge samples, US sound waves dissipate a vast amount of energy while moving through the sludge medium (Chu et al., 2001), and contribute to floc  38  disintegration and cell death via two methods: 1) utilization of pressure waves and cavitation, and 2) OH●, HO2●, and H● radical production.  Cavitation occurs at low pressure frequencies, and is the production of bubbles in a medium when US waves pass through. These bubbles then implode and produce localized conditions of intense heat and temperature within the sludge medium, causing microbial death and break-up of sludge flocs. Chemical reactions involving free radicals may also transpire from this increase in temperature and pressure, and occur at higher frequencies. Although both phenomena lead to floc disintegration, normally high power (200 W), lower frequency (20 kHz) sonication is used as the most efficient method for cell lysis (Takatani et al., 1981; Carrère et al., 2010). However, several negative aspects of US implementation include potential erosion of the sonotrode and higher energy consumption at increasing US frequencies (Pérez-Elvira et al., 2006).  Previous research has described US pretreatment of sludge as an effective method for improving CH4 gas production during AD (Wang et al., 1999). This mechanism also provides additional benefits to AD, including improvements in sludge solubilization, reduction of the SRT for AD, minimal clogging issues, little to no odour generation, and relative easiness to implement at full-scale sites (Nah et al., 2000; Salsabil et al., 2009). Moreover, previous research has indicated that US-pretreatment of WAS increased biogas production, CH4 gas production, and reduced levels of volatile sulphur compounds (i.e. H2S) produced during AD. Apul and Sanin (2010) treated WAS for 15 minutes using a 24 kHz ultrasonicator to apply 0.51 W/mL of the WAS sludge. During this 15 minute period, it was found that the soluble COD (sCOD) increased from 59 mg/L to 2500 mg/L, and achieved greater efficiency in the AD process. However, no cooling was performed during this US-pretreatment, and as WAS temperatures raised to 70°C, thermal effects may have also contributed to the otherwise mechanical effects of US pretreatment. Tiehm et al. (2001) have also shown that the duration of US pretreatments is an important factor to consider when pretreating sludge. When applying US for 30 min and 150 min to sludge  39  feed, the VS removal achieved a 21.2% and 36.2% increase over the control, respectively, during AD. Bougrier et al. (2005) determined that biogas yield plateaued as additional energy was applied to sludge feeds over 7,000 kJ/kg TS. After applying 6,950 kJ/kg TS, the specific energy input to feed was drastically increased to 14,547 kJ/kg TS. This increase only produced an additional 4.9% in biogas yield over the US pretreatment with the lower (6,950 kJ/kg TS) pretreatment, even with such extremely high specific energy inputs.  It has been indicated that microbial inactivation may occur at much lower specific energy inputs than the actual energy required for cellular lysis (i.e. expulsion of intracellular matter into the liquid phase of WAS). Disintegration of sludge flocs, even with minimal cell lysis, has been shown to still improve AD efficiency. Greater VS reduction in the AD digester is also indicated to be proportional to the level of cellular disintegration via US treatment (Tiehm et al., 2001; Chu et al., 2002). There are several additional benefits to US pretreatment. Under mesophilic temperatures, it has been reported that US pretreatment of excess sludges is helpful in reducing concentrations of micro-pollutants, such as naphthalene. However, several pollutants, such as pyrene, were unaffected as there was no difference in concentration before or after US pretreatment (Benabdallah El‐Hadj et al., 2006).  2.14. Current applications of molecular assays Over the past decade, there has been considerable advancement in the application of molecular and microbiological approaches to engineered biological systems, allowing for better understanding and control of concomitant biochemical processes (Narihiro and Sekiguchi, 2007). In order to better understand system productivity, substrate utilization, and biomass stability within AD processes, it is necessary to assess the viability and activity of bacteria within these engineered biological systems. Fluorophores are chemical compounds that can re-emit light once excited with a specific light wavelength, and, due to certain useful and cell-specific characteristics when applied to microbial populations, are one of the most common molecular markers used for  40  microbial characterization (Lew et al., 2010). Generally, fluorophores are compounds which designate microbial viability or activity via the emission of light (i.e. fluorescence) caused by certain actions or properties of micro-organisms. Notably, fluorophores are used as probes that indicate specific cells and tissues in analytical methods, such as epi-fluorescence microscopy and imaging, and flow cytometry. Many fluorophores have been designed to take advantage of the enzymatic activity or membrane integrity of bacterial cells, which are generally indicative of viability (Prorot et al., 2008; Cai et al., 2014). Intensities of fluorescence emitted by cells are considered to be proportional to the viability of the entire active or viable (depending on the type of fluorophore assay used) micro-organisms within a sample. The light emitted by a fluorophore is measured within a defined wavelength that can be easily determined. These respective wavelengths are termed the excitation and emission spectra. Many fluorophores contain various excitation and emission spectra within very different wavelengths, making these chemicals useful as tracers in fluids, as substrate for enzymatic assays, probes, or indicators in medical, industrial, and environmental applications.  The ability of a fluorophore to absorb and emit light photons is proportional to the product of the fluorescence quantum yield and the extinction coefficient, which is related to the fluorescence of the fluorophore (Johnson, 1998). Fluorescence-emitting compounds are able to absorb a specific wavelength of light, which results in the enhancement of electrons to greater energy levels. As electrons fall back to their initial (i.e. lower) energy levels, light is emitted at a longer and therefore less energetic wavelength. This longer wavelength is caused by some energy being lost as heat during the movement of electrons between energy levels. The difference of wavelength between the absorption and emission spectra is called the Stokes shift. When the excitation and the emission spectra are at different wavelengths, the measurement of fluorescence can be easily detected with a multitude of analytical methods. When the Stokes shift is small, it is difficult to maximize the excitation of the fluorophore, while simultaneously isolating the fluorescence emission from scattered sources (e.g. cells) within the sample. Therefore, the ideal fluorophore will have  41  a strong absorption at an excitation wavelength that is well-separated from the emission wavelength (Kogure et al., 2008). Although this is easily done when using a single fluorophore to a sample, when applying two or more fluorescence-emitting compounds in an assay, assay optimization becomes more difficult.  Traditionally, these fluorescence-emitting indicators have been established using micro-organisms in pure culture, but there is a shift to applying these compounds to environmental samples. Assays utilizing fluorophores are commonly selected due to their ability to measure viable but non-culturable (VBNC) micro-organisms within complex environmental samples, thus by-passing the need for often problematic culture-based assays. However, the fluorescence emitted is dependent on the number of microbial cells as well as the different metabolic rates of various species within a microbial population, which can be an issue in the application of fluorophores in complex environmental samples. Due to these reasons, it has been difficult to quantitate exact numbers of micro-organisms in heterogeneous microbial populations.  Although there are many different fluorophores that can be used in duality, it is essential to select these compounds based on their suitability to the environmental sample of interest. Tawakoli et al. (2013) looked at ten different viability assays, including FDA/Sytox Red, FDA/propidium iodide (PI), BacLight® Syto 9 (S9)/PI, carboxyfluorescein diacetate (CFDA)/Sytox Red, and calcein acetomethoxy/Sytox Red, and applied these combinations to quantify oral bacteria within biofilms. The bacteria were quantified with epifluorescence microscopy. When analyzed with a transmission electron microscope, it was shown that all assay procedures had considerable effect on the structural integrity of the cells within the biofilms as compared to the controls. This indicated aggressiveness caused by the fluorophore compounds during uptake, resulting in a loss of microbial viability shortly after intercalation. Fluorophore uptake in itself may cause loss of viability during measurement, regardless of their prior structural integrity (Wojcik and Dobrucki, 2008; Tawakoli et al., 2013). Although lysed cells may appear to have been assayed with the wrong fluorophore, this rather indicates former viability present at the initial time of assaying. This may lead to confusion when measuring these types of samples  42  with a direct counting method (i.e. manual microscopy). Previous literature has discussed fluorophore detection using various forms of microscopy or flow cytometry (Biggerstaff et al. 2006; Berney et al., 2007; Prorot et al., 2008). Very few studies have incorporated the use of microplate readers, although it has been proposed that rapid and accurate readings of fluorescence produced from fluorophores used in viability assays may be performed directly by using these readers (Pascaud et al., 2009; Cai et al., 2014). However, Pascaud et al. (2009) have also reported that simply extrapolating recommended protocols for certain commercial fluorophores in combination with the use of microplate readers can result in unsuccessful assays, and therefore sample-specific optimization is essential. The reasoning behind the selection of the fluorophores used in this research is detailed in chapter three. 2.14.1. S16:SO (live:dead) assay The combination of two fluorophore compounds that absorb and emit light at different wavelengths are often used in combination. Often, certain characteristics of these fluorophores, such as binding affinity and the ability of the fluorophores to permeate the cellular membrane, are utilized to indicate viability within a microbial population. For example, the S16 and SO fluorophores used in this research were chosen based on their compatibility (see chapter three).  S16 and SO are effective fluorophores for combinatory viability assays. Although there are many different types of fluorophores to choose from, these compounds were selected based on previous literature that had successfully applied these fluorophores to complex environmental samples, in particular water and wastewater samples (Biggerstaff et al., 2006; Falcioni et al., 2008). S16 is easily able to pass through cellular membranes to intercalate nucleic acids. In this sense, S16 is not selective and will permeate through all microbial cells within a sample. SO is a cell-impermeant fluorophore that binds to nucleic acids within microbial cells possessing compromised (i.e. lysed or perforated) membranes. The SO can only enter a cell through membrane tears or holes. When S16 is combined with SO, a relative ratio of live to dead cells (live:dead) within the entire microbial  43  population can be determined, as the SO is able to displace bound S16 due to its greater binding affinity to nucleic acids. However, as free nucleic acids are a substantial component of the extracellular matrix, these compounds are more indicative of the total sample biomass rather than the individualized population of micro-organisms (Whitchurch et al., 2002). The S16 is often termed as a “live cell” fluorophore, and the SO is called a “dead cell” fluorophore within the literature. Thus, these types of assays are often referred to as “live:dead” or “live/dead” assays. As these two compounds do not share emission spectra, they can be measured independently within one sample. When using such fluorophores, it is tempting to use only a live fluorophore (i.e. S9, S16) to gauge the total microbial population present. However, the addition of counter-fluorescing compounds, such as SO or PI, are essential to displace the live fluorophore that has entered perforated or lysed cells, and to quench additional background fluorescence caused by these compounds (Pascaud et al., 2009). The use of combinations of these compounds may be termed counter-fluorescing assays. 2.14.2. FDA assay The compound FDA has been reported to be metabolized by bacteria (Chrzanowski et al., 1984), algae (Gilbert et al., 1992), fungi (Söderström, 1977), and rotifers (Moffat and Snell, 1995), and is therefore a very good indicator of overall activity of biofilms (Battin, 1997) and other complex environmental samples such as activated sludge (Fontvieille et al., 1992). It is a cell-permeant, non-fluorescent esterase substrate that is hydrolyzed into fluorescein by non-specific intracellular esterases once introduced inside a metabolically active cell. Once FDA is cleaved, the fluorescein product is then able to be excited at a certain wavelength and emit fluorescence, which can be measured.  It has been reported that FDA is passively transported into cells across the cellular membrane (Chrzanowski et al., 1984), and that it cannot passively exit the cell and accumulate within cells possessing intact membranes. However, it has also been reported that once FDA is hydrolyzed by cellular esterases, the fluorescein product is able to rapidly leak from the cell. Due to this characteristic of fluorescein, fluorescence output from an  44  FDA assay cannot be traced accurately to individual cells (Prosperi et al., 1986; Clarke et al., 2001). Although similar compounds such as sulphofluorescein and CFDA are able to bypass this issue for researchers interested in individual cell-specific activity (Bölter et al., 2002), FDA is a simple and useful assay when measuring overall population activity with a high throughput microplate reader. The basis of the FDA assay is that only metabolically active cells are capable of hydrolyzing the FDA in significant amounts. Cells that are in a dormant state, such as when damaged or when present in a nutrient-poor environment, are expected to contribute minimal fluorescence to the overall detection within the sample of interest.  2.15. Summary The stabilization of sludge is an important aspect of waste management, and may be performed in such a way as to provide viable energy and resources via AD. To better facilitate the AD process, past research has indicated that the pretreatment (disintegration) of sludge prior to digestion can result in greater AD efficiency, resulting in greater CH4 yields, solids removal, and pathogen destruction, among other benefits. These advanced AD systems have utilized a variety of chemical, thermal, and mechanical methods of sludge pretreatment, in particular US (mechanical) pretreatment (which has successfully been applied at full-scale WWTPs) and, recently, MW (thermal) pretreatment (which has not yet been applied at full-scale). However, few studies have applied molecular and microbiological techniques to better understand important biological systems, such as those found in AD processes, nor to understand the effects of pretreatment on the cellular contribution to sludge solubilization and how this translates to AD efficiency. In this research, for the first time, fluorescence-emitting microbial viability assays in combination with a high throughput microplate reader were used to successfully assess the relative impact of thermal (MW) and mechanical (US) pretreatments on microbial cells in sludge samples in terms of cell lysis and enzymatic activity. The methodology of this research is outlined in the following sections.   45  Chapter 3 | Methodology 3.1. Sludge sample collection The feed sludge used for AD in this research was provided by the municipal WWTP in Kelowna, British Columbia, Canada. The design of this facility allows for the treatment of up to 70 million litres of wastewater per day, and is configured as a modified Bardenpho process. The modified Bardenpho process is used to administer BNR to remove phosphorus and nitrogen along with carbonaceous organics from treated wastewater prior to release back into the Okanagan Lake. For this experiment, both PS and tWAS were used as feed sludge (“mixed feed”) in a % volume ratio of 33:67, respectively, for the AD digesters. This feed mixture was used in order to replicate conditions at the Kelowna WWTP. The City of Kelowna currently composts its biosolids from the Kelowna WWTP, along with wood chips and incinerator ash, and produces a commercial product called “Ogogrow”. Due to the composting facility reaching its maximum capacity in addition to the high environmental footprint of the existing process, the City of Kelowna is considering AD as an alternative sludge treatment option in the future. This study, therefore, serves as a bench-scale AD feasibility study for Kelowna WWTP’s waste sludge streams. The PS and tWAS samples were collected bi-weekly from the WWTP to maintain a fresh batch of sludge feed, and all new sludge samples were characterized accordingly. Once collected, all sludges were stored in sealed containers at 4°C to prevent degradation.  3.2. Procedure for MW pretreatment of sludge  A Microwave Lab station (Ethos EZ, 2450 MHz, 0-1200 W; maximum temperature of 300°C and maximum pressure of 35 bars) with an ATC-400-CE temperature probe and 12 pressure-sealed vessels (100 mL capacity per vessel) was used to pretreat tWAS samples (Figure 3.1). During tWAS pretreatment, 50 g of tWAS was irradiated per pressure-sealed vessel (for a total volume of 600 g sludge per run). Once reaching the target (final) temperature, samples were maintained at this temperature for 1 minute. After MW pretreatment, all tWAS samples were allowed to cool to room temperature (22 ± 3°C)  46  within the pressure-sealed vessels to minimize evaporation during sample transfer. Once cooled, MW irradiated tWAS samples were transferred to clean, dry 1 L containers, sealed, and stored at 4°C until needed.     Figure 3.1. Set-up of MW irradiation of tWAS samples. The 12 pressure-sealed containers to hold tWAS samples are contained in a holding unit (top). The Microwave Lab station (Ethos EZ, 2450 MHz, 0-1200 W; maximum temperature of 300°C and maximum pressure of 35 bars) with an ATC-400-CE temperature probe is shown, with the holding unit containing the 12 pressure-sealed containers locked in place (bottom).  47  3.3. Procedure for US pretreatment of sludge A Fisher Scientific Model 500 Ultrasonic Dismembrator (with a maximum output of 400 W and frequency of 20 kHz), in combination with a Branson Model 102 Converter (101-135-066R) for Digital Sonifiers and a ¾ inch Branson Ultrasonic Disruptor Horn (stepped, tapped; 611-005-021) was used for the US treatment of tWAS samples (Figures 3.2 and 3.3, respectively). The power delivered to the convertor is measured in watts (W) and is displayed on the US system’s screen. The tWAS samples were sonicated at a total volume of 400 mL for a specified amplitude and time period corresponding to the specific energy input range chosen.   Figure 3.2. Set-up for US pretreatment of tWAS samples. The Fisher Scientific Model 500 Ultrasonic Dismembrator (with a maximum output of 400 W and frequency of 20 kHz), in combination with a Branson Model 102 Converter (101-135-066R) for Digital Sonifiers (with disruptor horn) is also shown with a beaker containing tWAS prior to US treatment (left). The disruptor horn is inserted into a beaker containing tWAS sample, which is held in a cold water bath containing crushed ice and plastic ice packs to minimize thermal effects during sonication (right). As thermal effects have been documented to occur after reaching temperatures of 40 –  48  45°C, it is important to maintain US pretreated sludge temperatures under this range in order to have purely mechanical effects administered to the tWAS samples. Therefore, containers holding the tWAS were immersed in cold water baths containing crushed ice and ice packs, which were used to maintain the temperature of tWAS samples below 35°C during sonication. This was also necessary to prevent the bench-scale US unit from shutting off due to overheating as well as to sustain US sludge pretreatment for a desired range of duration (10 – 60 min). After US pretreatment, all sonicated tWAS samples were transferred to clean, dry 1 L containers, sealed, and stored at 4°C until needed.    Figure 3.3. The ¾ inch Branson Ultrasonic Disruptor Horn (stepped, tapped; 611-005-021) used for the selected US intensities for tWAS pretreatment. 3.4. Selection of parameters to normalize pretreatment effects on sludge In order to properly observe the different effects of the MW and US pretreatments on digester feed both prior to and during AD, the two pretreatments needed to be compared under a normalized value in such a way as to eliminate the effects of individual influences inherent to the MW and US systems. It was therefore decided that the specific energy input to the tWAS samples of both the MW and US pretreatments would be calculated in units of kJ/g TS of tWAS and used for comparison. The specific energy input to sludge is defined as the amount of energy provided to a certain amount of sludge (Braguglia et al., 2012). Moreover, the specific energy inputs were chosen to compare the efficiency of these two pretreatments in order to normalize the assessments of AD operations, as each pretreatment  49  was administered to the tWAS samples using different frequencies and possessed very different organic solubilizations. All MW and US pretreatments used for the microbial characterization were performed immediately prior to application of the fluorophore assays, for reasons discussed in section 3.5. All MW and US pretreatments of tWAS samples for digester feed preparation were completed twice a week and stored at 4°C until required for feeding.  3.5. Methodology for microbial viability characterization The S16:SO and FDA assays were applied to a range of pretreated (MW and US) tWAS samples, in addition to two control sludges (live and dead). This range of intensities for each pretreatment was selected to observe potential patterns of microbial disruption based on pretreatment conditions, intensities, time duration, and temperatures; the intensities were chosen based on the specific energy inputs administered to the tWAS samples. 3.6. Selection and preparation of tWAS pretreatment intensities and controls As past research on sludge cake from the Kelowna WWTP has shown that MW pretreatment at temperatures between 80 – 160°C have increased the recoverable CH4 production of sludge stabilized  via AD (Mehdizadeh et al., 2013), the MW range utilized by Mehdizadeh et al. (2013) was used for this research. Therefore, the Milestone Microwave Lab station was used to achieve this temperature range. However, the corresponding specific energy input for each selected temperature intensity needed to be calculated. This calculation was performed by integrating the curve of a graph depicting power used to irradiate the tWAS samples over time, which was supplied by the Milestone Microwave Lab station (refer to Appendix C, section C.2 in Mehdizadeh, 2012, for sample calculations). After integration of the energy value, the energy input specific to the tWAS sample was calculated by dividing the energy applied to the mass of the dry solids within the sample that was MW-pretreated; this also normalized the energy input values for comparisons of digester feed samples between pretreatments. Therefore, based on the work by Mehdizadeh et al. (2013) and the specific energy calculations, a total of four MW temperatures (i.e. intensities) with a heating rate of 7.5oC/min were chosen for this  50  experiment: 1) 60°C (MW60), 2) 80°C (MW80),  3) 120°C (MW120), and 4) 160°C (MW160). The specific energy inputs calculated for each MW intensity, as well as their respective solubilizations, can be seen in Table 3.1. In addition to the range of MW intensities, a range of various amplitudes and sonication times were applied to tWAS samples to determine the closest possible US intensity in order to compare US and MW pretreatment for AD. As the US system used to sonicate the samples displayed the average power applied to the sample, as well as the exact amount of time the samples were sonicated, to calculate the energy applied to the tWAS samples the following equation (3.1) was used. 𝐸 = 𝑃 × 𝑇     (3.1) Where E is the energy required to pretreat the sludge sample (J), P is the power required (W), and T is the duration of the pretreatment (seconds, s). Again, the mass of the dry solids of sonicated tWAS samples were used to normalize the energy values based on feeds. Based on the specific energy inputs to tWAS trialed for US pretreatment, a total of five US pretreatments were chosen to correspond to the specific energy range of MW pretreatments selected (as described above) for comparative purposes:  1) 10 min at 60% amplitude (US1060), 2) 20 min at 60% amplitude (US2060),  3) 22 min at 80% amplitude (US2280),  4) 50 min at 100% amplitude (US50100), and   51  5) 60 min at 100% amplitude (US60100).  It should be noted that during sonication of tWAS samples, the % amplitude refers to the relative distance traveled by the horn, with 100% amplitude being the maximum distance able to be achieved. The amplitude is directly related to the intensity of US pretreatment. All of the selected MW and US intensities were administered to tWAS samples using the same methodology used in sections 3.2 and 3.3, respectively. Table 3.1 summarizes the pretreatment conditions, specific energies and particulate COD solubilization achieved for all US (in addition to MW) pretreatment intensities.  Two control tWAS sludges were also microbially characterized in order to provide a comparative index of microbial inactivation or lysis during both fluorophore assays. The first control was a non-pretreated tWAS, and used as a live control (CL). The CL gave an indication of naturally-occurring microbial activity and cell membrane integrity within the samples prior to pretreatment. The second control tWAS was used as a dead control (CD). All CD samples were sterilized in 50 mL undiluted volumes for 30 minutes at 126 ± 5°C at a pressure of 15 psi using a steam sterilizer (Model 25X 120V All American® Portable Sterilizer). This CD control was used to provide an indication of minimal microbial activity without complete liquefaction of the sludge. Although the use of steam sterilization at high temperature and pressure as an AD pretreatment has been implemented in previous research, CD was not used as an AD feed in this research (but only as a control sample to verify/optimize the accuracy of the analytical method of the selected assays) as previous literature has deemed this method of disintegration and cell lysis to be highly energy intensive and economically inefficient for full-scale application (Salsabil et al., 2010).  As mentioned in section 3.1, the tWAS samples were collected bi-weekly from the Kelowna WWTP and therefore not all runs of the assays were performed with identical tWAS samples. However, when an assay was applied to a tWAS sample, all controls were assayed along with the chosen intensities, and all samples measured within a single assay were from the same tWAS sample. By always including both controls in the assays, along with the pretreated samples being measured, variations in the tWAS samples were offset.  52  For this experiment, it is important to note that the numerical values of the emitted fluorescence was not particularly important, but emphasis is placed on the patterns of microbial disruption indicated by the detected fluorescence output. Additionally, the assay results become more robust by this inclusion of multiple samplings of tWAS. All MW, US, and CD sludge samples were prepared within the same three hour period and stored at 4°C to minimize time-dependent regeneration or reactivation of micro-organisms prior to microbial characterization.  In this experiment, although the samples were diluted, the cells were present in their natural environment (i.e. wastewater with high organic concentrations) and therefore cell dormancy due to substrate, nutrient, or vitamin deficiency can be assumed to be negligible. Thus, increases or decreases within the overall metabolic activity and viability of the measured samples are entirely assumed to be caused by the pretreatments administered.  3.7. Selection of microbial viability compounds for microbial characterization Bioassay-grade FDA in powdered form (Sigma-Aldrich) was stored at -20°C in the dark prior to microbial characterization. A LIVE/DEAD® BacLight™ Bacterial Viability Kit (containing S9 and PI) (Life Technologies, Molecular Probes®), S16 as 1 mM in dimethyl sulfoxide (DMSO) (Life Technologies, Molecular Probes®) and SO as 5 mM in DMSO (Life Technologies, Molecular Probes®) were stored at -20°C in the dark and thawed prior to use. 3.8. Preliminary study of selected viability assays in wastewater samples In the UBC Bioreactor Technology Group Laboratory, two different live:dead assays were first applied to the Kelowna WWTP diluted sludge samples to determine the appropriate assay by another researcher. Based on past literature (Boulos et al., 1999; Berney et al., 2007; Pascaud et al., 2009) the LIVE/DEAD® BacLight™ Bacterial Viability Kit (Molecular Probes®) containing a green-fluorescing Syto® 9 (S9) and a red-fluorescing PI fluorophore supplied in a 1:1 and 1:11 mmol/L ratio was first used to assay diluted sludge samples. In this kit, PI is a cell-impermeable, intercalating fluorophore that enters  53  cells that possess perforated membranes. The S9 fluorophore is a cell-permeable, intercalating compound that is able to penetrate all cell membranes within a sample. Both compounds possess strong binding affinity to nucleic acids, their target molecule. Once bound, the S9 is displaced by the PI, which has a stronger binding affinity to nucleic acids. Upon emission, S9 fluoresces green and the PI fluoresces red. When these fluorophores are combined in a single sample, a counter-fluorescence occurs and indicates relative ratios of live (S9) to dead (PI) cells. An epifluorescence microscope (Zeiss, Germany) was used to detect the fluorescence emitted by microbial cells marked with PI and S9 within tWAS samples, and images were obtained using a Zeiss Axioimager black and white camera at the respective wavelengths of the selected fluorophores. Results from this previous testing, for this particular sludge collected from the Kelowna WWTP, were ambiguous and required additional reviewing. Work from Biggerstaff et al. (2006) has shown that PI, a fluorophore commonly used in both eukaryotic cell and bacterial pure cultures and which is used in the BacLight™ assay kit, has non-specific binding (i.e. to non-biological matter within the sample) and causes autofluorescence when applied to wastewater samples due to its relatively low increase of fluorescence quantum yield once bound to nucleic acids. Similarly to PI, SO is a cell-impermeable, intercalating agent that fluoresces under a certain wavelength of excitation once bound to its target binding site (i.e. nucleic acids). The difference in SO is that it possesses a greater increase in its fluorescence quantum yield by 500 fold once bound to nucleic acids, whereas PI has an increase of only 20 – 30 fold. This enormous difference in fluorescence intensity emitted by the two respective fluorophores is the most pronounced when non-specific binding occurs, which, to some extent, is inevitable when assaying complex environmental samples containing high concentrations of both organic and nonorganic non-biological matter. When non-specific binding does occur, SO’s greater fluorescence quantum yield allows for a much more defined fluorescence between specific and non-specific binding (Biggerstaff et al., 2006). It was then shown that different fluorophores, such as SO, could successfully replace PI when assaying complex wastewater samples during live:dead assays. This greater quantum yield is helpful for direct count measurements and gives a more accurate detection by automated measurements, such as when using microplate readers or flow cytometers. 54  Table 3.1. Conditions of microwave irradiated (MW) and ultrasonicated (US) pretreatments chosen for microbial characterization. The live (CL) and dead (CD) controls were used for assay comparisons, and therefore their respective conditions are not shown. MW80 and US1060 were the only pretreated sludges used as feed for AD.  MW60 MW80 MW120 MW160 US1060 US2060 US2280 US50100 US60100 Temperature (°C) 60 80 120 160 < 35a < 35a < 35a < 35a < 35a Frequency 2,450 MHz 2,450 MHz 2,450 MHz 2,450 MHz 20 kHz 20 kHz 20 kHz 20 kHz 20 kHz Amplitude (%) - - - - 60 60 80 100 100 Pretreatment duration (min) 5.4b 9.1b 13.4b 19.6b 10 20 22 50 60 Heating rate  (°C/min) 7.5 7.5 7.5 7.5 - - - - - Specific energy input (kJ/g TS)c 2.17 2.62 4.89 6.48 2.37 4.74 6.73 23.09 27.71 sCOD/tCOD (%)d 12.6 14.9 19.0 36.4 9.1 10.6 12.7 25.3 24.5 a Temperature did not exceed 35°C during the sonication process. b Total time required to heat tWAS to target temperature and hold at this temperature for one minute. c Actual energy delivered to sample.  d % solubilization as sCOD/tCOD of the tWAS sample after pretreatment. MW60: MW at 60°C, MW80: MW at 80°C, MW120: MW at 120°C, MW160: MW at 160°C. US1060: US for 10 min at 60% amplitude, US2060: US for 20 min at 60% amplitude, US2280: US for 22 min at 80% amplitude, US50100: US for 50 min at 100% amplitude, US60100: US for 60 min at 100% amplitude.  55  Due to these results, it was decided that for this research, the PI used previously in the UBC Bioreactor Technology Group would be replaced with the fluorophore used by Biggerstaff et al. (2006), namely SO. Additionally, during preliminary testing with new assay compounds, SO and PI were both used to assay tWAS samples and compared using a fluorescence-detecting high throughput microplate reader. The SO again had much better capability than the PI, and therefore SO was used in place of PI for this research. Additionally, a similar fluorophore (S16) was used in lieu of S9 due to availability as well as for its excitation/emission spectra. Although past research has used a plethora of members from the green-fluorescing Syto® family (i.e. Syto® 13, S9), this group of fluorophores are nearly identical, as they possess little autofluorescence and, once bound to nucleic acids, increase their respective fluorescence quantum yield by more than 1000 fold. The Syto® family is permeable to all cells within a sample. The difference within this family is that each respective fluorophore may possess a slightly different range of excitation and emission wavelengths, which can be useful in determining the best compound based on the method of fluorescence measurement, among other characteristics (Invitrogen, 2010).  The FDA compound was chosen based on past research applying FDA to complex environmental samples, such as biofilms (Battin, 1997; Peeters et al., 2008), as well as wastewater (Fontvieille et al., 1992). 3.9. Assay optimization As shown by Biggerstaff et al. (2006), it is essential to first optimize the assays when applying molecular fluorophores to complex environmental samples. Therefore, CL and CD samples were used for optimization of the assays in terms of sludge sample dilution, fluorophore concentration, and incubation time. These control sludges were used for optimization as CL and CD represent the (relative) maximum live and maximum dead cells within the tWAS, respectively. Ultra-pure Type 1 water was used to dilute the control samples by a factor of 100, 50, 20, 10, 5, 1, and 0.5. All dilutions were performed after sludge pretreatment by MW, US, or steam sterilization. Each of these dilutions was then separately mixed with various concentrations of SO and S16 (each at 1 µM, 2 µM, 5 µM,  56  10 µM, 15 µM, and 20 µM in various combinations), as well as FDA dissolved in acetone (at 1 mg/mL, 2.5 mg/mL, 5 mg/mL, 10 mg/mL, 20 mg/mL, and 25 mg/mL). FDA-assayed samples were incubated at room temperature in the absence of light, and measured after 5 min, 10 min, 20 min, 45 min, 60 min, 120 min, and 180 min. Samples assayed with the S16:SO fluorophores were incubated in the dark and measured after 5 min, 10 min, 15 min, 20 min, and 25 min.  Serial dilutions using the previously mentioned dilution factors were pipetted into microplate wells for both CL and CD. Additionally, concentrations (previously described) of SO and S16, as well as FDA, were added separately to the dilutions. These assays were then incubated at the corresponding times, and measured. In terms of selection for the S16:SO assay, the optimal concentrations were found to plateau in fluorescence output once saturation of the fluorophores within the sample occurred (data not shown). A greater dilution was chosen to offset background noise possibly caused by the sludge, as well as to minimize the amount of fluorophore required. The time of incubation was based on previous literature (Biggerstaff et al., 2006; Invitrogen, 2010). It was also observed that at greater than 15 minutes incubation, the fluorescence output of both S16 and SO appeared to diminish. After the selection of the optimal fluorescence concentrations, serial dilutions were prepared using CL and CD volumetric ratios as 0:100, 10:90, 30:70, 50:50, 70:30, and 100:0 as live:dead cell solutions to ensure suitability for pretreated tWAS samples (i.e. to ensure fluorophore concentrations could efficiently indicate live:dead ratios) (Appendix A, Figure A.1).  The optimization of FDA was selected similarly to the S16:SO stains, although the incubation time of this FDA assay was kept short. Previous literature has reported FDA assay incubation times and temperatures for complex environmental samples to be anywhere from 10 min at 20°C for activated sludge (Fontvieille et al., 1992), 30 min at ambient temperature for stream sediment biofilms (Battin, 1997), and at 3 h at 37°C for clay and silt loam soil samples (Green et al., 2006). It was observed that with a non-limiting substrate, the relative fluorescence in CL would continue to greatly increase over a period of several hours, while CD did not greatly increase similarly (Appendix A, Figure A.2),  57  thus changing the relative comparison between groups; previous studies have incorporated the use of an inhibition solution (i.e. methanol, chloroform, pure acetone) to halt the continuous hydrolyzing of FDA by cells (Fontvieille et al., 1992; Green et al., 2006; Schumacher et al., 2015). This step was not performed as only the relative pattern of fluorescence, and not quantitation, was the goal of this research. Therefore, the incubation time was shortened to 45 minutes, which was deemed to be acceptable for the purpose of this research, and to minimize time-dependent differences between pretreated groups. Moreover, at concentrations greater than 20 mg/mL, the FDA would precipitate out of solution and flocculate. Additionally, at greater volumes than a 50:50 % ratio between acetone and sample, the acetone would melt the MicroWell™ optical-bottom plates used for fluorescence detection (refer to section 3.5.5) in the microplate reader. Each assay test was performed in duplicate, and was run thrice with two different tWAS samples. In summary, it was determined that a dilution factor of 20 was optimal for this particular tWAS generated by the Kelowna WWTP for both assays used in this research. Moreover, based on the chosen dilution factor and desired fluorescence intensity, the concentration of FDA to be used for this experiment was chosen to be 10 mg/mL of FDA. Additionally, the S16:SO assay was chosen to have concentrations at a 1:1 ratio of 10 µM. The selected incubation times of the FDA and S16:SO assays were 45 min and 15 min, respectively. 3.10. Microplate reader and fluorescence measurements To measure the fluorescence emitted by the assayed tWAS a Synergy™ HT Multi-Mode Microplate Reader (BioTek®) was used to measure all samples. All samples were measured in identical 400 µL 96-well black Nunc™ MicroWell™ optical-bottom plates with polymer base (Thermo Fisher Scientific Inc.) for both of the assays (Figure 3.3). Prior to beginning a full assay, all three fluorophores were added separately to Ultra-Pure Type 1 water and then pipetted into a MicroWell™ plate, and any emitted fluorescence was measured using the microplate reader; if any significant fluorescence was detected, it was assumed that either the water or MicroWell™ plate was contaminated by protein or nucleic acids, and a new MicroWell™ plate would be used. The Ultra-Pure water and the  58  MicroWell™ plate would then be retested until background fluorescence was nullified. Moreover, unassayed diluted tWAS was measured for additional fluorescence innate to the sludge. This type of background fluorescence was subtracted from the fluorescence output of each tWAS sample measured. Fluorescence values were analyzed using arbitrary fluorescence units (AFU). The fluorescence outputs were analyzed using Gen5 2.0 All-In-One Microplate Reader Software (BioTek®). For comparative purposes, the relative fluorescence output of the tWAS samples were normalized using VS concentration (% by weight) of sludge measurements, and all sample dilutions were factored into AFU comparisons. Fluorescence of cleaved FDA product (fluorescein) was measured at excitation wavelength of 490 nm and emission wavelength of 525 nm. The S16:SO assay fluorescence was measured at emission wavelength of 518 nm and excitation wavelength of 488 nm, and emission wavelength of 570 nm and excitation wavelength of 547 nm for S16 and SO, respectively. Although epifluorescence microscopy is a useful tool to measure this assay, leakage of fluorescein from perforated cells or from enzymes within the extracellular matrix may cause ambiguous results. By using an autosampling microplate reader, fluorescence outputs can be accurately read to give a total indication of microbial metabolic activity within samples. In this study, switching to a high throughput microplate reader from an epi-fluorescence microscope also allowed for a faster and less labor intensive quantification of live:dead cell ratios for cell lysis comparison between pretreatments.  3.11. Methodology for AD digester performance comparison A mechanical (US) and a thermal (MW) pretreatment were applied to tWAS in order to observe differences in disintegration techniques and AD efficiency, as well as to determine the effects of these pretreatments on microbial disruption within the tWAS samples.  3.12. Determination of pretreatment intensity for MW-AD digester feed All pretreatments were applied solely to the tWAS portion of the mixed feeds for the AD digesters. By only administering these pretreatments to the secondary sludges containing high concentrations of slowly biodegradable EPS, the energy input for feed preparation was minimized.  59   Figure 3.4. Microplate reader used for measuring fluorescence holding a 96-well black Nunc™ MicroWell™ optical-bottom microplate (top). A close-up of the MicroWell™ microplate (bottom) used for fluorescence measurements using the microplate reader. Past research on sludge cake from the Kelowna WWTP has shown that MW irradiation at temperatures between 80 – 160°C have increased the recoverable CH4 production (Mehdizadeh et al., 2013); however, it was determined that a final target temperature of 80°C applied at a ramp rate of 7.5°C/min was the optimum pretreatment temperature in terms of net energy generation. Using this information, it was decided that the MW pretreatment intensity chosen for AD would achieve a target temperature of 80°C (MW80),  Reading platform for assays  400µL microplate “wells” for assay samples  60  and would be administered at a ramp rate of 7.5°C/min. In order to compare the MW80 pretreatment to the selected US intensity (discussed in section 3.6.2), the specific energy input to the MW80-pretreated tWAS was calculated as described in section 3.5.1. The final calculation gave a specific energy input of 2.62 kJ/g TS for the MW80 intensity chosen for MW-AD feed (refer to Table 3.1).  3.13. Determination of pretreatment intensity for US-AD digester feed The US pretreatment intensity used for US-AD feed was chosen based on the specific energy input of the MW80 intensity selected for the MW-AD feed, as described in section 3.5.1, in order to provide similar intensities by both methods. Based on the ranges trialed for US pretreatment (as described previously in section 3.5.1), the chosen US intensity for AD was selected to have a sonication time of 10 min at an amplitude of 60% (US1060), which corresponded to a specific energy input of 2.37 kJ/g TS (Table 3.1). For full-scale sludge disintegration practices, it has been reported that a range of 2.5 – 5.0 kJ/g TS are generally selected for US-AD pretreatments (Braguglia et al., 2012), which coincided with several of the lower energy inputs calculated for this study. In this study, low specific energy pretreatments were selected in order to minimize the energy input and maximize the net energy production. 3.14. Acclimatization of AD digester inoculum For this study, the mesophilic (20 – 50°C) inoculum used for AD was supplied by the Penticton WWTP in British Columbia, Canada, which operates as a full-scale mesophilic AD digester. The inoculum was acclimatized to the respective pretreatment methods via acclimatization digesters prior to running the actual digester experiment in this research. Three acclimatization digesters were run for 1) MW-pretreated feed, 2) US-pretreated feed, and 3) non-pretreated (control) feed. This acclimatization of the inocula helps to prevent digester failure caused by toxic compounds that may be generated as a result of sludge pretreatments or shock when acid and CH4-forming micro-organisms are first introduced to new (i.e. altered) sludge feeds by pretreatments. These acclimatization digesters had an identical set-up to that described in section 3.6.4. In the months before the research  61  discussed in this thesis took place, the acclimatization digesters were operated at a safe SRT of 20 days for a duration of 50 days with an OLR of 3.04 g COD/L of digester/d in order to ensure that the micro-organisms were properly adapted to the pretreated feed. These acclimatization digesters were operated at 35 ± 2°C in a temperature-controlled incubating shaker rotating at 90 rpm to simulate complete-mixed reactor conditions. After this acclimatization period, the mesophilic inoculum was placed into the respective AD digester to begin the experiment. 3.15. Set-up of bench-scale AD digesters Semi-continuous, bench-scale digesters were used as the vessels of AD processes in this experiment. These AD digesters were made with side-armed 2 L Erlenmeyer flasks that were sealed with rubber stoppers at the top mouth of the flask to provide an anaerobic environment and to prevent biogas leakage. Holes with a diameter of 0.7 cm were drilled into the rubber stopper in the mouth of the flask. Through these holes, two glass pipes with a similar diameter were inserted. The first pipe possessed a length of around 35 cm, and had a 5 – 10 cm long piece of rubber tubing attached to its top; this was used for sludge removal (i.e. for effluent sludge sampling). The second glass pipe had a shorter length of around 15 cm, and had a shallow reach into the flask. This pipe also had a 10 cm piece of rubber tubing attached, but with a nozzle piece inserted into the mouth. This second pipe set-up was used for biogas collection, as a 2 L Tedlar® bag was connected to the nozzle piece using a piece of rubber hose that was around 50 cm long. The nozzle piece allowed for the biogas bag to be easily removed during volume and gas composition measurements. Additional rubber tubing (with a length of around 5 – 10 cm) was connected to the side-arm of each Erlenmeyer flask and used for sludge feed injection. When not in use, all rubber tubing was sealed with metal clamps to maintain the anaerobic environment. The described digesters are shown in Figure 3.4. In addition to the two digesters that were fed pretreated sludge using the previously described MW80 and US1060 pretreatments, a third digester was fed non-pretreated tWAS which acted as a control (CON). Therefore, each of the three digester feeds consisted of a 33:67 % volume mixture of PS and either MW80, US1060, or CON tWAS. The characteristics of the mixed feed sludge are summarized in  62  Table 3.2. Table 3.2. Characterization of both non-pretreated control (CON), ultrasonicated (US1060), and microwave irradiated (MW80) mixed sludge feeds for AD digesters.  CON US1060 MW80 pH 5.6 (0.1; 6)a 5.6 (0.1; 6) 5.6 (0.1; 6) Alkalinity (mg/L CaCO3) 765.5 (110.9; 6) 872.2 (191.2; 6) 771.3 (57.8; 6) tCOD (mg/L) 53512 (3156; 20) 52853 (3102; 20) 55012 (3979; 20) sCOD (mg/L) 4348 (528; 16) 5138 (500; 16) 7115 (313; 16) % COD solubilization (over control) - 20.2 60.6 TS (% by wt.) 4.2 (0.18; 20) 4.2 (0.14; 20) 4.3 (0.14; 20) VS (% by wt.) 3.5 (0.18; 20) 3.5 (0.15; 20) 3.6 (0.16; 20) Ammonia (mg N/L) 448.6 (181.8; 6) 501.7 (190.7; 6) 432.5 (189.8; 6) Total VFA (mg/L) 1639.1 (277.8; 12) 1760.6 (190.7; 12) 1736.8 (250.7; 12) a Arithmetic mean of replicates (standard deviation; number of replicates) COD: Chemical Oxygen Demand, tCOD: Total COD, sCOD: Soluble COD, TS: Total Solids, VS: Volatile Solids, Total VFA (Volatile Fatty Acids): Summation of acetic, propionic, and butyric acids  3.16. Start-up and operation of AD digesters Following the acclimatization period, the inocula were divided into one of three respective AD digesters: 1) MW-pretreated fed, 2) US-pretreated fed, and 3) non-pretreated (control) fed digester (Figure 3.5). Each digester possessed an active volume of 750 mL; the active volume is the liquid portion of the digester in which AD occurs. The active volume was maintained at 750 mL for the entire experimental period, and was identical for each SRT. The remaining volume of the digester was used as headspace for biogas accumulation. At the beginning of the experiment, each digestion system maintained a conservative SRT of 20 d to prevent organic overloading of the digesters. This 20 d SRT was held for a period  63  of 81 days, with an OLR of 2.81 ± 0.08 g COD/L of digester/d. Following this period, the SRT of each digester was decreased to 14 d and held for a period of 66 days with an OLR of 3.63 ± 0.11 g COD/L of digester/d. Lastly, a final shortened SRT of 7 d was held for 63 days with an OLR of 7.54 ± 0.11 g COD/L of digester/d to assess the stability of pretreated and non-pretreated digesters under organic overloading conditions. Each time a new SRT was introduced, all digesters stabilized within the first days of the change. Figure 3.6 shows the experimental set-up of this procedure, including sampling, feed preparation, and flow of feed and effluents through the digesters. 3.17. Analytical measurements of biogas volume and composition All biogas was collected within the headspace and the 2 L Tedlar® bags of each of the AD digesters. Both the volume of the biogas produced per day and the % composition of CH4, CO2, N2, and O2 in biogas were measured. Data were averaged and reported along with standard deviation. Biogas volume measurements were performed daily. This was not only to determine the total biogas and CH4 gas produced per day, but to also release pressure on the 2 L Tedlar® bags used to collect the biogas. A manometer was used to determine the total volume of the biogas produced daily at ambient temperature and pressure. The measured values were later converted to standard temperature and pressure (0°C; 100 kPa) to normalize the values over the respective SRTs. The calibration curve used for the manometer is shown in Appendix B, Figure B.1. The composition of biogas in terms of CH4, CO2, N2, and O2 in the headspace of the digesters was monitored bi-weekly using the method introduced by van Huyssteen (1967). An Agilent 7820A gas chromatograph (GC) with a packed column (Agilent G3591-8003/80002) and thermal conductivity detector (with an oven, inlet and outlet temperature of 70°C, 100°C, and 150°C, respectively) was used to perform these measurements. Helium was used as the carrier gas, with a flow rate of 25 mL/min. Once determined, the % CH4 composition was multiplied by the biogas volume produced daily in order to determine the average CH4 production produced per day throughout each SRT. All data were averaged and reported with standard deviation.  64   Figure 3.5. The three (US1060, MW80, and CON) AD digesters used in this experiment and their set-up.  Figure 3.6. Experimental outline of AD experiment for microwave irradiated tWAS feed at 80°C (MW80), ultrasonicated tWAS feed at 60% amplitude for 10 minutes (US1060), and non-pretreated (CON) tWAS feed. All feeds were mixed with primary sludge (PS) prior to being fed to the respective digester acclimatized to the designated pretreatment intensities. Also shown are the measurements performed with the biogas and biosolids (effluents) produced during the AD process over the 20 d, 14 d, and 7 d SRTs.  65  3.18. Measurements of TS and VS To measure the TS and VS of both the digester feeds and effluents, the standard methods established by APHA (2005) were used, corresponding to sections 2540 B and 2540 A. Samples were analyzed twice a week in duplicate for all three SRTs. In this procedure, clean, dry evaporating dishes were first burned at 550 ± 1oC to remove excess organics from the dishes. Once cooled, the dishes were then weighed to determine their original mass. The tWAS samples were then poured into the dishes (between 10 – 15 g of tWAS sample per dish), weighed, and the combined dish and sample weight were recorded. These dishes were then dried uncovered for 1 hour at 98°C, followed by an additional drying period of around 8 – 12 hours at 104 ± 1°C in a sealed oven with a fumigation pipe. The purpose of this drying time was to remove any water within the sample, without causing sample boiling and gasification of the water. Once the water was completely removed, the dishes containing the dry samples were again weighed. By subtracting the original mass of the dishes from the dry sample mass (including the dish mass), the TS of the tWAS sample was calculated. After weighing the dried samples, the dishes were then placed in a muffle furnace and burned at 550 ± 1°C for a minimum of 1 hour. The purpose of this step was to burn and thus remove all VS in the sample, leaving behind the fixed (inorganic) solids. Once cooled, these completely burned dishes were again weighed and recorded. By subtracting the mass of the burned (at 550°C) dishes from the mass of the evaporated (at 104°C) dishes measured previously, the total VS of the tWAS samples could be calculated. The equations (3.2) and (3.3) below were used to determine TS and VS, respectively, in terms of g/kg of the total sample tested. 𝑇𝑆 =(𝐷105 − 𝐷𝑒) × 1000𝐷𝑠 − 𝐷𝑒                               (3.2) and  𝑉𝑆 =(𝐷105 − 𝐷550) × 1000𝐷𝑠 − 𝐷𝑒      (3.3)  66  Where TS is the total solids of the sample (g/kg), VS is the volatile solids of the sample (g/kg), De is the original weight of the empty dry dish (g), Ds is the original weight of the dish plus the total mass of the added tWAS sample (g), D105 is the mass of Ds after evaporation at 105°C (g), and D550 is the mass of Ds after burning at 550°C (g). 3.19. Fecal coliform detection In this study, the level of fecal coliform contamination within the sludge effluents and feeds was measured using a membrane filtration technique to determine the number of colony forming units (CFU) per g dry solids. CFU refers to the number of viable bacterial cells that are able to replicate and produce distinguishable colonies once introduced to a specific growth medium. The membrane filtration technique is a highly sensitive process that can detect up to 1 CFU per 1000 mL of water (Cabral, 2010). However, it is limited in that it cannot detect VBNC pathogenic micro-organisms (George et al., 2001). In British Columbia, biosolids achieving Class B distinction must contain less than 2,000,000 CFU/ g dry solids, and as such, have stricter limitations in applicability. To achieve the highest level of biosolids distinction, Class A, biosolids must have 1,000 CFU/ g dry solids or less. Moreover, in order to achieve the vector attraction criteria of either Class A or Class B distinction, biosolids must reach a VS reduction of 38% during the stabilization (i.e. during AD or composting) of the sludge (BC OMRR, 2008). The fecal coliforms of both the digester feeds and effluents were quantified by lactose utilization identification using a membrane filtration method, according to the Standard Methods 9222D membrane filter procedure (APHA, 2005). Samples were analyzed during the last two weeks of each SRT in triplicate for all three digesters. For this procedure, the samples were first diluted with Ultra-Pure Type 1 water with a dilution factor of 100. After dilution, the samples were gently inverted to mix and a specific volume of the diluted samples were filtered through 0.45 µm membrane filters assisted by a vacuum filtration apparatus; additional Ultra-Pure Type 1 water was used to rinse the apparatus to ensure no sample accumulated on the filtering cone during sample transference. Once the samples were filtered, the membrane filters were placed on petri plates containing mFC solid  67  medium (mFC Nutrient Pad Sets, Sartorius, Germany), sealed with plastic wrap, and incubated at 44.5°C for 20 – 24 h. Fecal coliforms were identified based on the appearance of a dark blue colouration within colonies after the incubation period. These blue colonies were manually counted and recorded. Using the % TS measured for each effluent and feed, the total fecal coliform concentration in colony forming units (CFU) per g dry weight as TS was calculated. All dilutions made for the membrane filtration method were included in these calculations. In order to determine the exact volume of the diluted samples needed to ensure a countable plate (i.e. to prevent a “smear” of coliform colonies on the plate media), a trial and error procedure was completed using both the feed and the CON effluent, prior to measuring all of the samples. A range of filtered volumes (0.5 mL, 1 mL, 2 mL, 2.5 mL, 3 mL, and 5 mL) were used for this process for each SRT. It should be noted that data collected during the 20 d SRT were removed due to unreliability caused by the initial method developed for the highly concentrated sludge samples. All results were averaged and reported with standard deviation. 3.20. Analytical measurements for tCOD and sCOD In this study, the tCOD and sCOD concentrations were measured once a week using section 5250 D of the Standard Methods described by APHA (2005). The COD digestion solutions used for all measurements included dried K2Cr2O7 dissolved in Ultra-Pure Type 1 water (20.4 g/L), as well as a mixture of 98% concentrated H2SO4 (334 mL) and HgSO4 (34 g). These two solutions were added separately to each sample during COD digestion.  For tCOD, feed and effluent samples were first diluted and 2.5 mL was pipetted into test tubes prior to adding the COD digestion solution described above. The sCOD was measured by first centrifuging the digester feed and effluent samples at 10,000 rpm for 20 minutes. This supernatant was then filtered through membrane filters with 0.45 µm pore sizes, diluted, and 2.5 mL of each sample was pipetted into test tubes prior to adding the COD digestion solution.   68  After adding the diluted samples, 3.5 mL of K2Cr2O7 solution was added to each of the test tubes, followed by 1.5 mL of the H2SO4 and HgSO4 mixture. After the addition of the COD digestion solutions, the samples were digested at 150°C for 3 hours. All digested samples were measured for absorbency at 600 nm using a Thermo Scientific™ Genesys 10S UV-VIS spectrophotometer. Potassium hydrogen phthalate, or KHP, has a theoretical COD of 1.176 mg COD/L and was therefore prepared in a range of concentrations to be used as a standard. This range was used to determine the relative COD per sample in mg/L. The COD standard curve used for this experiment is shown in Appendix B, Figure B.2, and corresponds to 100 – 700 mg/L. The measurement of tCOD in the digester influent and effluent samples is indicative of the organic removal efficiency of the AD system. For this study, the % tCOD removal was calculated by taking the difference of the tCOD of the digester effluent from the tCOD of the influent feed over the influent feed tCOD concentrations.  3.21. Measurements of VFA concentrations  The VFA concentrations in digester feed and effluent samples were measured once a week to determine the overall stability of the digesters throughout each SRT. As the VFAs are intermediate products of CH4 production, accumulation of VFAs in the digester liquid phase indicate that the CH4 conversion route (i.e. the stepwise process of methanogenesis) is potentially disturbed and has been upset. For sample preparation, sludge samples were first centrifuged at 10,000 rpm for 30 min and then screened through a filter with 0.2 µm sized pores. The filtration was performed in order to prevent clogging within the GC column that was used to measure the liquid VFA concentrations. For the purpose of this experiment, three major VFAs were measured:  1) acetic acid,  2) propionic acid, and  69   3) butyric acid. The GC used for VFA measurement was an Agilent 7890A GC equipped with an Agilent 19091F-112, HP-FFAP polyethylene glycol TPA capillary column (length x ID: 25 m x 320 μm) and a flame ionization detector. This GC was also equipped with an autosampler. The procedure used was established by Ackman (1972), and accordingly, isobutyric acid was used as an internal standard for each of the samples measured.  3.22. Alkalinity and pH measurements The pH of all digester feeds and effluents was measured daily with a Fisher Scientific pH probe. Alkalinity measurements were performed once a week using a diluted acid titration according to Method 2320B (APHA, 2005) following sample centrifugation at 8000 rpm for 20 minutes. All samples were measured at room temperature (20 – 22oC).  3.23. Ammonia measurements Ammonia concentrations of feeds and effluents were measured once a week using pH titration via a Fisher Scientific pH probe at 20 – 22oC. Ammonia concentrations were measured according to 4500D (APHA, 2005), following sample centrifugation at 8000 rpm for 20 min. The calibration curve used to analyze the ammonia data is shown in Appendix B. 3.24. Measurement of sludge dewaterability The dewaterability of the digester effluent and feed samples was measured during the last two weeks of each SRT in triplicate for all three digesters using a Fann Instrument Company Model 440 (Texas, USA) capillary suction timer. Sludge samples (4 mL) were injected via syringe into a metal cylinder placed on chromatography paper; the liquid of the sample then drains from the sample and is absorbed into the paper via capillary suction. The time required for the liquid to move a specific distance (as measured by the capillary suction timer) is defined as the capillary suction time (CST) in seconds. The CST indicates how fast a sample releases its water, and therefore represents the dewatering characteristics  70  of the measured sludge samples at room temperature. The procedure used for the CST measurements were performed according to section 2710 G of Standard Methods (APHA, 2005). 3.25. Net energy production The net energy produced by each of the digesters was determined by taking the difference of the energy input (i.e. the energy required by pretreatment administration, digester heating conditions) and the energy output (i.e. CH4 gas production). It is assumed that the potential energy provided by the biogas is wholly due to the CH4 component, and that the CH4 gas possesses an energy content of 37 kJ/L (Droste, 1997). The energy inputs needed to pretreat the MW80 and US1060 feeds were previously calculated (refer to Table 3.1), however, the heating requirement of the mesophilic digesters must be taken into consideration. The specific heat of the sludge feed was assumed to be similar to specific heat of water (4.2 kJ/kg °C) as sludge samples contain around 96% water by w/w. The heat transfer is related to the change in temperature as shown in equation (3.4) below.  𝑄H = 𝑚 × 𝐶 × (𝑇2 −  𝑇1),     (3.4) Where QH (kJ/g TS) is the energy required to heat the sample, m (g/d) is the mass flow rate of the feed, C (kJ/kg°C) is the specific heat of the sludge feed, and T1 (°C) and T2 (°C) are the digestion and sludge feed temperatures, respectively. The digestion temperature was 35°C (T1) and the temperature of the sludge feed was 20°C (T2) for the control (non-pretreated) digester. For the MW-pretreated digester, it was assumed that pretreated tWAS at 80oC would cool to the mesophilic digestion temperature of 35oC upon being mixed with PS at room temperature, before being fed to the digester. Therefore, MW-pretreated feed sludge did not require further heating (T1 = T2 = 35°C). For the US-pretreated sludge, the temperature of the tWAS was increased to (but not exceeding) 35oC during US, so it was determined from a temperature balance that upon being mixed with PS, the overall temperature of the mixed sludge stream became 27°C (T1 = 35°C, T2 = 27°C). Once calculated, the QH for each digester was added to the energy required for pretreatment, if  71  used. The calculated input energies for the CON, US1060, and MW80 digesters are shown in Table 3.3.  Table 3.3. The energy needed to heat the digester sludges to mesophilic (35°C) temperatures (QH) was calculated using equation (3.4), and added to the energy required to pretreat the tWAS samples in order to determine the total input energy administered to the sludge added per digester.  Calculated QH (kJ/g TSadded) Pretreatment energy input to tWAS (kJ/g TSadded) Total input energy (kJ/g TSadded) Total input energy (kJ/g VSadded) CON 1.5 0 1.50 1.80 MW80 0 2.62 2.62 3.49 US1060 0.8 2.37 3.17 4.12 Once the total input energy was calculated (as kJ/g VSadded) for each digester, the average daily CH4 gas produced per SRT was multiplied by the energy content of CH4 gas (37 kJ/L), thus giving the average output energy produced by the digesters. The difference between this output energy and the total input energy (Table 3.3) was the net energy produced for the AD systems. 3.26. Statistical analysis for microbial characterization The microbial characterization data from both the S16:SO and FDA viability assays were analyzed with an Analysis of Variance (ANOVA) test, with α = 0.05 representing 95% confidence limits. Due to unequal sample sizes between the tWAS samples of each assay, post-hoc analyses of all microbial viability assay ANOVA results were performed using the Tukey-Kramer multiple comparison method; equation (C.1) was used to calculate the critical range value required by the Tukey-Kramer method (refer to Appendix C for details of the Tukey-Kramer method used in this research). Readings were considered significant when P ≤ 0.05. The values of the significant results of the microbial characterization are shown in Appendix D (Tables D.1 – D.4). Additionally, a correlation analysis was performed to determine if results of the S16:SO and FDA were directly related (Appendix E).   72  Chapter 4 | Results and Discussion 4.1. Microbial characterization using S16:SO assay Microbial characterization using the S16:SO fluorophores can be seen in Figure 4.1. An ANOVA test was run for the S16:SO assay results, as shown in Appendix D, Table D.1, where the p value = 1.5 x 10-34, which was less than the chosen alpha value (0.05). This indicated that not all samples had similar levels of cell death caused by the pretreatments. In other words, pretreatment method (i.e. MW vs. US) was a statistically significant factor for cell viability. As hypothesized, CL had the highest statistically significant of live cells compared to the numbers of dead cells within the sludge and CD had the lowest (P < 0.05). All pretreated tWAS samples had significantly lower live cell values than CL, indicating that both US and MW pretreatments (at the selected intensities) are successful in the killing, and most likely lysis, of microbial cells within the sludge samples. At MW intensities greater than MW60, there was no significant difference between CD, MW80, MW120, and MW160. This indicates that these MW intensities have an equal level of cellular disruption as that of steam sterilization, even though steam sterilization of the samples was conducted for a longer period of time (i.e. 30 minutes) compared to all the MW pretreatments. Yet the energy input required for steam sterilization can be expected to be many times greater than that required for MW pretreatment. Although the specific energy input was not calculated for this experiment as CD was only used as a comparative control during the assays, previous literature has calculated the specific energy input of steam sterilization via autoclave at 121°C under 1 bar for 15 minutes to be around 665 kJ/g TS for waste activated sludge containing 14.26 g/kg TS (Salsabil et al., 2010). These results show that at temperatures of 80°C and higher, MW pretreatment causes a similar level of cellular death as the universally accepted method of microbial sterilization, albeit at shorter exposure times and at much lower specific energy inputs. It is also interesting to note that there was no significant difference among the three highest MW intensities (80°C,  73  120°C, and 160°C) on the level of cell death caused within the tWAS samples. This may indicate that after around 80°C, irradiation of sludge does not further contribute significantly to the level of cellular death, and may just result in wasted energy. These results can help explain why MW80 was observed to be the optimum pretreatment condition yielding maximum net energy for similar waste sludge during a previous study (Mehdizadeh et al., 2013).  Figure 4.1. Ratios of live to dead cells (S16:SO) within sludge samples at various MW and US pretreatment intensities, CL, and CD (bar graph) based on level of fluorescence emitted (as arbitrary fluorescence units, AFU). Specific energy inputs of pretreatment intensities administered to tWAS samples are shown as kJ/g TS of sludge (scatter graph). Specific energy for CL is zero and was not considered for CD. CD: Dead Control (number of data points or n = 8), CL: Live Control (n = 10), MW60: MW at 60°C (n = 4), MW80: MW at 80°C (n = 8), MW120: MW at 120°C (n = 6), MW160: MW at 160°C (n = 4), US1060: US for 10 min at 60% amplitude (n = 6), US2060: US for 20 min at 60% amplitude (n = 4), US2280: US for 22 min at 80% amplitude (n = 4), US50100: US for 50 min at 100% amplitude (n = 4), US60100: US for 60 min at 100% amplitude (n = 4). There was a statistically significant difference between all four tWAS samples pretreated with MW and the five US methods, in that the MW intensities caused several times greater cellular destruction than US. Interestingly, although the MW60 pretreatment had the lowest specific energy input (2.17 kJ/g TS), it had a 50% greater level of cellular death than the US pretreatment with the second highest specific energy input (23.09 kJ/g TS) for US50100 (P < 0.05). The MW60 and US60100 were not significantly different in fluorescence output  74  (P > 0.05). This discrepancy may have been caused by the expulsion and subsequent liquefaction of the target binding sites (i.e. nucleic acids) of the intercalating fluorophores in the liquid phase of the tWAS samples at the highest US pretreatment intensity, which also corresponded to the greatest specific energy administered to the tWAS (27.71 kJ/g TS). If nucleic acids were displaced into the liquid fraction of the tWAS samples during the US60100 exposure time (1 hour), then they will have been directly affected by US treatment (i.e. production of cavitation bubbles) for a substantial period of time, which may have caused the nucleic acids to liquefy at such high temperatures and shear forces. The two pretreatment intensities that were used for the AD experiment (MW80 and US1060) have similar energy inputs (2.62 kJ/g TS and 2.37 kJ/g TS, respectively), yet US1060 had an 89.9% increase in S16:SO ratio than MW80, indicating that less cells were killed by US1060. This indicates that, in terms of the assumptions of this study, thermal effects, rather than mechanical effects, appear to be the most effective in cell destruction and floc disintegration within the tWAS. However, it is ambiguous as to whether or not these effects caused by MW are purely thermal. It is most likely a combination of both thermal and controversial athermal effects (Hong et al., 2004; Eskicioglu et al., 2007b). Chu et al. (2002) have reported that subsequent sludge floc deagglomeration and microbial lysis equates to the specific energy applied. However, this may only be valid for intensities within a single pretreatment, and not when comparing different pretreatment methods, as the results of this study indicate that there are additional effects that must be considered.  In terms of the S16:SO (live:dead) ratios, there were no statistically significant differences among US1060, US2060, and US2280, and neither was the difference between the US50100 and US60100 significant (P > 0.05). However, there was a significant difference between these two separate US groups in terms of cell death quantified by the SO fluorescence intensity only. Zhang et al. (2007) have reported that US pretreatment at 25 kHz with a power density of 0.5 W/mL not only disrupted the sludge flocs but lysed the microbial cells within the sludge samples. The lysed cells released large amounts of proteins and nucleic acids into the liquid fraction of the sludge samples, which was thought to occur around 10 – 30 min of US pretreatment.   75  4.2. Microbial characterization using FDA assay The results from the FDA assay related to enzymatic activity of the microbial cells can be seen in Figure 4.2. An ANOVA test was run for these results, as shown in Appendix D, Table D.3, where the p value = 2.7 x 10-34, which was less than the chosen alpha value (0.05). This again indicated that several of the samples possessed significantly different levels of cell death caused by the pretreatments.  Figure 4.2. FDA assay results as a function of specific energy input of pretreatment. Relative fluorescence (as arbitrary fluorescence units, AFU) emitted by metabolically active cells within sludge samples at various MW and US pretreatment intensities, CD, and CL (bar graph). Specific energy inputs of pretreatment intensities administered to the sludge samples are shown as kJ/g TS of sludge (scatter graph). Specific energy for CL is zero and was not considered for CD. CD: Dead Control (n = 7), CL: Live Control (number of  data points or n = 6), MW60: MW at 60°C (n = 6), MW80: MW at 80°C (n = 6), MW120: MW at 120°C (n = 6), MW160: MW at 160°C (n = 6), US1060: US for 10 min at 60% amplitude (n = 9), US2060: US for 20 min at 60% amplitude (n = 6), US2280: US for 22 min at 80% amplitude (n = 6), US50100: US for 50 min at 100% amplitude (n = 6), US60100: US for 60 min at 100% amplitude (n = 6). As hypothesized, the relative metabolic activity detected by the FDA assay strongly correlated (r = 0.97, p = 0.05, Appendix E), with the virtual levels of both live and dead micro-organisms within the tWAS samples. Overall, the FDA results support the results of the S16:SO assay (Figure 4.1). As seen in the S16:SO assay, the CL and CD possessed  76  the highest and lowest metabolic activities for the FDA assay, respectively (P < 0.05). The results of the FDA assay closely follow the pattern of decreased cell activity for CL, CD, and all MW and US pretreatments, excluding the US50100 and US60100 intensities. The US50100 was not significantly different in fluorescence output than the US60100 (P > 0.05). Moreover, there were no statistically significant differences in the metabolic activities between any of the MW-pretreated tWAS samples and CD, nor were the differences between US1060, US2060, and US2280 deemed to be significantly different.  Again there were significant differences between the US- and MW-pretreated groups in terms of the relative fluorescence emitted by metabolically active micro-organisms. Previous research has reported that there may be certain chemical reactions occurring during US pretreatment of sludge. These chemical reactions may cause inhibition of metabolic activity within sludge micro-organisms, without destroying floc structures (Zhang et al., 2007). Generally, the FDA and the membrane integrity assays used in this study were successfully applied to the tWAS sludges collected from the Kelowna WWTP. For both assays, it was found that the lowest specific energy inputs of MW still had many times greater inactivity and microbial death than the highest US specific energy input. Interestingly, although US2280 and MW60 had nearly identical sCOD/tCOD solubilization (12.6% and 12.7%, respectively in Table 3.1), the US2280 pretreatment achieved this level of solubilization by applying more than 3 fold greater specific energy input than MW60; additionally, both the S16:SO and FDA assays showed 70.1% and 70.0% greater  viability and metabolic activity, respectively, for MW60. This outcome is supported by previous research conducted by Salsabil et al. (2009), in which a microbial fluorophore was applied to US-pretreated WAS at various input energies (36 kJ/g TS, 31.5 kJ/g TS, and 108 kJ/g TS), and measured using flow cytometry. It was reported that the bulk of the WAS solubilization by US pretreatment was not caused by the expulsion of intracellular materials from microbial cells, but was caused by floc disintegration.   77  4.3. Production of CH4 in AD digesters The specific daily biogas production from each digester over each respective SRT is shown in Figure 4.3. As mentioned in the methodology section, the OLRs were 2.81 ± 0.08 g COD/L of digester/d, 3.63 ± 0.11 g COD/L of digester/d, and 7.54 ± 0.11 g COD/L of digester/d at corresponding SRTs of 20, 14 and 7 days, respectively. As expected, the daily specific biogas productions increased as SRT was reduced due to higher OLRs applied to all three digesters. Also, as it was clear from Figure 4.3, due to the highest OLR of 7.54 ± 0.11 g COD/L of digester/d at SRT of 7 d, the highest amount of fluctuations in daily biogas data were observed.    Figure 4.3. Specific daily biogas production (mL/d/L of digester) for microwave irradiated at 80°C (MW80), ultrasonicated for 10 minutes at 60% amplitude (US1060), and non-pretreated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs obtained at steady state. The average daily specific CH4 gas production from control and pretreated AD digesters utilizing Kelowna mixed sludge over the three SRTs can be seen in Figure 4.4. As hypothesized, as the SRT was reduced, the CH4 produced daily by the digesters increased due to increased OLRs. The CH4 gas production of both the US1060 and MW80 digesters did not appear to improve over the CON digester during the safest (longest) SRT of 20 d. Moreover, the difference in CH4 production obtained between the US1060 and MW80  78  digesters also appeared to have no change. At this SRT all of the digesters were given adequate time for hydrolysis and methanogenesis, and therefore any increase in biogas production caused by sludge pretreatment was overshadowed. The organic substrates (i.e. VFAs) in the CON digester were sufficiently converted to biogas during this longer period of time.   Figure 4.4. Average daily specific CH4 production for microwave irradiated at 80°C (MW80), ultrasonicated for 10 minutes at 60% amplitude (US1060), and non-pretreated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent  average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of samples (n) = 64 for MW80, n = 62 US1060, and n = 65 for CON. At 14 d SRT, n = 50 for MW80, n = 51 US1060, and n = 49 for CON. At 7 d SRT, n = 50 for MW80, n = 48 US1060, and n = 51 for CON. Throughout the entire experiment, the MW80 digester possessed the highest overall CH4 production per day during the 20 d SRT at 256.8 L CH4/kg VSfed. At the 14 d SRT, the MW80 digester had a higher CH4 production (248.2 L CH4/kg VSfed) over the CON digester (220.7 L CH4/kg VSfed) and the US1060 digester (230.4 L CH4/kg VSfed), showing an increase of 11.1% and 7.2%, respectively. This greater CH4 production seen in the digesters fed pretreated sludge indicates that at shorter SRTs, the AD digester fed non-pretreated sludge requires longer periods of time for micro-organisms to complete  79  methanogenesis. The CON digester is thus challenged to produce greater rates of CH4 gas due to limitations caused by utilizing sludge possessing a more complex molecular structure (Wahidunnabi and Eskicioglu, 2014). This is highlighted by comparing the CH4 gas production rate between the 20 d and 14 d SRTs. Therefore, it can be concluded that by disintegration of the sludge floc structure and expulsion of intracellular materials into the liquid phase by MW and US pretreatments, the total time requirement of AD is decreased and a significantly higher production of biogas can occur. Again, at the shortest SRT of 7 days under the highest OLR of 7.54 ± 0.11 g COD/L of digester/d, the specific CH4 gas production of the MW80 digester (222.6 L CH4/kg VSfed) appeared to be greater than the CON digester (211.2 L CH4/kg VSfed), as well as the US1060 digester (216.7 L CH4/kg VSfed); additionally, there was an incremental increase in CH4 production seen in the US1060 digester over the CON digester. This shows that at much shorter SRTs (i.e. 7 d) that are close to approaching organic overloading of the digesters, the MW pretreatment is able to produce more readily biodegradable substrate for the methanogenic bacteria to convert into CH4 gas than the US pretreatment. Although the MW80 digester had a greater CH4 production at the 14 d SRT than the 7 d SRT, these results show that irradiation is able to significantly lower the retention time. This can allow for a decreased digester volume requirement at the full-scale, in comparison to non-pretreated and US-pretreated sludges. As described previously, this may be due to both a greater floc disintegration and a greater ability to destroy the cellular membranes of micro-organisms within the sludge via deeper electromagnetic wave penetration, ultimately releasing the intracellular substrates into the liquid fraction of the tWAS to become more available for the AD process.  4.4. VS and TS removals in AD digesters The TS and VS removal efficiencies were measured for all digesters. There is a relationship between CH4 production and VS concentration reduction across the digesters, as organic (volatile) compounds are converted to CH4 during AD. Both the TS and VS % removals are shown in Figures 4.5 and 4.6, respectively. It can be seen that the removal efficiency decreased as the length of SRT decreased for all of the digesters for both TS and VS.   80   Figure 4.5. Total solids (TS) % removal for microwave irradiated at 80°C (MW80), ultrasonicated for 10 minutes at 60% amplitude (US1060), and non-pretreated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent  average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of data points (n) = 76 for MW80, US1060, and CON. At 14 d SRT, n = 67 for MW80, US1060, and CON. At 7 d SRT, n = 59 for MW80, US1060, and CON.  Figure 4.6. Volatile solids (VS) % removal for microwave irradiated at 80°C (MW80), ultrasonicated for 10 minutes at 60% amplitude (US1060), and non-pretreated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent  average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of  data points (n) = 76 for MW80, US1060, and CON. At 14 d SRT, n = 67 for MW80, US1060, and CON. At 7 d SRT, n = 59 for MW80, US1060, and CON.  81  The highest overall TS % and VS % removals were achieved by the MW80 digester at 44.0% and 53.5%, respectively, during the 20 d SRT. An increase in TS removal over the CON digester was achieved by the US1060 digester at both the 20 d and 14 d SRTs. It has been previously reported that VS removal was achieved for US-treated sludge (at 31 kHz for 64 s) at 50.3%, and 45.8% for non-pretreated sludge in semi-continuously fed fermenters at an SRT of 22; however, there was no significant increase in biogas production over the control even with a higher VS removal (Tiehm et al., 1997). 4.5. Removal of tCOD in AD digesters In addition to VS measurements, tCOD removals are also indicative of the organic removal efficiency of AD (Droste, 1997). The tCOD % removals for MW80, US1060, and CON digesters are shown in Figure 4.7.   Figure 4.7. The % organics removal as depicted by total chemical oxygen demand (tCOD) of the AD digester effluents for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent average and error bars represent the standard deviations of data collected during each SRT over the steady state.  At 20 d SRT, number of data points (n) = 18 for MW80, US1060, and CON. At 14 d SRT, n = 14 for MW80, US1060, and CON. At 7 d SRT, n = 10 for MW80, US1060, and CON.  82  As the SRT was gradually reduced, each digester had a reduction in its solids removal, which is due to the increased OLR. During the 20 d SRT, the MW80 digester had the greatest tCOD removal, with a 7.7% and a 7.3% increase over the CON and US1060 digesters, respectively. However, the greatest relative improvement was achieved by the MW80 digester during the 14 d SRT, with a 9.4% increase in tCOD removal over CON. The US1060 digester did not achieve a noticeable increase in its tCOD removal over the CON during the 20 d nor 14 d SRT; surprisingly, at the 7 d SRT, the US1060 possessed a worse tCOD removal than both the CON and MW80 digester. Previous research has reported that increased intensities of US pretreatment can enhance the disintegration of organic solids and increase the sCOD with more refractory characteristics in the liquid fraction of sludge samples (Wang et al., 2005; Benabdallah El‐Hadj et al., 2006), which may contribute to poorer tCOD % removals at shorter SRTs. 4.6. Alkalinity and pH The alkalinity levels and pH of the digesters over the respective SRTs are shown in Figures 4.8 and 4.9, respectively. Throughout all SRTs, the alkalinity and pH are related in that the digester alkalinity buffered drastic changes in pH potentially caused by VFA accumulation and digester failure. At the 20 d SRT, both the pH and alkalinity were at the overall greatest levels, due to the lower OLR and the subsequent conversion of VFA to biogas. The MW80 digester possessed a marginally greater pH and alkalinity (at 7.34 and 4787.5 mg/L as CaCO3, respectively) which may indicate a more efficient methanogenesis process following pretreatment at the 20 d SRT. However, upon lowering the SRT to 14 d and later to 7 d, both alkalinity and pH decreased for all three digesters. This is likely caused by an accumulation of VFA within the digesters linked to higher OLRs, bringing the alkalinity and pH to lower levels (Kim and Lee, 2012).   83   Figure 4.8. Alkalinity (mg/L as CaCO3) of the AD digester effluents for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of data points (n) = 6 for MW80, US1060, and CON. At 14 d SRT, n = 4 for MW80, US1060, and CON. At 7 d SRT, n = 4 for MW80, US1060, and CON.  Figure 4.9. The pH of the AD digester effluents for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of data points (n) = 35 for MW80, US1060, and CON. At 14 d SRT, n = 15 for MW80, US1060, and CON. At 7 d SRT, n = 11 for MW80, US1060, and CON.  84  4.7. Ammonia concentrations The ammonia concentrations measured for the digester effluents are shown in Figure 4.10. At all SRTs, each digester possessed ammonia concentrations below 1750 mg/L.   Figure 4.10. The ammonia concentrations measured for the AD digester effluents for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs achieved at steady state. Data represent average and error bars represent the standard deviations of data collected during each SRT over the steady state. At 20 d SRT, number of data points (n) = 8 for MW80, US1060, and CON. At 14 d SRT, n = 6 for MW80, US1060, and CON. At 7 d SRT, n = 4 for MW80, US1060, and CON. Moen et al. (2000) has reported that anaerobic digesters maintaining an ammonium concentration below 1900 – 2400 mg/L do not show inhibitory effects, and digester failure is unlikely to occur due to ammonia accumulation. Ammonia toxicity occurs during an increase of amino acids in the liquid phase by protein degradation. At the 20 d SRT, the MW80 digester had the greatest amount of ammonia in the digester effluent, at an increase of 7.4% over the CON. This indicates that MW pretreatment may have caused an increase in proteinaceous material within the feed sludge prior to AD, providing more substrate for fermentative micro-organisms. Woo et al. (2000) has shown increased concentrations of proteins and nucleic acids within the liquid phase of pure cultures irradiated with MW as temperatures were increased at nonlethal levels from 20 – 80°C. There are increases of  85  MW80 and US1060 over the CON digester effluent at 21.8% and 5.6% at the 14 d SRT, and 18.4% and 2.4% at the 7 d SRT, respectively. These results also indicate that the two pretreatments used in this research resulted in breakdown of proteinaceous materials that may have in part originated from lysed microbial cells. 4.8. Dewaterability of AD digester effluents The extent of water released by the digestate (digester effluent) sludge samples was measured by CST, and indicated the effects of the pretreatment on digested sludge dewaterability (Figure 4.11).   Figure 4.11. Dewaterability of digester effluents for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control (CON) digesters over the 20 d, 14 d, and 7 d SRTs, as seconds per total solids (TS % by weight) achieved at steady state during the last two weeks of measurements. At 20 d SRT, number of data points (n) = 11 for MW80, US1060, and CON. At 14 d SRT, n = 9 Data represent  average and error bars represent the standard deviations of data collected.for MW80, US1060, and CON. At 7 d SRT, n = 9 for MW80, US1060, and CON. Normalization of the sludge samples was performed by dividing the CST data by the respective sample’s % TS concentration by weight (w/w) (APHA, 2005). This normalization reduces the effects of any varying solids concentrations present in the sludge  86  samples. The CST values of the digester effluents were measured during each SRT. In general, the dewaterability rate of all digestates increased (i.e. CST values reduced) as SRTs were shortened from 20 d to 10 d and 7 d. This result was expected as AD as a process deteriorates the sludge dewaterability compared to feed sludge and aerobic digestion due to generation of colloidal matter and EPS. Therefore as the extent (SRT) of digestion increases, dewaterability of digestate (that needs volume reduction before final disposal such as land application) deteriorates. At the 20 d SRT, the CON digesters (790.4 s/%TS, w/w) did not appear to possess a different rate of dewaterability from the MW80 digester (788.9 s/%TS, w/w). Yet the US1060 digester had the fastest rate of dewaterability (the lowest CST value) at this SRT over the CON digester, showing a 14.4% improvement over CON. However, after reducing the SRT, this pattern changed. The MW80 digester had a faster dewaterability than both the US1060 and CON digesters, showing an 18.5% and 19.7% enhancement of dewaterability, respectively at the 14 d SRT. The difference in dewaterability rate between the US1060 and the CON digester appeared incremental. At the 7 d SRT, a similar pattern was seen, as the MW80 (533.1 s/%TS, w/w) had a faster dewaterability than both the CON (639.7 s/%TS, w/w) and US1060 (576.0 s/%TS, w/w) digesters, showing a 16.7% and a 7.4% enhancement, respectively. The US1060 digester was also significantly better than the CON digester.  These results are most likely due to the boiling of water molecules in sewage sludge caused by the release of frictional heat during the oscillation of water dipoles during MW pretreatment. In addition to the destruction of surrounding microbial cells within the tWAS samples, this dipole oscillation ultimately releases bound water accompanying the tWAS flocs (Wu, 2008), causing the release of water from the MW-pretreated digester effluent to be much faster than the other digesters. In regards to the MW80 digester, the better dewaterability also agrees with the greater biogas production and solids removal previously discussed. It has been reported that the poorer dewaterability of the digesters fed US-pretreated sludge could potentially be caused by the detrimental effect of higher US intensities on sludge dewaterability (Quarmby et al., 1999). Saha et al. (2011) have  87  reported that some mechanical pretreatments (i.e. high-pressure homogenization and US pretreatment) of sludge can deteriorate the dewaterability of the digester feed more than that caused by thermal pretreatments; thus, the CSTs of these mechanical pretreatments are significantly higher. Although this only explains the poor dewaterability of US1060 seen in the 14 d and 7 d SRTs, the dewaterability of US1060 during the 20 d SRT was also most likely poor, but was offset by the longer SRT duration.  4.9. Fecal coliform detection in AD digester effluents In this study, the level of fecal coliforms present within the digester effluents and control feed sludge were monitored in order to gauge the soil amendment applicability of these digested biosolids, as shown in Figure 4.12. At the 14 d and 7 d SRTs, all of the effluents achieved fecal coliform densities as required by Class B biosolids distinction, as defined by BC OMRR (2008).  Figure 4.12. Concentrations of fecal coliforms detected in effluents for microwave irradiated at 80°C (MW80), ultrasonicated for 10 minutes at 60% amplitude (US1060), and non-pretreated control (CON) digesters, as well as for feed sludge, over the 14 d and 7 d SRTs achieved at steady state. Data represent  average and error bars represent the standard deviations of data collected during the last two weeks of measurement. At 14 d SRT, number of data points (n) = 9 for MW80, US1060, and CON. At 7 d SRT, n = 6 for MW80, US1060, and CON. The limits for both Class A and Class B biosolids (BC OMRR, 2008) are displayed to show soil ammendment applicability. Fecal coliform levels were measured as colony forming units (CFU) per gram of dry sample solids.  88  Although Class B effluents are able to be used as soil amendment, there are restrictions as to how these biosolids may be applied to land. These restrictions include no grazing of animals on applied land for 30 days after application, food crops with harvested parts above ground (i.e. orchard fruit, wheat) cannot be harvested for 14 months after application of biosolids, food crops with harvested parts below ground (i.e. potatoes) cannot be harvested for 20 – 38 months after biosolids application, application of biosolids on private, enclosed farmland must be closed to personnel/workers for a minimum of 30 days, and application of biosolids in high-traffic public areas (i.e. public park, athletic fields) must be closed to the public for 1 year (WEF, 1995; Tchobanoglous et al., 2003). Over the 14 d and 7 d SRTs, the MW80 digester achieved the greatest coliform reduction at 73.4% and 69.8% fecal coliforms detected compared to CON (respectively) at these SRTs. As applied pretreatment temperature and intensity are major factors in pathogen destruction within biosolids, this decrease in fecal coliform concentrations is likely due to both thermal and athermal effects of MW pretreatment on the tWAS samples; these effects are enhanced when temperature and irradiation intensity are increased. Pino-Jelcic et al. (2006) reported that, compared to CH with an oven, significant cell destruction may be achieved in sludge MW irradiated for shorter treatment durations and at lower temperatures.  In regards to the US1060 digester, it is interesting to note that the concentration of fecal coliforms detected within the digester effluent appeared to increase over the CON digester. It can be seen that the US1060 digester had a 31.0% and 39.6% increase over the CON at the 14 d and 7 d SRT, respectively. This increase is most likely caused by the disintegration of sludge flocs, rather than cellular lysis and subsequent expulsion of intracellular components into the liquid fraction (Bougrier et al., 2005). By disintegrating the sludge flocs, this may release more substrate into the liquid fraction of the samples. Additionally, if the micro-organisms within these samples are not killed, they will be able to metabolize this supplementary substrate and replicate at a faster rate. Previous literature has indicated that as the specific energy input is increased to the sludge via US, the higher concentrations of solubilized compounds detected could not be linked to the lysis of cells within the  89  sludge; it was reasoned that the increase in sludge solubilization was in fact due to floc disintegration and EPS destruction (Yu et al., 2008). Additionally, it was hypothesized that the greater solubilization of the pretreated sludge samples were caused by a shift of extracellular organic materials from the inner layer to the outer layer of the sludge flocs, rather than microbial death (Yu et al., 2008; Salsabil et al., 2009). Finally, a similar response was observed in other mechanical pretreatment techniques in which fecal coliforms in high-pressure pretreated digestates were higher than the control (non-pretreated) digestates. The authors postulated that in the pretreated digestates, fecal coliforms were present at higher concentrations outside the EPS structure as a result of floc disintegration which promoted their coliform growth during membrane filtration/selective medium enumeration method (Wahidunnabi and Eskicioglu, 2014).  4.10. Net energy production from AD digesters The net energy produced by the digesters is shown in Figure 4.13. At the 20 d SRT, both the US1060 and the MW80 digesters had lower net energy generation values than the CON digester; the CON digester had a 27.0% and a 17.4% increase over the US1060 and the MW80 digesters, respectively. Additionally, there appeared to be an increase in net energy production by the MW80 digester over the US1060 digester as the total input energy (i.e. digester feed heating requirement) was greater for US1060 than MW80 (refer to Table 3.3). The daily CH4 gas production previously described also agrees with this data, as little differences were observed during biogas production between MW80 and US1060, which indicates that the longest SRT (20 d) allowed for complete conversion of the digester VFAs to biogas. The CON digester had a much greater energy output than both of the digesters fed pretreated sludge, as little energy input was applied to the non-pretreated CON feed and therefore had fewer energy deductions. Moreover, the increase seen between the MW80 and US1060 digesters was most likely due to the greater input pretreatment energy required by US1060 (refer to Table 3.3).  At the 14 d SRT, the MW80 digester appeared to have a higher net energy output (5.69 kJ/g VSfed) than the US1060 (4.41 kJ/g VSfed), which agreed with the CH4 produced daily, as  90  expected. The difference between the MW80 digester and the US1060 digester was also different at the 7 d SRT, as the MW80 digester possessed a 17.9% increase. This indicates that although the input energy is the same at each SRT for the respective pretreatment, the shortest SRT is not enough time to complete the AD process; however, the MW80 digester attains a greater net energy production via pretreatment over the US1060, as well due to the smaller energy input required by MW80 pretreatment. However, less CH4 gas is produced as SRT is shortened and therefore does not offset the energy-intensive pretreatments as much, as seen during the 14 d SRT.  Figure 4.13. The net energy production (kJ/g VSfed) of the AD digesters produced over the 20 d, 14 d, and 7 d SRTs for microwave irradiated at 80°C (MW80), sonicated for 10 minutes at 60% amplitude (US1060), and non-treated control (CON) digesters achieved at steady state. Data represent  average and error bars represent the standard deviations of data collected during steady state. At 20 d SRT, number of data points (n) = 64 for MW80, n = 62 US1060, and n = 65 for CON. At 14 d SRT, n = 50 for MW80, n = 51 US1060, and n = 49 for CON. At 7 d SRT, n = 50 for MW80, n = 48 US1060, and n = 51 for CON.  During the 14 d and 7 d SRTs, the CON digester again possessed the highest net energy outputs, at 6.37 kJ/g VSfed and 6.02 kJ/g VSfed, although these values were lower than the 20 d SRT value of 7.46 kJ/g VSfed. The same patterns could be seen for the two digesters fed pretreated sludge over the three SRTs. This decrease in net energy production also agrees with the previous data regarding the CH4 gas production (Figure 4.4), in that as the  91  SRT decreased, less time was available for methanogenesis to be completed. It is also important to emphasize that the US and MW systems used for this research was generalized lab-scale research equipment not specifically designed for pretreatment of wastewater sludge, therefore energy transfer efficiencies to the sample may be significantly lower than expected by custom-designed units at the full-scale. Therefore, the net energy yields obtained by custom made US and MW units may surplus the control digesters at the full-scale application, complementing other benefits of pretreatments in CH4 production, organics removal, fecal coliform destruction and improvement in digestate dewaterability.  It was also interesting to observe that although the microbial viability assays (live:dead and enzymatic activity assays) indicated (4 – 5 folds) greater cell lysis for MW-pretreated tWAS over the US-pretreated tWAS (Figures 4.1 and 4.2), the improvements observed from MW80-AD performance over US1060-AD was much less pronounced (3 – 10%) in terms of CH4 production (Figure 4.4), and organic removal efficiencies (Figures 4.5 – 4.7). The possible explanations for these results are as follows. Nielsen (2002) have reported the relative composition of the organic materials typically found in WAS flocs, and included cell biomass (10 – 20%), fibrous organic material (10 – 30%) and EPS (40 – 60%). As cell biomass is only 10 – 20% of the total organics possessing CH4 potential in AD, even a 4 – 5 fold greater cell lysis achieved by MW over US pretreatment has a diminished impact in the overall CH4 yields.  Furthermore, in this study, digesters received a mixture of pretreated secondary (tWAS) and untreated PS at a ratio of 67:33 % by volume, representing a typical full-scale scenario. Therefore, the advantage gained from enhanced cellular destruction in MW irradiated tWAS samples were further reduced in the mixed digester feed, resulting in much lower overall improvements in AD performance.      92  Chapter 5 | Conclusion 5.1. Summary In the first part of this research, a range of MW and US intensities were applied to tWAS samples from Kelowna’s WWTP, and were microbially characterized in terms of live to dead (live:dead) cell ratios and metabolic activity. Two microbial fluorophore assays were applied to these ranges, in addition to a live and a dead control, in order to observe the effects of the two different pretreatments on tWAS in terms of microbial death and inactivation. The two assays used included a live:dead counter-fluorescence assay using S16 and SO intercalating fluorophore stains, and the FDA enzymatic activity assay. The range of MW and US pretreatments was chosen based on previous research, and possessed varying specific energy inputs. Four MW pretreatments were measured: MW at 60°C (MW60), MW at 80°C (MW80), MW at 120°C (MW120), and MW at 160°C (MW160). Another five US pretreatments were also selected for comparison: US for 10 min at 60% amplitude (US1060), US for 20 min at 60% amplitude (US2060), US for 22 min at 80% amplitude (US2280), US for 50 min at 100% amplitude (US50100), and US for 60 min at 100% amplitude (US60100).  In the second part of this research, two sludge pretreatments, microwave irradiation at 80°C (MW80) and ultrasonication for 10 minutes at 60% (US1060) were applied to tWAS portions of feed sludge mixed with PS (67:33 % of feed volume, respectively), and fed to bench-scale AD digesters at mesophilic reactor temperatures (35°C). The effects of pretreating the sludge prior to AD were observed and compared to a digester fed non-pretreated sludge (CON), in terms of CH4 production, dewaterability, fecal coliform destruction, net energy production, and solids removal. The results are summarized as follows.  For the first time, microbial viability assays were successfully utilized for better understanding of cellular destruction during US and MW pretreatments of wastewater sludge (tWAS) under similar specific energy inputs.  93   MW irradiation attained higher tWAS solubilization at lower specific energy inputs than US pretreatment, and both the S16:SO and FDA assays show many times greater cell death and inactivation for MW-pretreated than US-pretreated tWAS at similar specific energy inputs. These results indicate that even at lower energy inputs, MW irradiation of sludge causes greater microbial destruction than US pretreatments, indicating that greater energy input cannot necessarily achieve greater cellular lysis.   MW60 and US2280 had nearly identical sCOD/tCOD solubilizations (12.6% and 12.7%, respectively), yet both the S16:SO and FDA assays showed several times greater cell death and inactivity in the MW60 sample. This indicates that increased solubilization of sludge by US is caused by floc disruption, rather than cell lysis.   The digester pH and alkalinity results followed a similar trend. Both digesters fed US1060 and MW80 pretreated sludge had marginally higher pH and alkalinity than the CON digester. The ammonia concentrations within the pretreated digester effluents were greater than the CON, and were likely due to the effect of the pretreatments (i.e. enhanced hydrolysis/degradation of proteins) on the sludge feed.   The MW80 and US1060 have similar energy inputs (2.62 kJ/g TS and 2.37 kJ/g TS, respectively), yet US1060 had an 89.9% and 81.9% increase in the S16:SO and FDA values, respectively, than MW80. This indicates that less cells were killed by US1060. This may also suggest that thermal effects, rather than mechanical effects, appear to be the most effective in cell destruction and floc disintegration within the tWAS. Yet it is ambiguous as to whether or not these effects caused by MW are purely thermal.   Overall, the FDA and S16:SO results support one another as both assays resulted in similar patterns of fluorescence output for the pretreated and control samples.  From AD experiments, the highest overall CH4 production was achieved by the MW80 digester (256.8 L CH4/kg VSfed). Neither of the digesters fed pretreated  94  sludge significantly improved in CH4 gas production over the CON digester during the 20 d SRT. This was not surprising as all of the digesters were given adequate time for methanogenesis to be completed. At shorter SRTs (i.e. 7 d), that are close to approaching the organic overloading of the digesters, the MW80 pretreatment is able to produce better substrate for the methanogenic bacteria to convert into CH4 gas than the US1060 pretreatment.   The highest overall TS % and VS % removals were achieved by the MW80 digester at 43.2% and 53.5%, respectively, during the 20 d SRT. The VS and TS % removal efficiency decreased as the length of SRT decreased for all of the digesters. Moreover, as the SRT was gradually reduced, each digester had a reduction in its % tCOD solids removal, which is due to the increased OLR. The MW80 digester had the greatest tCOD removal (54.7%) at the 20 d SRT.  The US1060 had the best dewaterability at the 20 d SRT (676.5 s/%TS, w/w), yet the MW80 achieved a better dewaterability rate at the 14 d (571.9 s/%TS, w/w) and 7 d (533.1 s/%TS, w/w) SRTs.   Over all SRTs measured, the MW80 digester achieved the greatest fecal coliform reduction at 73.4% and 69.8% less fecal coliforms than CON during the 14 d and 7 d SRTs, respectively. The fecal coliform concentrations in the US1060 digester increased over the CON digester at the 14 d (31.0 % increase) and 7 d (39.6 % increase) SRTs. This increase is most likely due to the disintegration of sludge flocs, rather than cellular lysis, caused by US pretreatment. Moreover, the digester effluents achieved Class B biosolids distinction, according to BC OMRR (2008).  The digester fed non-pretreated (control) feed had the highest overall net energy output (7.46 kJ/g VSfed) at the 20 d SRT, while digesters fed MW- and US-pretreated sludge had net energy outputs of 6.16 kJ/g VSfed and 5.44 kJ/g VSfed, respectively. This greater net energy output of the control digester can be offset by the additional benefits provided by MW pretreatment of sludge prior to AD.   95   Results from this research indicate that microbial lysis is important for increased substrate availability for methanogenesis. However, improvements from the AD digester fed MW-pretreated sludge (although higher compared to the AD digester fed US-pretreated and control sludge quantified by microbial assays) was diminished due to smaller contributions (10 – 20%) of sludge micro-organisms and addition of PS to the digester feed.  5.2. Overall conclusion In general, the microbial viability assay results from this research are very valuable to provide a deeper understanding for why thermal pretreatments (i.e. MW and CH) have been historically more successful than mechanical pretreatments (i.e. US, high-pressure homogenization) in enhancing biodegradation potential of sludge in digesters and improving digestate quality in terms of fecal coliforms. The following is a finalized summary of the hypotheses (refer to section 1.3) tested in this thesis: 1. Although it was found that both AD digesters fed pretreated tWAS produced a greater amount of CH4 gas as well as a greater fecal coliform destruction than the control digester, these increases were minor.  2. It was validated that the level of enzymatic activity and cellular integrity of microbial cells within the pretreated samples were much lower than those found in the non-treated control samples.  3. Although it was validated that the level of cellular destruction was greater in microbially characterized MW- and US-pretreated sludge (r = 0.97, p = 0.05), this greater cellular disruption only marginally corresponded to greater overall AD efficiency within the two digesters fed pretreated tWAS over the control.  4. MW-pretreatment indicated a better substrate for AD processes than US-pretreatment of the tWAS feed. However, it was not validated as to whether or not this was due to thermal or athermal MW effects.     96  5.3. Recommendations for future work The following are recommendations for future areas of research, in relation to the results discussed in this thesis.  Comparison of sludge pretreatment using multiple pretreatment methods, such as high pressure homogenization, CH, or MW treatment at various frequencies, as well as combinations of these, measured using the microbial viability assays used in this research.  Additionally, it would be interesting to measure the concentrations of proteins, nucleic acids, sugars, and lipids before and after sludge pretreatment, and compare it to the assay results.  Application of the microbial assays to both non-pretreated and pretreated sludges that are stored for long periods of time (i.e. prior to composting or incineration), to gauge the abilities of micro-organisms to regenerate/recover over time.        97  Bibliography Ackman, R.G. 1972. Porous polymer bead packing and formic acid vapor in the GLC of volatile free fatty acids. Journal of Chromatographic Science. 10, 560 – 565. American Public Health Association (APHA). 2005. Standard Methods for the Examination of Water and Wastewater. Washington DC, USA. Aoi, Y., Miyoshi, T., Okamoto, T., Tsuneda, S., Hirata, A., Kitayama, A., Nagamune, T. 2000. Microbial ecology of nitrifying bacteria in wastewater treatment process examined by fluorescence in situ hybridization. Journal of Bioscience and Bioengineering, 90 (3), 234 – 240. Appels, L., Baeyens, J., Degrève, J., Dewil, R. 2008. Principles and potential of the anaerobic digestion of waste-activated sludge. Progress in Energy and Combustion Science, 34, 755 – 781. Apul, O.G., Sanin, F.D. 2010. Ultrasonic pretreatment and subsequent anaerobic digestion under different operational conditions. Bioresource Technology, 101 (23), 8984 – 8992. Barber, W.P.F. 2009. Observing the effects of digestion and chemical dosing on the calorific value of sewage sludge. Proceedings of IWA Specialist Conference: Moving forward. Wastewater biosolids sustainability: Technical, managerial, and public synergy. Moncton, New Brunswick, Canada, 351 – 358. Battin, T.J. 1997. Assessment of fluoresce in diacetate hydrolysis as a measure of total esterase activity in natural stream sediment biofilms. The Science of the Total Environment, 198, 51 – 60.  Benabdallah El‐Hadj, T., Dosta, J., Marquez‐Serrano, R., Mata‐Alvarez, J. 2006. Effect of ultrasound pretreatment in mesophilic and thermophilic anaerobic digestion with emphasis on naphthalene and pyrene removal. Water Research, 41 (1), 87 – 94.  98  Berney, M., Hammes, F., Bosshard, F., Weilenmann, H.U., Egli, T. 2007. Assessment and interpretation of bacterial viability by using the LIVE/DEAD® BacLight Kit in combination with flow cytometry. Applied and Environmental Microbiology, 73 (10), 3283 – 3290.  Bhattacharya, S.N. 1981. Flow characteristics of primary and digested sewage sludge. Rheologica Acta, 20, 288 – 298. Biggerstaff, J.P., Le Puil, M., Weidow, B.L., Prater, J., Glass, K., Radosevich, M., White, D.C. 2006. New methodology for viability testing in environmental samples. Molecular and Cellular Probes, 20, 141 – 146.  Bölter, M., Bloem, J., Meiners, K., Möller, R. 2002. Enumeration and biovolume determination of microbial cells – a methodological review and recommendations for applications in ecological research. Biology and Fertility of Soils, 36, 249 – 259. Botheju, D., Bakke, R. 2011. Oxygen effects in anaerobic digestion-a review. The Open Waste Management Journal 4, 1 – 19. Bougrier, C., Carrère, H., Delgenes, J.P. 2005. Solubilization of waste-activated sludge by ultrasonic treatment. Chemical Engineering Journal, 106 (2), 163 – 169. Boulos, L., Prévost, M., Barbeau, B., Coallier, J., Desjardins, R. 1999. LIVE/DEAD BacLight: application of a new rapid staining method for direct enumeration of viable and total bacteria in drinking water. Journal of Microbiological Methods, 37, 77 – 86.  Braguglia, C.M., Gianico, A., Mininni, G. 2012. Comparison between ozone and ultrasound disintegration on sludge anaerobic digestion. Journal of Environmental Management, 95, S139 – S143.  British Columbia Organic Matter Recycling Regulation (BC OMRR). 2008. Land application guidelines for the organic matter recycling regulation and the soil  99  amendment code of practice, best management practice. BC OMRR and Soil Amendment Code of Practice (SACoP) Land Application Guidelines. Prepared for: BC Ministry of Environment. Prepared by: SYLVIS Environmental. New Westminster, British Columbia. Bruss, J.H., Christensen, J.R., Rasmussen, H. 1993. Anaerobic storage of activated sludge: effects on conditioning and dewatering performance. Water Science Technology, 28, 350 – 357. Cabral, J. P. 2010. Water microbiology. Bacterial pathogens and water. International Journal of Environmental Research and Public Health, 7 (10), 3657 – 3703. Cai, Y., Strømme, M., Welch, K. 2014. Bacteria viability assessment after photocatalytic treatment. Biotechnology. 3 Biotech, 4, 149 – 157.  Carey, R.O., Migliaccio, K.W. 2009. Contribution of wastewater treatment plant effluents to nutrient dynamics in aquatic systems: a review. Environmental Management, 44, 205 – 217.  Carrère, H., Dumas, C., Battimelli, A., Batstone, D.J., Delgenès, J.P., Steyer, J.P., Ferrer, I. 2010. Pretreatment methods to improve sludge anaerobic degradability: A review. Journal of Hazardous Materials, 183, 1 – 15.  Chiu, Y., Chang, C., Lin, J., Huang, S. 1997. Alkaline and ultrasonic pretreatment of sludge before anaerobic digestion. Water Science and Technology, 36 (11), 155 – 162.   Chrzanowski, T.H., Crony, R.D., Hubbard, J.G., Welch, R.P. 1984. Applicability of the fluorescein diacetate method of detecting active bacteria in freshwater. Microbial Ecology, 10, 179 – 185. Chu, C.P., Chang, B.V., Liao, G.S., Jean, D.S., Lee, D.J. 2001. Observations on changes in ultrasonically treated waste activated sludge. Water Research, 35 (4), 1038 – 1046.   100  Chu, C.P., Lee, D.J., Chang, B.V., You, C.S., Tay, J.H. 2002. “Weak” ultrasonic pretreatment on anaerobic digestion of flocculated activated biosolids. Water Research, 36 (11), 2681 – 2688.  Chyi, Y.T., Dague, R.R. 1994. Effects of particulate size in anaerobic acidogenesis using cellulose as a sole carbon source. Water Environment Research, 66 (5), 670 – 678. Chynoweth, D.P., Owens, J.M., Legrand, R. 2001. Renewable methane from anaerobic digestion of biomass. Renewable Energy, 22, 1 – 8.  Clarke, J.M., Gillings, M.R., Altavilla, N., Beattie, A.J. 2001. Potential problems with fluorescein diacetate assays of cell viability when testing natural products for antimicrobial activity. Journal of Microbiological Methods, 46, 261 – 267. Coelho, N.M.G. 2012. Application of microwaves and thermophilic anaerobic digestion to wastewater sludge. Ph.D. thesis, Department of Civil Engineering, University of Ottawa, Ontario, Canada. Corbitt, R.A. 1990. Standard Handbook of Environmental Engineering, Second Edition. The McGraw-Hill Companies, Inc. New York, USA. Davis, M.L., Cornwell, D.A. 2008. Introduction to Environmental Engineering, Fourth Edition. The McGraw-Hill Companies, Inc. New York, USA. de la Rubia, M.A., Perez, M., Romero, L.I., Sales, D. 2006. Effect of solids retention time (SRT) on pilot scale anaerobic thermophilic sludge digestion. Process Biochemistry, 41 (1), 79 – 86. Demirel, B., Scherer, P. 2008. The role of acetotrophic and hydrogenotrophic methanogens during anaerobic conversion of biomass to methane: A review. Reviews in Environmental Science and Biotechnology, 7, 173 – 190. Dreyfuss, M.S., Chipley, J.R. 1980. Comparison of effects of sublethal microwave irradiation and conventional heating on the metabolic activity of Staphylococcus  101  aureus. Applied and Environmental Microbiology, 39, 13 – 16.  Droste, R.L. 1997. Theory and practice of water and wastewater treatment. John Wiley & Sons, New Jersey, USA. Eastman, J.A., Ferguson, J.F. 1981. Solubilization of particulate organic carbon during the acid phase of anaerobic digestion. Water Pollution Control Federation, 53 (3), 352 – 366. Epstein, E. 1997. The science of composting. Technomic Publishing Co., Lancaster, PA. Eskicioglu, C., Kennedy, K.J., Droste, R.L. 2006. Characterization of soluble organic matter of waste activated sludge before and after thermal pretreatment. Water Research, 40, 3725 – 3736. Eskicioglu, C., Droste, R. L., Kennedy, K. J. 2007a. Performance of anaerobic waste activated sludge digesters after microwave pretreatment. Water Environment Research, 79 (11), 2265 – 2273.   Eskicioglu, C., Terzian, N., Kennedy, K.J., Droste, R.L., Hamoda, M. 2007b. Athermal microwave effects for enhancing digestibility of waste activated sludge. Water Research, 41, 2457 – 2466.  Eskicioglu, C., Kennedy, K.J., Droste, R.L. 2009. Enhanced disinfection and CH4 production from sewage sludge by microwave irradiation. Desalination, 248, 279 – 285.  Falcioni, T., Papa, S., Gasol, J.M. 2008. Evaluating the flow-cytometric nucleic acid double-staining protocol in realistic situations of planktonic bacterial death. Applied and Environmental Microbiology, 74 (6), 1767 – 1779.  Ferrer, I., Ponsá, S., Vázquez, F., Font, X. 2008. Increasing biogas production by thermal (70°C) sludge pretreatment prior to thermophilic anaerobic digestion. Biochemical Engineering Journal, 42 (2), 186 – 192.  102  Fontvieille, D.A., Outaguerouine, A., Thevenot, D.R. 1992. Fluorescein diacetate hydrolysis as a measure of microbial activity in aquatic systems: application to activated sludges. Environmental Technology, 13, 531 – 540. Fleming, J. 1961. Microwave irradiation in relation to biological systems and neural activity. In: Biological effects of microwave radiation. Plenum Press, New York, USA. Forster-Carneiro, T., Ricardo, I., Pérez, M., Schvartz, C. 2012. Anaerobic digestion – Pretreatments of substrates. Biogas Production. Edited by: Mudhoo, A. Scrivener Publishing LLC, USA, 1 – 25. Gavala, H.N., Yenal, U., Skiadas, I.V., Westermann, P. Ahring, B.K. 2003. Mesophilic and thermophilic anaerobic digestion of primary and secondary sludge. Effect of pretreatment at elevated temperature. Water Research, 37 (19), 4561 – 4572. Ge, H., Jensen, P. D., Batstone, D. J. 2010. Pretreatment mechanisms during thermophilic – mesophilic temperature phased anaerobic digestion of primary sludge. Water Research, 44 (1), 123 – 130. George, I., Crop, P., Servais, P. 2001. Use of β-D-Galactosidase and β-D-Glucuronidase activities for quantitative detection of total and faecal coliforms in wastewater. Canadian Journal of Microbiology, 47, 670 – 675.  Gilbert, F., Galiani, F., Cadiou, Y. 1992. Rapid assessment of metabolic activity in marine microalgae: application in ecotoxicological tests and evaluation of water quality. Marine Biology, 112, 119 – 205.  Green, V.S., Stott, D.E., Diack, M. 2006. Assay for fluorescein diacetate hydrolytic activity: optimization for soil samples. Soil Biology and Biochemistry, 38, 693 – 701. Hai, F.I., Li, X., Price, W.E., Nghiem, L.D. 2011. Removal of carbamazepine and  103  sulfamethoxazole by MBR under anoxic and aerobic conditions. Bioresource Technology, 102 (22), 10386 – 10390. Haug, R.T., Stuckey, D.C., Gossett, J.M., McCarty, P.L. 1978. Effect of thermal pretreatment on digestibility and dewaterability of organic sludges. Journal of the Water Pollution Control Federation, January, 73 – 85.  Hong, S.M., Park, J.K., Lee, Y.O. 2004. Mechanisms of microwave irradiation involved in the destruction of fecal coliforms from biosolids. Water Research, 38, 1615 – 1625. Hong, S.M., Park, J.K., Teeradej, N., Lee, Y.O., Cho, Y.K., Park, C.H. 2006. Pretreatment of sludge with microwaves for pathogen destruction and improved anaerobic digestion performance. Water Environment Research, 78 (1), 76 – 83.  Invitrogen. 2010. Molecular Probes® Handbook: A Guide to Fluorescent Probes and Labeling Technologies, 11th Edition. Edited by: Johnson, I. and Spence, M.T.Z. Iranpour, R., Cox, H.H.J., Oh, S., Fan, S., Kearney, R.J., Abkian, V., Haug, R.T. 2006. Thermophilic-Anaerobic Digestion to Produce Class A Biosolids: Initial Full-Scale Studies at Hyperion Treatment Plant. Water Environment Research, 78 (2), 170 – 180. Johnson, I. 1998. Fluorescent probes for living cells. Journal of Histochemistry and Cytochemistry, 30, 123 – 140. Jones, D.A., Lelyveld, T.P., Mavrofidis, S.D., Kingman, S.W., Miles, N.J. 2002. Microwave heating applications in environmental engineering – a review. Resources, Conservation, and Recycling, 34, 75 – 90. Jönsson, O., Polman, E., Jensen, J.K., Eklund, R., Schyl, H., Ivarsson, S. 2003. Sustainable gas enters the European gas distribution system. Danish Gas Technology Center. Kashyap, D.R., Dadhich, K.S., Sharma, S.K. 2003. Biomethanation under psychrophilic  104  conditions: a review. Bioresource Technology, 87 (2), 147 – 153. Khursheed, A., Kazmi, A.A. 2011. Retrospective of ecological approaches to excess sludge reduction. Water Research, 45, 4287 – 4310.  Kim, M.H., Hao, O.J. 1990. Comparison of activated sludge stabilization under aerobic or anoxic conditions. Research Journal of the Water Pollution Control Federation, 62 (2), 160 – 168. Kim, J., Park, C., Kim, T.H., Lee, M., Kim, S., Kim, S.W., Lee, J. 2003. Effects of various pretreatments for enhanced anaerobic digestion with waste activated sludge. Journal of Bioscience and Bioengineering, 95 (3), 271 – 275. Kim, D.J., Lee, J. 2012. Ultrasonic sludge disintegration for enhanced methane production in anaerobic digestion: effects of sludge hydrolysis efficiency and hydraulic retention time. Bioprocess and Biosystems Engineering, 35 (1-2), 289 – 296. Kleinig, A.R., Middelberg, A.P.J. 1998. On the mechanism of microbial cell disruption in high pressure homogenization. Chemical Engineering Science, 53, 891 – 898. Kogure, T. Kawano, H., Abe, Y., Miyawaki, A. 2008. Fluorescence imaging using a fluorescent protein with a large Stokes shift. Methods, 45, 223 – 226. Koupaie, E.H., Eskicioglu, C. 2015. Below and above boiling point comparison of microwave irradiation and conductive heating for municipal sludge digestion under identical heating/cooling profiles. Bioresource Technology, 187, 235 – 245. Kropp, J., Dichtl, N. 2001. Sludge production and characterization. Sludge into Biosolids: Processing, Disposal. Edited by: Spinosa, L., Vesilind, P.A. IWA Publishing, London, UK, 19 – 39. Lawrence, A.W., McCarty, P.L. 1970. A unified basis for biological treatment design and operation. Journal of Sanitary Engineering Division, American Society of Civil Engineers, 96 (SA3).  105  Lew, S., Lew, M., Mieszczyński, T., Szarek, J. 2010. Selected fluorescent techniques for identification of the physiological state of individual water and soil bacterial cells – review. Folia Microbiologica, 55 (2), 107 – 118.  Lovley, D.R., Goodwin, S. 1988. Hydrogen concentrations as an indicator of the predominant terminal electron-accepting reactions in aquatic sediments. Geochimica et Cosmochimica Acta, 52, 2993 – 3003. Mackie, R.I., Bryant, M.P. 1981. Metabolic activity of fatty acid-oxidizing bacteria and the contribution of acetate, propionate, butyrate, and CO2 to methanogenesis in cattle waste at 40 and 60°C. Applied and Environmental Microbiology, 41, 1363 – 1373. Mata-Alvarez, J., Macé, S., Llabrés, P. 2000. Anaerobic digestion of organic solid wastes. An overview of research achievements and perspectives. Bioresource Technology, 74, 3 – 16.  McInernay, M.J., Bryant, M.P. 1981. Basic principles of bioconversions in anaerobic digestion and methanogenesis. Biomass conversion processes for energy and fuels. Edited by: Sofer, S.S., Zaborsky, O.R. Plenum Publishing Corp., New York, USA, 277 – 296.  McInernay, M.J., Bryant, M.P., Hespell, R.B., Costerton, J.W. 1981. Syntrophomonas wolfei, gen. nov. sp. nov., an anaerobic, syntrophic, fatty acid-oxidizing bacterium. Applied and Environmental Microbiology, 41, 1029 – 1039. McLoughlin, A.J. 1994. Controlled release of immobilized cells as a strategy to regulate ecological competence of inocula. Advances in Biochemical Engineering/Biotechnology, 51, 2 – 45. Mehdizadeh, S.N. 2012. Enhancement of Kelowna’s biosolids to energy conversion with thermal pretreatments. M.A.Sc. thesis, Department of Civil Engineering, University of British Columbia, Kelowna, British Columbia, Canada.  106  Mehdizadeh, S.N., Eskicioglu, C., Bobowski, J., Johnson, T. 2013. Conductive heating and microwave hydrolysis under identical heating profiles for advanced anaerobic digestion of municipal sludge. Water Research, 47, 5040 – 5051. Méndez-Contreras, J.M., Rendón-Sagardi, J.A., Ruiz-Espinoza, J.E., Alvarado-Lassman, A., Martínez-Delgadillo, S.A. 2009. Behavior of the mesophilic and thermophilic anaerobic digestion in the stabilization of municipal wastewater sludge (part 1). Revista Mexicana de Ingeniería Química, 8 (3), 283 – 290. Moffat, B.D., Snell, T.W. 1995. Rapid toxicity assessment using an in vivo enzyme test for Brachionus plicatilis (Rotifera). Ecotoxicology and Environmental Safety, 30, 47 – 53.  Murthy, S.N., Novak, J.T. 1999. Factors affecting floc properties during aerobic digestion: implications for dewatering. Water Environment Research, 71, 197 – 202. National Research Council (NRC). 1993. In situ bioremediation - When does it work? National Academies Press, Washington D.C., USA, 224. Narihiro, T., Sekiguchi, Y. 2007. Microbial communities in anaerobic digestion processes for waste and wastewater treatment: a microbiological update. Current Opinion in Biotechnology, 18 (3), 273 – 278. Nah, I.W., Kang, Y.W., Hwang, K.Y., Song, W.K. 2000. Mechanical pretreatment of waste activated sludge for anaerobic digestion process. Water Research, 34 (8) 2362 – 2368. Neyens, E., Baeyens, J. 2003. A review of thermal sludge pre-treatment processes to improve dewaterability. Journal of Hazardous Materials, 98 (1-3), 51 – 67.  Nges, I.A., Liu, J. 2010. Effects of solid retention time on anaerobic digestion of dewatered-sewage sludge in mesophilic and thermophilic conditions. Renewable Energy, 35 (10), 2200 – 2206.  107  Nielsen, P.H. 2002. Activated sludge – the floc. Encyclopedia of Environmental Microbiology. Edited by Bitton, G. Wiley-Interscience, Hoboken, New Jersey, USA. Nielsen, P.H., Saunders, A.M., Hansen, A.A., Larsen, P., Nielsen, J.L. 2012. Microbial communities involved in enhanced biological phosphorus removal from wastewater – a model system in environmental biotechnology. Current Opinion in Biotechnology, 23, 452 – 459. Novak, J.T., Sadler, M.E., Murthy, S.N. 2003. Mechanisms of floc destruction during anaerobic and aerobic digestion and the effect on conditioning and dewatering of biosolids. Water Research, 37 (13), 3136 – 3144. Olofsson, A.-C., Zita, A., Hermansson, M. 1998. Floc stability and adhesion of green-fluorescent-protein-marked bacteria to flocs in activated sludge. Microbiology, 144, 519 – 528. Park, C., Helm, R.F. 2008. Application of metaproteomic analysis for studying extracellular polymeric substances (EPS) in activated sludge flocs and their fate in sludge digestion. Water Science and Technology, 57 (12), 2009 – 2015. Park, C., Novak, J.T., Helm, R.F., Ahn, Y.-O., Esen, A. 2008. Evaluation of the extracellular proteins in full-scale activated sludges. Water Research, 42, 3879 – 3889. Pascaud, A., Amellal, S., Soulas, M.L., Soulas, G. 2009. A fluorescence-based assay for measuring the viable cell concentration of mixed microbial communities in soil. Journal of Microbiological Methods, 76, 81 – 87.  Peeters, E., Nelis, H.J., Coenye, T. 2008. Comparison of multiple methods for quantification of microbial biofilms grown in microtiter plates. Journal of Microbiological Methods, 72 (2), 157 – 165. Pilli, S., Bhunia, P., Yan, S., LeBlanc, R.J., Tyagi, R.D. 2011. Ultrasonic pretreatment of  108  sludge: a review. Ultrasonics Sonochemistry, 18, 1 – 18. Pino-Jelcic, S.A., Hong, S.M., Park, J.K. 2006. Enhanced anaerobic biodegradability and inactivation of fecal coliforms and Salmonella spp. in wastewater sludge by using microwaves. Water Environment Research, 78 (2), 209 – 216.  Pelczar, M.J., Reid, R.D. 1958. Microbiology. The McGraw-Hill Companies, Inc. New York, USA. Pérez-Elvira, S.I., Nieto Diez, P., Fdz-Polanco, F. 2006. Sludge minimization technologies. Reviews in Environmental Science and Biotechnology, 5, 375 – 398.  Polprasert, C. 1989. Organic waste recycling. John Wiley & Sons, New Jersey, USA. Popova, N.M., Bolotina, O.T. 1963. The present state of purification of town sewage and the trend in research work in the City of Moscow. International Journal of Air and Water Pollution, 7, 145 – 158. Prorot, A., Eskicioglu, C., Droste, R., Dagot, C., Leprat, P. 2008. Assessment of physiological state of micro-organisms in activated sludge with flow cytometry: application for monitoring sludge production minimization. Journal of Industrial Microbiology and Biotechnology, 35, 1261 – 1268. Prosperi, E., Croce, A.C., Bottiroli, G., Supino, R. 1986. Flow cytometric analysis of membrane permeability properties influencing intracellular accumulation and efflux of fluorescein. Cytometry, 7, 70 – 75. Quarmby, J., Scott, J.R., Mason, A.K., Davies, G. and Parsons, S.A. 1999. The application of ultrasound as a pre-treatment for anaerobic digestion. Environmental Technology, 20 (11), 1155 – 1161. Rajeshwari, K.V., Balakrishnan, M., Kansal, A., Kusum Lata, Kishore, V.V.N. 2000. State-of-the-art of anaerobic digestion technology for industrial wastewater treatment. Renewable and Sustainable Energy Reviews, 4, 135 – 156.  109  Rasi, S., Läntelä, J., Rintala, J. 2011. Trace compounds affecting biogas energy utilization – a review. Energy Conversion and Management, 52 (12), 3369 – 3375.  Rocher, M., Goma, G., Pilas Begue, A., Louvel, L., Rols, J.L. 1999. Towards a reduction in excess sludge production in activated sludge processes: biomass physicochemical treatment and biodegradation. Applied Microbiology and Biotechnology, 51, 883 – 890. Roediger, H. 1987. Using quicklime – hygienization and solidification of dewatered sludge, Water Environment Federation. WEF Operations Forum, 18 – 21.  Rojas Oropeza, M., Cabirol, N., Ortega, S., Castro Ortiz, L.P., Noyola, A. 2001. Removal of fecal indicator organisms and parasites (fecal coliforms and helminth eggs) from municipal biologic sludge by anaerobic mesophilic and thermophilic digestion. Water Science and Technology, 44 (4), 97 – 101. Ross, M. 2013. Operator Essentials – What every operator should know about biological phosphorus removal. Waste, Environment, and Technology Magazine (July), Water Environment Federation, 48 – 50. Saha, M., Eskicioglu, C., Marin, J. 2011. Microwave, ultrasonic and chemo-mechanical pretreatments for enhancing CH4 potential of pulp mill wastewater treatment sludge. Bioresource Technology, 102 (17), 7815 – 7826. Sahlström, L. 2003. A review of survival of pathogenic bacteria in organic waste used in biogas plants. Bioresource Technology, 87, 161 – 166. Salsabil, M.R., Prorot, A., Casellas, M., Dagot, C. 2009. Pretreatment effects of activated sludge: Effect of sonication on aerobic and anaerobic digestibility. Chemical Engineering Journal, 148, 327 – 335.  Salsabil, M.R., Laurent, J., Casellas, M., Dagot, C. 2010. Techno-economic evaluation of thermal treatment, ozonation and sonication for the reduction of wastewater biomass  110  volume before aerobic or anaerobic digestion. Journal of Hazardous Materials, 174, 323 – 333. Santos, H.M., Lodeiro, C., Capelo-Martínez, J.L. 2009. The power of ultrasound. Ultrasound in Chemistry: Analytical Applications. Edited by: Capelo-Martínez, J.L. Wiley-VCH Verlag GmbH and Co. KGaA, Weinheim, Germany. Sato, S., Shibata, C., Yazu, M. 1996. Nonthermal killing effect of microwave irradiation. Biotechnology Techniques, 10 (3), 145 – 150. Schumacher, T.E., Eynard, A., Chintala, R. 2015. Rapid cost-effective analysis of microbial activity in soils using modified fluorescein diacetate method. Environmental Science and Pollution Research, 22, 4759 – 4762. Shimizu, T., Kudo, K., Nasu, Y. 1993. Anaerobic waste-activated sludge digestion – A bioconversion mechanism and kinetic model. Biotechnology and Bioengineering, 41 (11), 1082 – 1091. Siripong, S., and Rittmann, B.E. 2007. Diversity study of nitrifying bacteria in full-scale municipal wastewater treatment plants. Water Research, 41 (5), 1110 – 1120. Smet, E., Van Langenhove, H., De Bo, I. 1999. The emission of volatile compounds during the aerobic and the combined anaerobic/aerobic composting of biowaste. Atmospheric Environment, 33, 1295 – 1303. Söderström, B.E. 1977. Vital staining of fungi in pure cultures and in soil with fluorescein diacetate. Soil Biology and Biochemistry, 9, 59 – 63. Tabatabaei, M., Zakaria, M.R., Rahim, R.A., Wright, A.D.G., Shirai, Y., Abdullah, N., Sakai, K., Ikeno, S., Mori, M., Kazunori, N., Sulaiman, A., Hassan, M.A. 2009. PCR-based DGGE and FISH analysis of methanogens in an anaerobic closed digester tank for treating palm oil mill effluent. Electronic Journal of Biotechnology, 12 (3), 2 – 12.  111  Takatani, S., Takayama, S., Yamauchi., T. 1981.  A study of anaerobic digestion for sewage sludge. Mitsubishi Juko Giho (Japan), 18, 1 – 7. Tawakoli, P.N., Al-Ahmad, A., Hoth-Hannig, W., Hannig, M., Hannig, C. 2013. Comparison of different live/dead stainings for detection and quantification of adherent micro-organisms in the initial oral biofilm. Clinical Oral Investigations, 17, 841 – 850.  Tchobanoglous, G., Schroeder, E.D. 1985. Water Quality.  Addison-Weslev Publishing Company. Inc., U.S.A. Tchobanoglous, G., Burton, F.L., Stensel, H.D. 2003. Wastewater engineering: treatment and reuse, 4th Edition. The McGraw-Hill Company, Inc., New York, USA. Tiehm, A., Nickel, K., Neis, U. 1997. The use of ultrasound to accelerate the anaerobic digestion of sewage sludge. Water Science Technology, 36 (11), 121 – 128. Tiehm, A., Nickel, K., Zellhorn, M., Neis, U. 2001. Ultrasonic waste activated sludge disintegration for improving anaerobic stabilization. Water Research, 35 (8), 2003 – 2009. Toreci, I., Kennedy, K. J., Droste, R. L. 2009. Evaluation of continuous mesophilic anaerobic sludge digestion after high temperature microwave pretreatment. Water Research, 43 (5), 1273 – 1284. Turovskiy, I.S., Mathai, P.K. 2006. Wastewater sludge processing. Wiley Publishing, New York, USA. Tyagi, V.K., and Lo, S.L. 2013. Microwave irradiation: A sustainable way for sludge treatment and resource recovery. Renewable and Sustainable Energy Reviews, 18, 288 – 305. United States Environmental Protection Agency (US EPA). 1983. Process design manual for land application of municipal sludge. EPA 625/1-83-016. Municipal  112  Environmental Research Laboratory, Cincinnati, OH.  United States Environmental Protection Agency (US EPA). 2006. Biosolids technology fact sheet – Multi-stage anaerobic digestion. EPA 832-F-06-031. Office of Water.  Urbain, V., Block, J.C., Manem, J. 1993. Bioflocculation in activated sludge: An analytical approach. Water Research, 27, 829 – 83. Valo, A., Carrère, H., Delgenès, J.P. 2004. Thermal, chemical and thermo-chemical pretreatment of waste activated sludge for anaerobic digestion. Journal of Chemical Technology and Biotechnology, 79, 1197 – 1203. van Huyssteen, J.J. 1967. Gas chromatographic separation of anaerobic digester gases using porous polymer. Water Research, 1, 237 – 242. Vergara, S.E. 2012. Composting – Encyclopedia of Consumption and Waste: The Social Science of Garbage. Thousand Oaks, CA: SAGE, 138 – 141. Wahidunnabi, A.K., Eskicioglu, C. 2014. High pressure homogenization and two-phased anaerobic digestion for enhanced biogas conversion from municipal waste sludge. Water Research, 66, 430 – 446.  Wang, F., Wang, Y., Ji., M. 2005. Mechanisms and kinetics models for ultrasonic waste activated sludge disintegration. Journal of Hazardous Materials, 23, 145 – 150. Wang, Q., Kuninobu, M., Kakimoto, K., Ogawa, H.I., Kato, Y., 1999. Upgrading of anaerobic digestion of waste activated sludge by ultrasonic pretreatment. Bioresource Technology, 68, 309 – 313.  Water Environment Federation (WEF). 1995. Wastewater residuals stabilization. Manual of Practice, no. FD-9. WEF, Alexandria, VA. Water Environment Federation (WEF). 1998. Design of wastewater treatment plants. Manual of Practice, fourth edition, volume 8, no. 3. WEF, Alexandria, VA.  113  Water Environment Federation (WEF). 2004. High performance anaerobic digestion. Residuals and Biosolids Committee, Bioenergy Technology Subcommittee. WEF, Alexandria, VA. Wellinger, A., Sutter, K. 1988. Biogas production at low temperatures. Proceedings of Energy from Biomass and Wastes XII, New Orleans. Institute of Gas Technology, Chicago, IL, 21. Whitchurch, C.B., Tolker-Nielsen, T., Ragas, P.C., Mattick, J.S. 2002. Extracellular DNA required for bacterial biofilm formation. Science, 295, 1487.  Willis, J., Schafer, P. 2006. Advances in thermophilic anaerobic digestion. Water Environment Federation, Atlanta, Georgia, 5378 – 5392. Wilson, C.A., Novak, J.T. 2009. Hydrolysis of macromolecular components of primary and secondary wastewater sludge by thermal hydrolytic pretreatment. Water Research, 43 (18), 4489 – 4498. World Health Organization (WHO). 2008. Guidelines for drinking-water quality, incorporating 1st and 2nd Addenda, Recommendations, 3rd Edition, Volume 1, Geneva, Switzerland. Wojciechowska, E. 2005. Application of microwaves for sewage sludge conditioning. Water Research, 39 (19), 4749 – 4754. Wojcik, K., Dobrucki, J.W. 2008. Interaction of a DNA intercalator DRAQ5, and a minor groove binder SYTO17, with chromatin in live cells – influence on chromatin organization and histone-DNA interactions. Cytometry Part A, 73, 555 – 562.  Woo, I.S., Rhee, I.K., Park, H.D. 2000. Differential damage in bacterial cells by microwave radiation on the basis of cell wall structure. Applied and Environmental Microbiology, 66 (5), 2243 – 2247. Wu, T. 2008. Environmental perspectives of microwave applications as remedial  114  alternatives: review. Practice Periodical of Hazardous, Toxic, and Radioactive Waste Management, 12 (2), 102 – 115.  Xing, C.H., Yamamoto, K., Fukushi, K. 2006. Performance of an inclined-plate membrane bioreactor at zero excess sludge discharge. Journal of Membrane Science, 275, 175–186. Yeow, S.K., Peng, W.L. 2012. Application of ultrasound pretreatment for sludge digestion. Biogas Production. Edited by: Mudhoo, A. Scrivener Publishing LLC, USA, 91 – 136.  Yu, G.H., He, P.J., Shao, L.M., Zhu, Y.S. 2008. Extracellular proteins, polysaccharides, and enzymes impact on sludge aerobic digestion after ultrasonic pretreatment. Water Research, 42 (8-9), 1925 – 1934. Yu, Q., Lei, H.Y., Li, Z., Li, H.L., Chen, K., Zhang, X.H., Liang, R.L. 2010. Physical and chemical properties of waste-activated sludge after microwave treatment. Water Research, 44, 2841 – 2849. Zeeman, G., Sutter, K., Vens, T., Koster, M., Wellinger, A. 1988. Psychrophilic digestion of dairy and pig manure: startup procedure of batch, fed-batch and CSTR-type digesters. Biological Wastes, 26, 15 – 31. Zhang, P., Zhang, G., Wang, W. 2007. Ultrasonic treatment of biological sludge: Floc disintegration, cell lysis and inactivation. Bioresource Technology, 98 (1), 207 – 210. Zhang, S., Zhang, P., Zhang, G., Fan, J., Zhang, Y. 2012. Enhancement of anaerobic sludge digestion by high-pressure homogenization. Bioresource Technology, 118, 496 – 501.    115  Appendices Appendix A | Preliminary results for S16:SO and FDA assays  Figure A.1. Preliminary results in terms of live:dead cell ratios for S16:SO assay (n = 4) showing stepwise dilutions of  % live cells (CL) to % dead cells (CD).  Figure A.2. Preliminary results for FDA assay (n = 6) showing fluorescence measurements (as arbitrary fluorescence units, AFU) of both live control (CL) and control dead (CD) tWAS samples  116  over a 3 hour time period. Where the equation for live control is y = 47969x + 0.752, and the equation for dead control is y = 1111x + 148.45. Appendix B | Calibration curves for measurements of AD data  Figure B.1. Calibration curve for biogas measurement using a manometer (at STP: 0°C, 1 atm), showing the height of the displaced volume within the manometer converted into volume of gas measured (equation is y = 5.0671x – 1E-05).      117  Figure B.2. Calibration curve for chemical oxygen demand (COD), measured at 600 nm using a spectrophotometer (equation is y = 2359.5x – 0.9308).   Figure B.3. Calibration curve for ammonia concentration (mg/L) measured against mV reading of samples (equation is y = -21.49ln(x) – 66.757).                   118  Appendix C | Tukey Kramer method for post hoc analyses of assay data All microbial characterization data were statistically analyzed using the following steps: 1) ANOVA of all test and control groups 2) Post-hoc analysis using the Tukey Kramer method ANOVA tests run for both the S16:SO and FDA viability assays possessed P < 0.05, indicating that there was an overall difference between all groups analyzed. However, the one way ANOVA used does not specify significant values between individual groups. Therefore, a post-hoc analysis was run to determine whether three or more individual means differed significantly. The Tukey Kramer method was chosen as it is designed for experiments in which the n-values are not equal. In this method, the absolute difference between the means of two groups are compared to a critical range that is calculated using the studentized range distribution. Any observed difference greater than the calculated critical range is considered significant with 95% confidence, as α = 0.05 for this study. The Tukey Kramer test was performed for each pair of groups selected (see Appendix D) for both the S16:SO and FDA assays, respectively. An equation (C.1) was used to calculate the critical range between compared groups. 𝐶𝑟𝑖𝑡𝑖𝑐𝑎𝑙 𝑟𝑎𝑛𝑔𝑒 =  𝑄𝑢 ×  √𝑀𝑆𝑊2× (1𝑛1+1𝑛2)     (C.1) Where MSW is the mean of squares within groups provided by the initial ANOVA, n1 and n2 are the respective n values of Group 1 and Group 2 being compared, and Qu is the value selected using the standardized range distribution (found in any statistical textbook), α value (for this study, α = 0.05), and the numerator and denominator degrees of freedom (DF).  The numerator DF is the total number of groups analyzed in the experiment. The denominator DF is the sum of the number of n values for each tested group in the experiment.  119  Appendix D | Statistical analyses for microbial characterization data Table D.1. ANOVA results and summary for S16:SO assay. Table includes the sum of squares (SS), degrees of freedom (df), mean of squares (MS), F value (F), P value, and the F critical value (F crit).        Source of Variation SS df MS F P value F crit Between Groups 66.517 10 6.65 155.2 1.47E-34 2.02 Within Groups 2.1854 51 0.043           Total 68.703 61     Summary       Groups Count Sum Average Variance 1 CL 10 29.338 2.934 0.0897 2 CD 8 1.859 0.232 0.00392 3 MW60 4 2.255 0.564 0.00225 4 MW80 8 1.737 0.217 0.00285 5 MW120 6 1.787 0.298 0.00307 6 MW160 4 1.027 0.257 3.26E-06 7 US1060 6 12.868 2.145 0.120 8 US2060 4 8.077 2.019 0.0414 9 US2280 4 8.131 2.033 0.0832 10 US50100 4 4.983 1.246 0.0732 11 US60100 4 3.564 0.891 0.0378  Table D.2. Tukey Kramer post-hoc analysis of S16:SO assay results for live control (CL), dead control (CD), microwave (MW) irradiated, and ultrasonicated (US) tWAS samples at designated temperatures and intensities. The data are significantly different if the absolute difference (AD) between the two compared groups (Group 1 and Group 2) > calculated critical range (CR). Results show “true” for a statistically significant difference between the two groups under comparison, and “false” for differences between results that are not statistically significant. Group 1 Group 2 AD CR Results Dead Control Live Control 2.70 0.33 TRUE MW 60°C Live Control 2.37 0.41 TRUE MW 80°C Live Control 2.71 0.33 TRUE MW 120°C Live Control 2.63 0.36 TRUE MW 160°C Live Control 2.67 0.41 TRUE US 60%, 10min Live Control 0.78 0.36 TRUE US 60%, 20min Live Control 0.91 0.41 TRUE US 80%, 22min Live Control 0.90 0.41 TRUE  120  Group 1 Group 2 AD CR Results US 100%, 50min Live Control 1.68 0.41 TRUE US 100%, 60min Live Control 2.04 0.41 TRUE MW 60°C Dead Control 0.33 0.42 FALSE MW 80°C Dead Control 0.015 0.34 FALSE MW 120°C Dead Control 0.065 0.37 FALSE MW 160°C Dead Control 0.024 0.42 FALSE US 60%, 10min Dead Control 1.91 0.37 TRUE US 60%, 20min Dead Control 1.78 0.42 TRUE US 80%, 22min Dead Control 1.80 0.42 TRUE US 100%, 50min Dead Control 1.01 0.43 TRUE US 100%, 60min Dead Control 0.65 0.42 TRUE MW 80°C MW 60°C 0.34 0.42 FALSE MW 120°C MW 60°C 0.26 0.45 FALSE MW 160°C MW 60°C 0.30 0.49 FALSE MW 120°C MW 80°C 0.081 0.37 FALSE MW 160°C MW 80°C 0.039 0.42 FALSE MW 160°C MW 120°C 0.041 0.45 FALSE US 60%, 20min US 60%, 10min 0.12 0.45 FALSE US 80%, 22min US 60%, 10min 0.11 0.45 FALSE US 100%, 50min US 60%, 10min 0.89 0.45 TRUE US 100%, 60min US 60%, 10min 1.25 0.45 TRUE US 80%, 22min US 60%, 20min 0.013 0.49 FALSE US 100%, 50min US 60%, 20min 0.77 0.49 TRUE US 100%, 60min US 60%, 20min 1.12 0.49 TRUE US 100%, 50min US 80%, 22min 0.78 0.49 TRUE US 100%, 60min US 80%, 22min 1.14 0.49 TRUE US 100%, 60min US 100%, 50min 0.35 0.49 FALSE US 60%, 10min MW 60°C 1.45 0.45 TRUE US 60%, 20min MW 60°C 1.45 0.49 TRUE US 80%, 22min MW 60°C 1.46 0.49 TRUE US 100%, 50min MW 60°C 0.68 0.49 TRUE US 100%, 60min MW 60°C 0.32 0.49 FALSE US 60%, 10min MW 80°C 1.92 0.45 TRUE US 60%, 20min MW 80°C 1.80 0.49 TRUE US 80%, 22min MW 80°C 1.81 0.49 TRUE US 100%, 50min MW 80°C 1.02 0.49 TRUE US 100%, 60min MW 80°C 0.67 0.49 TRUE    121  Table D.3. ANOVA results and summary for FDA enzymatic assay. Table includes the sum of squares (SS), degrees of freedom (df), mean of squares (MS), F value (F), P value, and the F critical value (F crit).        Source of Variation SS df MS F P value F crit Between Groups 7060549026 10 706054902.6 109.7 2.7E-34 1.99 Within Groups 379642450.2 59 6434617.8           Total 7440191476 69     Summary       Groups Count Sum Average Variance 1 CL 6 215348.6 35891.4 22209229.5 2 CD 7 12011.5 1715.9 35143.3 3 MW60 6 33098.1 5516.3 441619.8 4 MW80 6 21061.2 3510.2 171185.0 5 MW120 6 11417.3 1902.9 42284.8 6 MW160 6 13865.0 2310.8 104174.1 7 US1060 9 174419.9 19380.0 10356525.3 8 US2060 6 107914.7 17985.8 6729301.0 9 US2280 6 110787.9 18464.7 22327137.9 10 US50100 6 50113.9 8352.3 5120022.8 11 US60100 6 64569.4 10761.6 2170922.8  Table D.4. Tukey Kramer post-hoc analysis of FDA assay results for live control (CL), dead control (CD), microwave (MW) irradiated, and ultrasonicated (US) tWAS samples at designated temperatures and intensities. The data are significantly different if the absolute difference (AD) between the two compared groups (Group 1 and Group 2) > calculated critical range (CR). Results show “true” for a statistically significant difference between the two groups under comparison, and “false” for differences between results that are not statistically significant. Group 1 Group 2 AD CR Results Dead Control Live Control 13696.6 4767.0 TRUE MW 60°C Live Control 12150.0 4946.9 TRUE MW 80°C Live Control 12952.4 4946.9 TRUE MW 120°C Live Control 13595.4 4946.9 TRUE MW 160°C Live Control 13432.2 4946.9 TRUE US 60%, 10min Live Control 6604.5 4515.9 TRUE US 60%, 20min Live Control 7162.2 4946.9 TRUE US 80%, 22min Live Control 6970.7 4946.9 TRUE US 100%, 50min Live Control 11144.1 4946.9 TRUE  122  Group 1 Group 2 AD CR Results US 100%, 60min Live Control 10051.9 4946.9 TRUE MW 60°C Dead Control 1546.5 4767.0 FALSE MW 80°C Dead Control 744.1 4767.0 FALSE MW 120°C Dead Control 101.1 4767.0 FALSE MW 160°C Dead Control 264.3 4767.0 FALSE US 60%, 10min Dead Control 7092.0 4318.0 TRUE US 60%, 20min Dead Control 6534.3 4767.0 TRUE US 80%, 22min Dead Control 6725.8 4767.0 TRUE US 100%, 50min Dead Control 2552.4 4767.0 FALSE US 100%, 60min Dead Control 3644.6 4767.0 FALSE MW 80°C MW 60°C 802.4 4946.9 FALSE MW 120°C MW 60°C 1445.3 4946.9 FALSE MW 160°C MW 60°C 1282.2 4946.9 FALSE MW 120°C MW 80°C 642.9 4946.9 FALSE MW 160°C MW 80°C 479.7 4946.9 FALSE MW 160°C MW 120°C 163.1 4946.9 FALSE US 60%, 20min US 60%, 10min 557.6 4515.9 FALSE US 80%, 22min US 60%, 10min 366.1 4515.9 FALSE US 100%, 50min US 60%, 10min 4539.5 4515.9 TRUE US 100%, 60min US 60%, 10min 3447.3 4515.9 FALSE US 80%, 22min US 60%, 20min 191.5 4946.9 FALSE US 100%, 50min US 60%, 20min 3981.8 4946.9 FALSE US 100%, 60min US 60%, 20min 2889.6 4946.9 FALSE US 100%, 50min US 80%, 22min 4173.4 4946.9 FALSE US 100%, 60min US 80%, 22min 3081.2 4946.9 FALSE US 100%, 60min US 100%, 50min 1092.2 4946.9 FALSE US 60%, 10min MW 60°C 4987.7 4515.9 TRUE US 60%, 20min MW 60°C 4987.7 4946.9 TRUE US 80%, 22min MW 60°C 5179.3 4946.9 TRUE US 100%, 50min MW 60°C 1005.8 4946.9 FALSE US 100%, 60min MW 60°C 2098.0 4946.9 FALSE US 60%, 10min MW 80°C 6347.9 4515.9 TRUE US 60%, 20min MW 80°C 5790.2 4515.9 TRUE US 80%, 22min MW 80°C 5981.7 4515.9 TRUE US 100%, 50min MW 80°C 1808.3 4515.9 FALSE US 100%, 60min MW 80°C 2900.5 4515.9 FALSE     123  Appendix E | Correlation analysis for microbial characterization data A correlation analysis was performed using Pearson’s r value (correlation coefficient) in order to determine if a directly linear relationship correlated between the S16:SO and FDA assays by using Microsoft Excel. There are three possible outcomes to the correlation analysis, pertaining to a value from 1 ≥ r ≥ -1: 1. Positive correlation (1 ≥ r ≥ 0) 2. Negative correlation (-1 ≤ r ≤ 0) 3. No correlation (r ≈ 0) The results of the correlation analysis are shown in Figure E.1 and Table E.1. The positive value (r = 0.97) indicates that there is a strong correlation between the two microbial characterization assays, and that as the live:dead cell ratio increases the relative enzymatic activity of the microbial population increases as well. The r value was calculated for the 95 % confidence interval (p value = 0.05).  Figure E.1. Graph of correlation analysis of the S16:SO (live:dead) cell ratios compared to the relative fluorescence of the FDA results. A positive correlation is indicated by the increasing relative fluorescence to increasing live:dead cell ratio. Sample size (n) is n = 11.   124  Table E.1. Result of correlation analysis indicating a positive direct linear correlation between the two microbial characterization assays, S16:SO and FDA assays. The sample size (n) is n = 11.  Live/Dead Ratio FDA Live/Dead Ratio 1  FDA 0.97 1  

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