text:
UBC Theses and Dissertations /
Biogeochemical characterization of the hyporheic zone...
UBC Theses and Dissertations
Biogeochemical characterization of the hyporheic zone below the Fraser River near Vancouver, British… Zima, Michael K. 2016
Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.
Title Biogeochemical characterization of the hyporheic zone below the Fraser River near Vancouver, British Columbia
Creator Zima, Michael K.
Publisher University of British Columbia
Date Issued 2016
Description The aim of this thesis is to characterize the biogeochemical conditions in the hyporheic zone of the Fraser River in order to help assess the fate of creosote-derived contaminants in groundwater. By investigating hyporheic biogeochemistry, a framework is established for further studies which could quantitatively assess the potential for monitored natural attenuation as a remedial option for groundwater contamination. At our site on the North Arm of the Fraser, groundwater flows horizontally towards the river at a velocity of 0.1 to 0.2 m/day. Transverse mixing between fresh groundwater and underlying saline groundwater is believed to be responsible for cation exchange reactions. From intertidal monitoring wells out to the hyporheic zone (HZ), Fe and Mn concentrations are found to increase despite dilution, and this is believed to be a result of either cation exchange, or oxidation of organic matter (OM), H₂S, or CH₄. As groundwater reaches the river, it turns upward and discharges into the HZ. In the river channel, fresh groundwater discharges to the HZ at distances between 70 and 85 m from the shoreline's high-water mark (HWM), while the deeper, saline groundwater discharges at distances greater than 100 m. A custom-made cryogenic probe was designed to collect representative, high-quality sediment samples from the HZ. A vertical characterization of biogeochemistry is presented. Chloride is used as a conservative tracer to determine a dilution factor of 170 in the top 85 cm of sediment, but dilution is believed to persist beyond this depth. DNA sequencing results infer a large variety of geochemical processes. Since multiple known aerobic microorganisms are found to have highest proportional abundances in the top 30 cm of the HZ, this depth range is considered to represent the portion of the HZ which is continually exposed to significant amounts of oxygen from the river. Porewater geochemistry also supports this delineation, as Fe(II) is non-detectable above the depth of 30 cm. The estimation of residence time in the HZ, the delineation of the aerobic portion of the HZ, and the characterization of dilution rates provide valuable information on the biogeochemical and physical components of natural attenuation in the HZ.
Genre Thesis/Dissertation
Rights Attribution-NonCommercial-NoDerivatives 4.0 International
URI http://hdl.handle.net/2429/59069
Degree Master of Science - MSc
Program Geological Sciences
Affiliation Science, Faculty of
Earth, Ocean and Atmospheric Sciences, Department of
Degree Grantor University of British Columbia
GraduationDate 2016-11
Campus UBCV
Scholarly Level Graduate
Rights URI http://creativecommons.org/licenses/by-nc-nd/4.0/
AggregatedSourceRepository DSpace
24-ubc_2016_november_zima_michael.pdf [ 6.06MB ]
JSON: 24-1.0314121.json
JSON-LD: 24-1.0314121-ld.json
RDF/XML (Pretty): 24-1.0314121-rdf.xml
RDF/JSON: 24-1.0314121-rdf.json
Turtle: 24-1.0314121-turtle.txt
N-Triples: 24-1.0314121-rdf-ntriples.txt
Original Record: 24-1.0314121-source.json
24-1.0314121-fulltext.txt
24-1.0314121.ris
i    BIOGEOCHEMICAL CHARACTERIZATION OF THE HYPORHEIC ZONE BELOW THE FRASER RIVER NEAR VANCOUVER, BRITISH COLUMBIA   by  MICHAEL K. ZIMA  B.Sc., University of Guelph, 2013      A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE   in   THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES  (Geological Sciences)    THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)   August 2016   © Michael K. Zima, 2016 ii  Abstract   The aim of this thesis is to characterize the biogeochemical conditions in the hyporheic zone of the Fraser River in order to help assess the fate of creosote-derived contaminants in groundwater. By investigating hyporheic biogeochemistry, a framework is established for further studies which could quantitatively assess the potential for monitored natural attenuation as a remedial option for groundwater contamination.  At our site on the North Arm of the Fraser, groundwater flows horizontally towards the river at a velocity of 0.1 to 0.2 m/day. Transverse mixing between fresh groundwater and underlying saline groundwater is believed to be responsible for cation exchange reactions. From intertidal monitoring wells out to the hyporheic zone (HZ), Fe and Mn concentrations are found to increase despite dilution, and this is believed to be a result of either cation exchange, or oxidation of organic matter (OM), H2S, or CH4. As groundwater reaches the river, it turns upward and discharges into the HZ. In the river channel, fresh groundwater discharges to the HZ at distances between 70 and 85 m from the shoreline's high-water mark (HWM), while the deeper, saline groundwater discharges at distances greater than 100 m.   A custom-made cryogenic probe was designed to collect representative, high-quality sediment samples from the HZ. A vertical characterization of biogeochemistry is presented. Chloride is used as a conservative tracer to determine a dilution factor of 170 in the top 85 cm of sediment, but dilution is believed to persist beyond this depth. DNA sequencing results infer a large variety of geochemical processes.  Since multiple known aerobic microorganisms are found to have highest proportional abundances in the top 30 cm of the HZ, this depth range is considered to represent the portion of the HZ which is continually exposed to significant amounts of oxygen from the river. Porewater geochemistry also iii  supports this delineation, as Fe(II) is non-detectable above the depth of 30 cm. The estimation of residence time in the HZ, the delineation of the aerobic portion of the HZ, and the characterization of dilution rates provide valuable information on the biogeochemical and physical components of natural attenuation in the HZ.   iv  Preface  This thesis is composed mainly of unpublished material, with some reference to work previously published by other researchers. The author of this thesis is responsible for all field and lab work done under the supervision of Dr. Roger Beckie. The thesis was reviewed by Dr.'s Roger Beckie, Sean Crowe, and Uli Mayer, who provided mainly stylistic edits. The Methodology section entitled "DNA Extraction and Sequencing" was composed in part by Dr. Rachel Simister, who was also responsible for sequencing and analysis of DNA samples presented in this thesis. Figure 4.26 and Appendix G were provided by Dr. Mati Raudsepp and Lan Kato from the UBC Electron Microbeam / X-ray Diffraction Facility, who completed Rietveld refinement on mineralogical samples. Figures 2.2 and 3.1 are adapted with permission from applicable sources.   v  Table of Contents  Abstract ..................................................................................................................................................... ii Preface ..................................................................................................................................................... iv Table of Contents ...................................................................................................................................... v List of Tables ......................................................................................................................................... viii List of Figures .......................................................................................................................................... ix List of Reactions ..................................................................................................................................... xii List of Equations .................................................................................................................................... xiii List of Abbreviations ............................................................................................................................. xiv Acknowledgements ................................................................................................................................. xv Chapter 1 – Introduction ............................................................................................................................ 1 1.1 Purpose ................................................................................................................................................ 1 1.2 Research Objectives ............................................................................................................................ 1 1.3 The Hyporheic Zone ........................................................................................................................... 2 1.4 Monitored Natural Attenuation ........................................................................................................... 9 1.5 Site Description and History ............................................................................................................. 10 Chapter 2 – Characterization of Groundwater Discharge to the Fraser River .................................. 14 2.1 Objectives ......................................................................................................................................... 14 2.2 Previous Studies ................................................................................................................................ 14 2.3 Conceptual Model ............................................................................................................................. 17 2.4 Methodology ..................................................................................................................................... 20 2.5 Results and Discussion ..................................................................................................................... 24 Monitoring Well Conductivity ............................................................................................................ 24 Hyporheic Zone Conductivity ............................................................................................................. 25 Water Type Characterization .............................................................................................................. 28 Conceptual Model ............................................................................................................................... 34 Manganese Reduction in the Aquifer .................................................................................................. 36 Iron Reduction in the Aquifer ............................................................................................................. 39 Iron and Manganese Reduction in the Hyporheic Zone ...................................................................... 40 2.6 Conclusion ........................................................................................................................................ 42 Chapter 3 – Design and Development of a Sediment-Freezing Sampler ............................................. 44 3.1 Previous Approaches ........................................................................................................................ 44 vi  3.2 Design Considerations ...................................................................................................................... 46 Motivation ........................................................................................................................................... 46 Objectives ........................................................................................................................................... 46 Constraints .......................................................................................................................................... 47 3.3 Production and Testing ..................................................................................................................... 48 3.4 Sampling Methodology ..................................................................................................................... 54 3.5 Assessing Contamination from River Water..................................................................................... 58 3.6 Challenges in the Field ...................................................................................................................... 60 Chapter 4 – Biogeochemical Characterization of Hyporheic Zone Sediments ................................... 62 4.1 Introduction ....................................................................................................................................... 62 Motivation and Research Objectives .................................................................................................. 65 4.2 Methodology ..................................................................................................................................... 66 Core Sectioning ................................................................................................................................... 66 Porewater Extraction ........................................................................................................................... 67 Sequential Extractions......................................................................................................................... 69 DNA Extraction and Sequencing ........................................................................................................ 71 Sediment Mineralogy .......................................................................................................................... 73 4.3 Results and Discussion ..................................................................................................................... 74 Core Description ................................................................................................................................. 74 Porewater Geochemistry ..................................................................................................................... 75 Sequential Extractions......................................................................................................................... 83 DNA Sequencing ................................................................................................................................ 89 Identifiable Microbial Metabolisms .................................................................................................... 92 Sediment Mineralogy ........................................................................................................................ 102 Interpretation ..................................................................................................................................... 107 4.4 Conclusion ...................................................................................................................................... 112 Major Microbial Metabolisms and Geochemical Processes ............................................................. 112 Defining the Aerobic Portion of the Hyporheic Zone ....................................................................... 113 Implications for Discharging Contaminants ..................................................................................... 114 Chapter 5 – Summary and Recommendations ..................................................................................... 115 5.1 The River District Site .................................................................................................................... 115 5.2 Characterization of Groundwater Discharge to the Fraser River .................................................... 115 5.3 Design and Development of a Sediment-Freezing Sampler ........................................................... 116 vii  5.4 Biogeochemical Characterization of Hyporheic Zone Sediments .................................................. 118 5.5 Delineation of the Aerobic Hyporheic Zone and its Implications .................................................. 120 5.6 Direction of Research and Recommendations ................................................................................ 121 References ................................................................................................................................................ 124 Appendices ............................................................................................................................................... 134 Appendix A – Hyporheic Zone and Monitoring Well Chemistry Data ................................................ 134 Appendix B – Core Porewater Chemistry Data .................................................................................... 139 Appendix C – Figures 4.1, 4.2, 4.3, and 4.6 Plotted in mM Concentrations ........................................ 141 Appendix D – Gran Titration Data for Core Porewater ........................................................................ 143 Appendix E – Sediment Extraction Data .............................................................................................. 147 Appendix F – Extended DNA Sequencing Results............................................................................... 151 Appendix G – Additional Spectra for Rietveld Refinements ................................................................ 155 Appendix H – Additional Lab Photos ................................................................................................... 158  viii  List of Tables  Table 1.1 – Hydrostratigraphy and hydraulic conductivity of geological units at the River District ......... 13 Table 2.1 – Values used to calculate normalized specific conductance (NSC) .......................................... 34 Table 4.1 – Sequential extraction steps used to target exchangeable, amorphous, crystalline, and residual phases of Fe and Mn, and their original authors ......................................................................................... 69 Table 4.2 – Summary of microorganisms with known metabolisms categorized based on peak abundance in the aerobic zone, the transition zone, and the anaerobic zone .............................................................. 102 Table 4.3 – Results of quantitative phase analysis by Rietveld refinements, with relative amounts of crystalline phase minerals normalized to 100% ........................................................................................ 103 Table 4.4 – Summary of inferred geochemical processes in the HZ based on results from porewater analyses and sediment extractions ............................................................................................................ 112      ix  List of Figures  Figure 1.1 – Turbidity and discharge data supplied by Environment Canada .............................................. 3  Figure 1.2 – Primary water level in the Fraser River recorded near Whonnock, BC in 2015 ...................... 4  Figure 1.3 – Depiction of redox microzones in the hyporheic zone ............................................................. 7  Figure 1.4 – Typical Eh at which redox couples are equimolar for pH = 7 .................................................. 7  Figure 1.5 – Map showing regional and local setting of the Fraser River delta near Vancouver ............... 11  Figure 2.1 – Visual representation of the saline groundwater circulation cell which persists at the lowest reaches of the Fraser ................................................................................................................................... 15  Figure 2.2 – The maximum extent of the seawater wedge ......................................................................... 16  Figure 2.3 – Vertical profiles for naphthalene concentrations and specific conductance in five intertidal zone monitoring wells ................................................................................................................................. 18  Figure 2.4 – Conceptual site model of groundwater discharging to the Fraser River ................................. 19  Figure 2.5 – Method of transect completion across the Fraser River ......................................................... 21  Figure 2.6 – The KIST® sampler, designed by Peter Krahn of Environment Canada ............................... 22  Figure 2.7 – Effect of delayed titration on apparent alkalinity for HZ samples rich in Fe ......................... 24  Figure 2.8 – Specific conductance (SC) of groundwater from intertidal zone monitoring wells ............... 25  Figure 2.9 – Spatial distribution of specific conductance for all HZ groundwater samples ....................... 26  Figure 2.10 – Effect of chloride and sodium concentrations on specific conductance for monitoring well samples ........................................................................................................................................................ 27  Figure 2.11 – Effect of chloride and sodium concentrations on specific conductance in the HZ ............... 28  Figure 2.12 – Piper plot showing major ion geochemistry of all water samples ........................................ 29  Figure 2.13 – Dissolved concentrations of Mg2+, Ca2+, and Na+ for monitoring well samples .................. 33  Figure 2.14 – pH measurements from multi-level monitoring wells .......................................................... 33  Figure 2.15 – Normalized specific conductance for HZ and intertidal zone samples with inferred flow paths ............................................................................................................................................................ 35  Figure 2.16 – Manganese concentrations for the intertidal zone monitoring wells .................................... 37  Figure 2.17 – Correlation between Mn and Ca concentrations for HZ and monitoring well samples ........ 38 x  Figure 2.18 – Ca and Mn concentrations in the hyporheic zone plotted against their location relative to the high water mark .......................................................................................................................................... 39  Figure 2.19 – Iron concentrations for the intertidal zone monitoring wells ................................................ 40  Figure 2.20 – Mn and Fe concentrations in the HZ with distance from the high water mark .................... 41  Figure 3.1 – Side-view diagram showing Roschinski's sampling method ................................................. 45  Figure 3.2 – Diagram showing unification of sediment freezing sampler and KIST sampler .................... 49  Figure 3.3 – Diagram showing the distribution of CO2 orifices along the length of the manifold ............. 51  Figure 3.4 – Laboratory setup for sampler freezing tests............................................................................ 52  Figure 3.5 – Temperature data from a laboratory freezing test ................................................................... 53  Figure 3.6 – Resulting ice core from a laboratory freezing test .................................................................. 53  Figure 3.7 – Sequence of video frames shows progress of sampler insertion ............................................ 54  Figure 3.8 – Freeze pipe with frozen sediment being removed from sampler assembly ............................ 54  Figure 3.9 – Detailed photo of frozen sediment core .................................................................................. 56  Figure 3.10 – Photo of sediment core being covered in plastic wrap ......................................................... 57  Figure 3.11 – Photo of sediment core being placed into dry ice cooler ...................................................... 57  Figure 3.12 – Piper plot showing major ion chemistry of extracted porewater samples to assess contaminantion by river water .................................................................................................................... 59  Figure 4.1 – Dissolved concentrations of Na, Ca, Mg, and K from core porewater ................................... 75  Figure 4.2 – Total Fe, Fe(II), and Mn concentrations from core porewater ............................................... 76  Figure 4.3 – Cl, NO3, and SO4 concentrations from core porewater .......................................................... 77  Figure 4.4 – Specific conductance of core porewater ................................................................................. 79  Figure 4.5 – pH of core porewater .............................................................................................................. 79  Figure 4.6 – Dissolved O2 concentrations from core porewater ................................................................. 81  Figure 4.7 – Alkalinity of core porewater ................................................................................................... 81  Figure 4.8 – Piper plot showing trend in major ion chemistry for core porewater ..................................... 82  Figure 4.9 – Results from sequential Fe extractions ................................................................................... 84 Figure 4.10 – Results from sequential Mn extractions ............................................................................... 85  xi  Figure 4.11 – Relationship between porewater concentrations and exchangeable fractions of Fe/Mn ...... 87  Figure 4.12 – Results from Fe speciation during extraction of amorphous sediment fractions .................. 88  Figure 4.13 – Distribution of major phyla with depth in percent of total sequences .................................. 90  Figure 4.14 – Distribution of major classes with depth in percent of total sequences ................................ 90  Figure 4.15 – Total number of sequences identified per depth interval ...................................................... 91  Figure 4.16 – Variation in proportional abundance with depth for α-proteobacteria ................................. 93  Figure 4.17 – Variation in proportional abundance with depth for β-proteobacteria ................................. 94  Figure 4.18 – Variation in proportional abundance with depth for γ-proteobacteria .................................. 95  Figure 4.19 – Variation in proportional abundance with depth for δ-proteobacteria .................................. 96  Figure 4.20 – Variation in proportional abundance with depth for Sulfuricurvum .................................... 97  Figure 4.21 – Variation in proportional abundance with depth for Acidobacteria ..................................... 98  Figure 4.22 – Variation in proportional abundance with depth for Nitrospira ........................................... 99  Figure 4.23 – Variation in proportional abundance with depth for Planctomycetacia ............................... 99  Figure 4.24 – Variation in proportional abundance with depth for Chthoniobacter ................................. 100  Figure 4.25 – Variation in proportional abundance with depth for Candidatus Methanoperedens and Marine Benthic Group D .......................................................................................................................... 101  Figure 4.26 – Rietveld refinement plot for sample depth 22.5 cm showing relative amounts of crystalline phase minerals normalized to 100% ......................................................................................................... 104  Figure 4.27 – Scanning electron microscope (SEM) image showing white particulate accumulations on sediment grains believed to be iron precipitates ....................................................................................... 105  Figure 4.28 – Qualitative evaluation of elemental composition from "area A" in Figure 4.27 ................ 106  Figure 4.29 – Iron-to-chloride and manganese-to-chloride ratios for core porewater .............................. 108            xii  List of Reactions  Reaction 2.1 – Cation exchange with Ca and Na ........................................................................................ 31 Reaction 2.2 – Calcite precipitation generating acidity .............................................................................. 32 Reaction 3.1 – Dissolution of CO2 .............................................................................................................. 48 Reaction 3.2 – Formation of carbonic acid ................................................................................................. 48 Reaction 4.1 – Sulfate reduction via oxidation of organic matter ............................................................... 65     xiii  List of Equations  Equation 2.1 – Calculation of normalized specific conductance (NSC) ..................................................... 34  Equation 5.1 – Calculation of specific discharge with Darcy's Law ......................................................... 121 Equation 5.2 – Calculation of linear groundwater velocity ...................................................................... 121 Equation 5.3 – Calculation of vertical residence time in the HZ .............................................................. 121    xiv  List of Abbreviations  BTEX Benzenes, toluenes, ethylbenzenes and xylenes DDIW Distilled, de-ionized water DNA  Deoxyribonucleic acid DNAPL Dense non-aqueous phase liquid DO Dissolved oxygen EDX Energy-dispersive x-ray EPA Environmental Protection Agency (US) HAC Halogenated aromatic compounds HWM High-water mark HZ Hyporheic zone IZ Intertidal zone MNA Monitored natural attenuation MTBE Methyl tert-butyl ether NPT National pipe thread taper NSC Normalized specific conductance NTU Nephelometric turbidity unit OM Organic matter OTU Operational taxonomic unit PAH Polycyclic aromatic hydrocarbons PCE Tetrachloroethylene (perchloroethylene) RNA Ribonucleic acid SC Specific conductance SEM Scanning electron microscope SRB Sulfate-reducing bacteria TCE Trichloroethylene VC Vinyl chloride XRD X-ray diffraction    xv  Acknowledgements   I'd like to thank the members of my supervisory committee – Dr.'s Roger Beckie, Sean Crowe, and Uli Mayer. Your vast reserves of knowledge and experience made my thesis work smooth and enjoyable. Your patience, guidance, and advice helped me learn as a student and grow as a hydrogeologist. I am extremely appreciative of the detailed comments and suggestions you provided for my thesis. Your edits helped build on its completeness and added coherence to my arguments.  Thank you to Dr. Mario Bianchin, Tilman Roschinski, and Kun Jia who helped pave the way for hyporheic zone research in our group. Your respective theses helped me to direct and compose my own. Thank you to Jörn Unger for your workmanship and dedication. Your resourcefulness and attention to detail were highly valued in our engaging conversations during sampler development. To Chris Payne and Lora Pakhomova, thank you for your ingenuity and patience in the field. I have no doubt that our success was in large part because of your steady hands at the helm of the Kraken.  Thank you to Laura Laurenzi, Maureen Soon, Dr. Mati Raudsepp, Lan Kato, and Elisabetta Pani for your supportive analytical expertise. Thank you to Dr. Rachel Simister and Céline Michiels for your direction and insightfulness which helped to ignite my interest in geomicrobiology. Thank you to Peter Krahn for your guidance in the field as well as your willingness to share techniques and experience from your own career. Thank you to the Hydrogeology group at Golder Associates Ltd. of Vancouver, who conferred such a superb research opportunity.  Lastly, I am obliged to thank my assistants in the field – or perhaps I should say, my companions in the field; Laura Ramsden, Jarod Devries, Jordan Zak, Andrea Chong, Keelin Scully, Cassie Ragan, Brendan Gibbs, Olenka Forde, Elliott Skierszkan, and Coni Nicolau. You were all instrumental in solving the problems we encountered in the field, thus helping to keep me "afloat" on the Fraser. 1  Chapter 1 – Introduction  1.1 Purpose  This thesis presents findings from research conducted on the hyporheic zone (HZ) underlying the North Arm of the Fraser River near Vancouver, British Columbia. The HZ is a dynamic region beneath a stream or river where groundwater and surface water interact. HZs have been mainly characterized in the context of smaller hydrological systems because fewer logistical challenges exist in the field. Consequently, our knowledge of HZs in large river deltas such as the Fraser is less established. Only recently has our understanding of dominant mixing mechanisms in these settings and their relative importance emerged [1, 2]. Although physical and chemical processes in large, tidally-affected HZs have been characterized to some degree, the biogeochemistry of such systems is still not well understood.  The aim of this research was to elucidate basic relationships between geochemistry and microbiology in the hyporheic sediments below the Fraser.  In this thesis, discussion is provided in the context of biogeochemistry, referring to the control and dependence of microbes on their geochemical environment. By investigating hyporheic biogeochemistry, we aim to lay the framework for further studies which could quantitatively assess the potential for monitored natural attenuation as a remedial option for local groundwater contamination.  1.2 Research Objectives  This research was driven by two major objectives. The first objective is to examine principal locations of groundwater discharge as evidenced by salinity and major ion geochemistry. This work was focused at the 10-100m scale and yielded insight on groundwater flow paths, fresh-saline mixing, and discharge patterns to the Fraser River. Major applications of this research include improved predictions of potential contaminant discharge points and improved groundwater model calibration for hydrogeologists currently 2  working at the River District site. The second objective is to investigate the biogeochemistry of the hyporheic sediments at the 0.1-1m scale to better understand how groundwater contaminants may be transformed prior to entering a river. This portion of the research involved the development of innovative field equipment for sediment sampling, and provided a snapshot of the biogeochemical conditions which exist in the HZ below the Fraser River estuary.  1.3 The Hyporheic Zone  HZs are known to provide a wide array of ecological functions.  Microbial communities fostered in HZs are highly diverse and adaptable [3-7]. Where waters of differing chemical composition mix, reactivity is high, and a large range of environmental conditions persist [5, 8-10]. Apart from serving as essential habitat and spawning ground for many organisms [11], commonly characterized functions of the HZ include the removal of organic matter [12, 13] and excess nutrients [5, 14, 15] from rivers thus mitigating eutrophication [3, 16]. HZs also have the ability to attenuate anthropogenic contaminants from both surface water and groundwater [10, 17, 18]. Accordingly, the HZ has been termed a "river's liver" [3].  Mixing between freshwater and groundwater is controlled by mechanisms which operate at varying spatial and temporal scales [2]. In the Fraser River near Vancouver, two dominant factors are the seasonal peak in discharge due to inland spring melting (i.e. freshet) and tidal pumping. As shown in Figure 1.1, fluvial discharge increases by about an order of magnitude during freshet. This rise in river stage increases hydraulic head in the river, reversing hydraulic gradients in the HZ so that oxygenated freshwater recharges the underlying aquifer [2]. On a shorter time scale, tides impact groundwater-surface water interaction by inducing oscillatory flow between periods of flooding and ebbing tide [2]. Bianchin et al. (2010) quantified the magnitude of tidally-induced hyporheic exchange using a time series of temperature and hydraulic head data from various depths and concluded that solutes discharging to the river are significantly diluted. As a result of oscillatory flow and the effects of hydrodynamic dispersion 3  in the HZ, dilution of discharging groundwater was estimated to be on the order of 99.9% and 84% during flooding and ebbing tides, respectively.   Figure 1.1 – Turbidity and discharge data supplied by Environment Canada. Turbidity measured from water quality buoy on Main Arm near Tilbury Island. Discharge measured from hydrometric station at Hope, BC.  During periods of high fluvial discharge, tidal signals along the Fraser become less noticeable (see Figure 1.2). Concurrently, dune-induced (or bed-form-induced) flow becomes more prominent in the HZ [2]. Dune-induced mixing is a function of hydrostatic head variations along the surface of a dune. This results in the downwelling of surface water into the stoss (or upstream) side of a dune, with subsequent emergence of that flow path on the lee side. Consecutive sections of downwelling and upwelling can be referred to as hyporheic flow cells or hyporheic exchange flows [19, 20] which can also develop through interaction with woody debris on the riverbed [21].  0200040006000800010000120001400001/2012 04/2012 06/2012 09/2012 12/2012 03/2013 06/2013 09/2013 12/2013 03/2014 06/2014 09/2014 12/2014050100150200250Discharge (m³/s) Turbidity (NTU) Turbidity and Discharge on the Fraser River near Vancouver - 2012 to 2014 Tubidity on Main Arm (NTU)Discharge at Hope, BC (m³/s)22-Jun 11700 m³/s 1-May 173 NTU 19-Apr 164 NTU 24-Apr 186 NTU 22-Apr 205 NTU 28-May 9920 m³/s 17-May 10100 m³/s 4   Figure 1.2 – Primary water level in the Fraser River recorded near Whonnock, BC in 2015. Diurnal variations are produced by tides. During freshet, the magnitude of variations decreases substantially. Data obtained from Environment Canada.  Hester et al. (2013) modelled the interaction of groundwater and surface water in hyporheic flow paths induced by riverbed dunes. Aside from demonstrating the system's high sensitivity to geological heterogeneity and dispersivity, the modelling results implied that mixing between groundwater and surface water occurs in a remarkably thin layer [19]. The effects of tidal pumping or freshet were not assessed, however. It is expected that diurnal variability in river stage due to tides would act to "smear" this mixing zone in the vertical direction. Dunes and bed-form structures have previously been surveyed along the Fraser River Main Arm. Along measurement transects, mean crest-to-trough heights of 1.47m [22] and 1.2m [23] have been calculated. In homogeneous sediments such as those in the Fraser, however, hydraulic conductivity in the HZ and upwelling flow rate were believed to be much more important for mixing than dunes [19].  33.544.555.566.515-Apr19-Apr23-Apr27-Apr01-May05-May24-Apr28-Apr02-May06-May10-May14-May18-May22-May26-May30-May03-Jun07-Jun11-Jun15-Jun19-Jun23-Jun27-Jun01-Jul05-Jul10-Jul14-Jul18-Jul22-JulStage (m) Primary Water Level on the Fraser River at Whonnock, BC 2015 freshet 5  In addition to the formation of hyporheic flow cells, dunes encourage HZ mixing through bedload transport. As the stoss side of a dune is eroded, sediment is transported over the crest and deposited on the lee side, trapping fresh surface water in the pore space [24]. This freshly-deposited porewater then interacts with upwelling groundwater until the next erosion sequence begins. Furthermore, layered deposition of sediment on the lee side of dunes is hypothesized to generate small-scale anisotropy in the HZ [25]. In the Fraser River estuary, dune-associated transport is thought to constitute nearly all bed-material transport [23]. Measurements from bathymetric soundings on the Main Arm by Villard and Church (2003) during the freshet of 1989 show dune migration velocity to be on the order of 10 metres per day. Although periods of low flow generally result in smaller dunes and lower migration velocities [23], dune-associated sediment transport is still an important mechanism that shifts secondary minerals away from locations in the HZ where they originally formed. This ultimately makes studying secondary mineral precipitation in the HZ much more challenging.  Secondary precipitation of iron onto sediment grains in the HZ occurs where reduced upwelling groundwater, enriched in ferrous iron, encounters oxygen-rich freshwater. As the aqueous ferrous iron is exposed to an increasingly positive redox potential, iron oxidation occurs, largely through biotic pathways [6, 10, 26]. Evidence of oxidative iron precipitation has been previously noted by Charette and Sholkovitz (2002) in Waquoit Bay, a shallow estuarine environment off the shores of Cape Cod, MA. Iron oxide-coated sands were evident in sediment cores by their dark red, yellow, and orange colours. Peak concentrations of iron precipitates were found at depths between 100 and 130 cm [27]. Other studies have also confirmed the presence of iron coatings on sediment grains [28-30]. Furthermore, similar oxidative precipitation is expected for manganese in the HZ [6, 10, 18]. In addition to serving as a visual indicator for the extent of mixing in the HZ, these precipitate coatings serve as adsorption sites for trace metals [18]. These processes may also be beneficial in filtering out toxic elements such as arsenic which would otherwise discharge into water bodies [10]. In many environments, reductive dissolution may simultaneously act to release arsenic bound or co-precipitated to the iron [31-33].  6   Although high concentrations of ferrous iron have been noted in the HZ of the Fraser River, the discovery of visible iron coatings on sediment grains has remained elusive [34]. One hypothesis for the lack of visible secondary iron precipitation is that high concentrations of organic matter generate localized, strongly reducing conditions, which sequester any available oxygen [34]. It is also likely, however, that continual erosion and deposition of fresh sediment through bedload transport make iron precipitation difficult to identify because it can't develop in significant concentrations.  In addition to controlling abiotic geochemistry, the downward diffusion of oxygen from the river into the HZ generates a spectrum of microbial niches, ranging from predominantly aerobic, to anaerobic, with increasing depth. Within this primary redox gradient are also smaller, isolated microzones of redox conditions which may differ substantially from immediately adjacent regions. These redox microzones may be related to lenses of finer-grained material, or pore-scale heterogeneities (Figure 1.3) [35]. In microzones of lower hydraulic conductivity, groundwater velocity is slower, and thus residence times increase. When the ratio of residence time to reaction time (referred to as a Damköhler number) increases, favourable redox reactions have more time to proceed [36]. As oxygen is depleted, a sequence of reduction processes characterized by the "redox ladder", proceeds as follows: denitrification (or nitrate reduction), reduction of Mn-oxides, reduction of Fe-oxides, sulfate reduction, and finally methanogenesis (or fermentation) [37, 38]. The typical redox potentials at which these reactions occur are plotted in Figure 1.4. Since microzones typically exist as pockets of finer-grained material with high concentrations of organic matter, they may serve as isolated sinks for electron acceptors in an otherwise oxygenated medium [35].  7   Figure 1.3 – Depiction of redox microzones in the hyporheic zone. Redox microzones may exist as small pore-scale heterogeneities, or as finer-grained lenses. Reduced groundwater discharges upward to the hyporheic zone. With increasing depth, different biogeochemical processes dominate. In microzones, redox conditions may differ substantially from regions which are immediately adjacent.   Figure 1.4 – Typical Eh at which redox couples are equimolar, for pH = 7. Values from Langmuir 1997, p. 416 [39].  The concept of redox microzones is well-supported in biogeochemistry. It is understood that the boundary between microzones and active groundwater flow paths serves as an advantageous habitat for microorganisms in that they can access a greater variety of resources [40]. Microbial communities found in microzones are generally located in biofilms, and tend to be composed of diverse, interacting species whose metabolisms are interdependent [40, 41]. Biofilms, which are "matrix-enclosed microbial accretions", proliferate in the HZ by adhering to sediment particle surfaces [42]. This allows biofilms to 8  access energy substrates, nutrients, and electron acceptors delivered by the porewater, while taking advantage of the structural rigidity offered by the porous media [5, 6]. Biofilms in the HZ are dense in organic matter, composed mainly of cells and extracellular polymeric materials. This structure is thought to provide a high number of reaction sites for both organic and inorganic contaminants [43-45].   In addition to decreasing contaminant mobility through biosorption, microorganisms in the HZ can biodegrade sorbed organic contaminants much faster than abiotic hydrolysis reactions [46]. Microbial enzymes are used to alter the shape and orientation of sorbed organic compounds, exposing weaker points in the structure, and making compounds more susceptible to breakdown reactions [37].  A variety of microbial metabolisms may be able to degrade a contaminant, but the effectiveness of a given degradation pathway varies based on the type of contaminant as well as site-specific conditions. Compounds such as benzene, toluene, ethylbenzene, and xylene (BTEX compounds), for example, demonstrate their highest degradation coefficients under aerobic respiration [47]. Polycyclic aromatic hydrocarbons (PAHs) such as naphthalene which have large molecular weights and low solubility degrade at rates up to two orders of magnitude higher in aerobic conditions than in anaerobic conditions [48, 49]. Conversely, some chlorinated compounds like tetrachloroethylene (PCE) and trichloroethylene (TCE) breakdown more rapidly under reducing conditions due to the oxidation state of carbon [47]. The same is not always true for their intermediate breakdown products, however (i.e. vinyl chloride, VC).  For many organic contaminants in groundwater, the aerobic environment sustained in the HZ serves as a defence against surface water contamination. Bianchin et al. (2011) propose that PAH attenuation seen below the Fraser River is a function of dispersion due to the oscillatory pumping of tides, and is likely supplemented by biodegradation [1]. Researchers who investigated the degradation of fuel oxygenates at a variety of other sites on Long Island, NY confirmed the importance of aerobic biodegradation in such environments, but estimated its contribution to total mass attenuation at just 3% [17]. They concluded that 9  utilizing the HZ's natural attenuation capacity during  contaminated site assessment merits serious consideration [17]. As the concept of monitored natural attenuation becomes more popular amongst remediation specialists, the need to obtain a detailed biogeochemical understanding of the processes involved becomes increasingly important.  1.4 Monitored Natural Attenuation  Monitored natural attenuation (MNA) is defined as a combination of physical, chemical, and biological processes which, "under favorable conditions, act without human intervention to reduce the mass, toxicity, mobility, volume, or concentration of contaminants in soil or groundwater" [50]. Implying more than just biodegradation, this umbrella term can constitute abiotic transformation to a less toxic form, or reduction of mobility and bioavailability as a result of sorption onto sediment particle surfaces [50].  The US Environmental Protection Agency (EPA) states that MNA should not be considered a presumptive or default remediation technique, but rather an option that be considered in combination with other strategies. Where MNA meets statutory and regulatory requirements and has been deemed appropriate given site-specific conditions, it may be beneficial [50]. Primary advantages of MNA include a lower volume of extracted waste for remediation and a decreased potential for transfer between different phases (i.e. increased dissolution from non-aqueous phase liquid). Furthermore, cost savings are potentially significant and human exposure is reduced. Disadvantages of MNA consist of longer time frames for remediation and monitoring, as well as increased complexity in site characterization. Ensuring public acceptance of the efficacy of MNA may also be challenging. Additionally, site conditions may at some point evolve, rendering MNA less effective [50].   MNA is most appropriate in situations where source control has been achieved and long-term performance monitoring has been established [51]. At high contaminant concentrations, MNA loses 10  effectiveness due to toxicity effects [52, 53]. As a result, groundwater systems experiencing substantial dilution, such as tidally affected environments and the HZ, may be better suited for MNA.  To demonstrate that sufficient MNA is occurring at a site, the EPA advocates for the use of three lines of evidence. These include (1) documented decrease in contaminant concentrations, (2) documented geochemical conditions which are assumed to be a result of microbial activity, and (3) documented microbiological degradation activity (in- or ex-situ) [50]. Unique, identifiable breakdown products of certain contaminants may also serve to demonstrate that degradation is occurring.  Although MNA may be appropriate as part of a remediation plan at onshore sites, justifying the utilization of MNA in the HZ and demonstrating its effectiveness are much more problematic – logistically and legally. This is because in some jurisdictions, the HZ may be defined as being part of a river, or the "receptor" itself [54].   When groundwater near a surface water body is contaminated, the appropriate remediation standard is often defined by the tolerances of organisms living in the water body. In British Columbia, this includes "phytoplankton, zooplankton, benthos, macrophytes and fish" [54]. The inclusion of benthos, which comprises bacteria essential to the process of biodegradation, produces a discrepancy. Utilizing MNA in the HZ, by default, requires that bacteria be exposed to contaminant concentrations which are higher than regulatory standards permit. If MNA is to be permitted in the HZ, site-specific allowances for higher contaminant concentrations would be required, or else current guidelines would need to be amended.  1.5 Site Description and History  The study area is a large property currently undergoing development on the North Arm of the Fraser River in Vancouver, BC, known as the River District (see Figure 1.5). Since the late 1800s, land use on 11  the site was associated with the lumber industry. Known operations included creosoting of piles, structural timbers, and railway ties. Creosote, a distilled form of coal tar commonly used for lumber preservation, was stored and utilized on the site until the early 1910s. Sawmill infrastructure was also added and expanded in the following decades. Operations ceased in 2002, with site demolition following shortly thereafter [55].   Figure 1.5 – Map showing regional and local setting of the Fraser River delta near Vancouver, BC, Canada  Creosote consists of a wide variety of hydrocarbons. Its exact composition varies depending on the origin and distillation method. PAHs make approximately 85% of the creosote mixture – naphthalene being the most prevalent compound. Also included are phenolic compounds, heterocyclic aromatic compounds (HACs), and BTEX compounds [56, 57]. The specific gravity of coal tar creosote ranges from 1.05 to 1.08, meaning that it behaves as a dense, non-aqueous phase liquid (DNAPL) [57, 58]. DNAPLs are 12  minimally responsive to hydraulic gradients and are transported as a function of gravity, viscosity, and interfacial tension.  As a result, DNAPLs often pool at the bottom of aquifers, where they remain perched on impermeable layers [37].  The River District site was first investigated for potential contamination in 1991, with PAHs being identified as contaminants of concern. Groundwater monitoring has been in place since the mid-1990s. Remediation has been completed at many parts of the site, including removal of all PAH contamination in the top-most 3 metres. Residual contamination persists at greater depths, however. A pump-and-treat system is in place at the site, establishing a capture zone for the dissolved portion of PAHs, thus minimizing contaminant discharge to the Fraser River. After being extracted, groundwater is treated in an on-site facility [55].  Stratigraphy at the River District site is as follows; the uppermost layer consists of mineral and wood waste fill (1-5 m thickness), which is underlain by an organic silt unit (1-6 m thickness), the Fraser River Sands unit (15-25 m thickness), and finally a marine clay unit. The marine clay unit dips slightly away from the river, and acts as an impermeable surface along which residual creosote rests in a few locations. Two distinct aquifers can be characterized at the site; a shallow, unconfined aquifer within the upper fill material, and a deep, mostly-confined regional aquifer which extends throughout the Fraser River Sands unit [55]. The organic silt unit acts as a leaky aquitard, where groundwater flow is predominantly vertical into the lower aquifer. Groundwater velocity in both aquifers is on the order of 0.1 m/day, towards the river. Estimated hydraulic conductivity values for the fill unit, the organic silt unit, and the upper and lower halves of the Fraser River Sands unit are shown in Table 1.1 [55].      13  Table 1.1 - Hydrostratigraphy and hydraulic conductivity of geological units at the River District. From Golder (2003). Unit Hydrostratigraphy Hydraulic conductivity (m/s) Mineral and wood waste fill Unconfined aquifer 5 x 10-5 Organic silt Leaky aquitard 1 x 10-7 Upper Fraser River Sands Mostly-confined aquifer 3 x 10-5 Lower Fraser River Sands 4 x 10-4   The lower aquifer is hydraulically connected to the Fraser River, and thus it responds to diurnal fluctuations in river stage caused by tides [57]. Gradient reversals are common near the discharge zone and are amplified in magnitude during freshet [2]. Variations in hydraulic head measured in onshore monitoring wells follow the offshore tidal pattern, with amplitude reduction and time lag proportional to the distance from the discharge zone [2].  Although the regional flow system in the Fraser River Sands consists primarily of connate, saline groundwater, a layer of fresh, meteoric groundwater is found in the upper portion of the aquifer. Both fresh and saline groundwater have been found to discharge into Fraser at a nearby location similar to the River District site, although fresh groundwater discharge was found to occur over a much smaller area of the HZ [1].   14  Chapter 2 – Characterization of Groundwater Discharge to the Fraser River  2.1 Objectives  Deep, saline groundwater is known to underlie shallow, fresh groundwater at the River District site. A major objective of the research presented in this chapter is to characterize the specific conductance of groundwater discharging in the HZ. By doing so, discharge locations of fresh and saline portions of the aquifer can be delineated. Significant motivation for this research is derived from a contaminant monitoring initiative at the River District site. By identifying areas of fresh and saline groundwater discharge in the HZ, potential contaminant discharge locations can be predicted based on their depth in the onshore aquifer.  2.2 Previous Studies  The dynamics of groundwater-surface water interaction below the Fraser are investigated by a handful of researchers. Neilson-Welch and Smith (2001) examine saline water intrusion adjacent to the Fraser, modelling salinity distributions and groundwater flow under density-dependent conditions. During rising tide, a "wedge" of seawater from the Strait of Georgia is known to migrate up the Fraser River basin. When the river stage is high enough to reverse gradients in the discharge zone, diluted seawater infiltrates the permeable HZ sediments [59]. This seawater, in contrast with upwelling groundwater, operates under density-dependent conditions, sinking down to the bottom of the Fraser River Sands unit. A saline circulation cell then develops, with saline water migrating up to 500 m away from the river along the underlying aquitard [59]. As the density contrast fades due to dilution, the flow turns upward and reverses direction, rejoining the local groundwater flow system as it travels towards the river (see Figure 2.1). Two distinct mixing zones exist along this flow path. The "lower" mixing zone is found at the base of the saline circulation cell, where fresh water trapped in the underlying aquitard leaks upward. The "upper" 15  mixing zone is found at the top of the circulation cell, where overturned, brackish groundwater interacts with local, fresh groundwater [60].    Figure 2.1 – Visual representation of the saline groundwater circulation cell which persists at the lowest reaches of the Fraser. It is driven by the seawater wedge (seen here in green), which migrates up the channel and intrudes the underlying aquifer under density-dependent conditions. Upper and lower mixing zones are originally reported by Jia (2015). Small white arrows denote upward fresh groundwater seepage from the underlying aquitard.  Intrusion of seawater into the permeable sediments of the Fraser is restricted to the lowest reaches of the delta. When low flow conditions converge with high tides, the extent of the seawater wedge is greatest. At its maximum, the wedge limits are estimated at 12 km and 20 km upstream of the mouths of the North and Main Arms, respectively [61] (see Figure 2.2). Upstream of these limits, however, groundwater discharging up into the central portion of the Fraser River channel is still saline [1]. The origin of this salinity is likely trapped connate water. It is derived from the mixing of meteoric and marine water in river sediments, in combination with diagenetic reactions which occurred during the progression of the Fraser River delta [62, 63]. Diffusion acting over thousands of years is also an important transport mechanism for salinity [64]. 16   Figure 2.2 – The maximum extent of the seawater wedge is shown in orange. When low flow coincides with high tides, the wedge migrates farthest upstream. Original limits reported by Ages and Woollard (1976). Adapted here from Neilson-Welch and Smith (2001).  Groundwater discharge is examined in previous research along the North Arm of the Fraser River. Bianchin et al. (2011) define the HZ of a site located upstream of the River District. Using geophysical techniques in combination with groundwater analyses, they determine that fresh groundwater discharges between 88 and 105 m from the shoreline, while saline groundwater discharges at distances greater than 105 m. The researchers conclude from field work and modelling that tidally-driven mixing in the HZ occurs to an approximate depth of 1 m.  Jia (2015) examines biogeochemical processes occurring as a result of fresh and saline groundwater interaction at the same site as Neilson-Welch (2001). Unlike the River District or the site studied by Bianchin et al., Jia's site is sufficiently far downstream that it is affected by salt water intrusion from the Fraser. Jia finds that Mn and Fe reduction are occurring, in addition to cation exchange reactions, where 17  fresh and saline groundwater mix in the aquifer. Sulfate reduction and the resultant production of alkalinity are believed to be occurring. Upstream from Jia's site, aquifers connected to the Fraser are less and less likely to be affected by salt water intrusion due to the limited extent of the seawater wedge.  With respect to contaminant monitoring initiatives at the River District site, previous work by Golder Associates Ltd. from August 2014 characterizes PAH concentrations in the aquifer. Naphthalene is found to be measurable below the intertidal zone (see Figure 2.3). Peak concentrations are found in saline groundwater at a depth of approximately 16 m. Data indicate that the depth to saline groundwater remains stable, since 2014 and 2015 conductivity datasets are similar.  2.3 Conceptual Model  The conceptual model developed for this thesis is based on an earlier version by Bianchin et al. (2011). Produced for a similar site on the Fraser, Bianchin's conceptual model is derived from resistivity and geochemistry data. At Bianchin's site, both fresh and saline groundwater are found to be discharging from the onshore aquifer into the Fraser River [1].  Based on the hydrostratigraphy identified by Golder (2003) for the River District site, a new conceptual site model (see Figure 2.4) is developed. Preliminary analysis of samples from monitoring wells in the intertidal zone and the HZ provide guidance to the process. As more data were obtained during the research project, the conceptual model was revised. 18       0 500 1000 1500 200005101520250 5000 10000Naphthalene (ug/L) Depth (m) Specific conductance (uS/cm) SCNaphthalene21-Aug-2014 MW14-02 0 500 1000 1500 200005101520250 5000 10000Naphthalene (ug/L) Depth (m) Specific conductance (uS/cm) SCNaphthalene0 1000 200005101520250 5000 10000Naphthalene (ug/L) Depth (m) Specific conductance (uS/cm) SCNaphthalene0 500 1000 1500 200005101520250 5000 10000Naphthalene (ug/L) Depth (m) Specific conductance (uS/cm) SCNaphthalene0 500 1000 1500 200005101520250 5000 10000Naphthalene (ug/L) Depth (m) Specific conductance (uS/cm) SCNaphthalene21-Aug-2014 MW14-03 25-Aug-2014 MW14-04 25-Aug-2014 MW14-05 25-Aug-2014 MW14-06 Figure 2.3 – Vertical profiles for naphthalene concentrations and specific conductance in five intertidal zone monitoring wells are shown, along with a map of their relative position near the high water mark. Peak naphthalene concentrations are found at a depth of approximately 16 m, below the halocline. Data collected by Golder Associates Ltd. 19      Figure 2.4 – Conceptual site model of groundwater discharging to the Fraser River. Fresh and saline groundwater are denoted by blue and red, respectively. Groundwater dilution occurs as it mixes with river water in the HZ. 20  2.4 Methodology  To characterize groundwater discharge to the Fraser, sampling transects were completed in the HZ perpendicular to the shoreline. Findings from this work were then supplemented by observations from multi-level monitoring wells installed in the intertidal zone of the riverbank. Groundwater from the HZ was sampled on five multi-day excursions between July 2014 and January 2015, while groundwater from two intertidal monitoring wells was sampled in August 2015. The two wells selected for examination of inorganic chemistry were MW14-02 and MW14-06. They were equidistant from the shoreline's high water mark (HWM) and were thus expected to yield similar vertical salinity gradients. The distance separating the wells was approximately 30 m.  Field work on the Fraser River was completed on the "Kraken", UBC's 22-foot aluminum research vessel. GPS was used to verify positioning on the river, which was further confirmed through triangulation with range-finders and shoreline markers. Once a sampling location was confirmed, dual anchors were set with the bow of the vessel facing into the current. During ebbing tides, the boat faced upstream, and during flooding tides, the boat faced downstream. By incrementally varying the length of line in each anchor, transects were completed perpendicular to the shoreline (see Figure 2.5). Sampling locations were selected in a manner such that the geochemical variability of discharging groundwater could be spatially characterized. Where discharging groundwater shifted from fresh to saline, sampling locations were spaced closer together in an attempt to capture a finer resolution of the salinity gradient.  Groundwater samples were easiest to obtain from the most permeable sediments in the HZ. Sediment grab sampling transects indicated that from the HWM to approximately 70 m into the channel, the river bottom was composed predominantly of silt. Past this point, the sediment composition gradually shifted to fine sand, and then coarse sand, where large quantities of groundwater could be sampled at an efficient rate. The silt pinch-out point identified at 70 m from the HWM represented the top of the aquifer (Fraser River 21  Sands unit) where it intersected the river channel. At the River District site, groundwater entering the Fraser River discharged mainly through these sandy sediments, from the silt pinch-out to the centre of the channel. A comparable pattern of groundwater discharge was expected from the other side of the river.      Groundwater from the HZ was collected using the KIST® sampler (see Figure 2.6), developed by Peter Krahn in the Environment Canada Enforcement Division (Pacific and Yukon Region). After confirming that anchoring had yielded a stable position, the KIST® sampler was deployed from the boat deck and lowered to the riverbed. Once at the bottom, the sampler was manually hammered into the sandy sediments maintaining a vertical orientation. During periods of strong current, the sampler may have entered the HZ at a slight angle, estimated to be no more than 20 degrees from the normal. The sampler was inserted to a minimum depth of 1 m using the built-in slide hammer and attached cord. Depth of insertion was assessed when the percussion pad of the sampler was felt to reach the river bed, indicating that the extension coupling (Figure 2.6) was fully immersed. A cable length counter verified the amount of line released during hammering. current transect anchors Figure 2.5 – Method of transect completion across the Fraser River. Incremental variation in length of anchor line allows for lateral movement across channel. 22   Figure 2.6 – The KIST® sampler, designed by Peter Krahn of the Environment Canada Enforcement Division, Pacific and Yukon Region.   Once inserted, a Geotech peristaltic pump was turned on to start the flow of groundwater from the HZ up to the boat. Groundwater collected through the sampler's screened tip was extracted through HDPE tubing up to the boat deck. A flow through cell was fitted with a Thermo Orion 250A+ pH meter calibrated daily to 3-points (pH 4, 7, and 10) and a Thermo Orion 115 conductivity meter capable of automatic temperature correction. Electrical conductivity data were thus automatically converted into values of specific conductance (SC), with units of microSiemens per centimetre (µS/cm). A secondary conductivity probe (TetraCon 325 with Cond 3400i unit) was also used to verify the primary probe's accuracy. Once a minimum of three tube volumes had been extracted and pH/SC values were stable, sampling began. Groundwater was sampled directly from the supply line with a 60mL syringe and filtered through Millex-HV 25mm-diameter 0.45µm pore size cellulose acetate lockable syringe filters. Sample 23  containers were triple-rinsed and filled with no head space. Samples for dissolved metal analysis were acidified with HNO3 to pH 2 or lower for preservation, while samples for anion analysis remained untreated. Samples were kept on ice, and then refrigerated as soon as possible. Analyses were completed by AGAT Laboratories in Burnaby, BC. Cation concentrations were determined by ICP-OES while anion concentrations were analyzed by ion chromatography.   Alkalinity of each sample was determined with the Gran titration method using dilute sulfuric acid. When titrations could not be done on-board immediately after sampling, they were completed in the evening upon return from the field. Because the majority of samples had high concentrations of ferrous iron, proton generation associated with iron oxidation and concurrent decline in alkalinity were expected. To assess deviation from true alkalinity in the hours after sampling, a titration time-series experiment was conducted on samples from different excursions (see Figure 2.7). Two samples and one duplicate were immediately transferred into separate containers during sampling, all of which were to be titrated at a later date. When samples were titrated within 6 hours of sampling, the apparent alkalinity was found to deviate by no more than 5%. Samples with higher iron content experienced greater deviation from true alkalinity. 24   Figure 2.7 – The effect of delayed titration on apparent alkalinity is shown for hyporheic groundwater samples rich in Fe2+. Although Fe concentrations in the legend pertain to total dissolved iron, Fe2+ is expected to be the predominant species. Initial calculated alkalinity values are 9.17 and 7.93 meq/L for D05 and E13, respectively. Sample exposure to oxygen is expected to occur mainly during filtering and bottling, but low-level diffusion of oxygen through the HDPE container wall is possible over longer periods.  2.5 Results and Discussion  Monitoring Well Conductivity  Figure 2.8 shows the vertical distribution of specific conductance (SC) measured in wells MW14-02 and MW14-06. Fresh, low-SC groundwater exists down to a depth of 10 m. Below this, a rapid shift in salinity (considered here a halocline) occurs, with SC values stabilizing at a depth of 13 m. From this point downward, the groundwater is observed to be saline. At the bottom of the aquifer, a slight decrease in SC is apparent, and is hypothesized to result from slow, upward seepage of fresh groundwater from the underlying aquitard. Similar observations were made by Jia at her site.  99% 90% 83% 100% 98% 93% 84% 100% 96% 91% 81% 53% 40%50%60%70%80%90%100%0.01 0.1 1 10 100Alkalinity relative to earliest measurement Days passed since sampling Effect of Delayed Titration on Apparent Alkalinity D05:          [Fe] = 32.4 mg/LD05-Dupl: [Fe] = 32.4 mg/LE13:          [Fe] = 138 mg/L25   Figure 2.8 – Specific conductance (SC) of groundwater below the intertidal zone is shown with measurements taken from multi-level monitoring wells. At depths of up to 10 m, groundwater is fresh. A rapid shift in SC is evident between depths of 10 and 13 m, and is referred to as a "halocline". Groundwater at depths of 13 m and below is saline. A slight decrease in SC for the deepest measurements is due to upward seepage of fresh groundwater from the underlying aquitard.  Hyporheic Zone Conductivity  To help visualize salinity gradients in the discharge zone, values of SC for sampling locations in the HZ are plotted in Figure 2.9. Most noticeable is the increase in SC towards the centre of the channel. Apart from a few HZ sampling locations close to shore with unexpectedly high SC attributed to silty sediments, groundwater samples collected from around and past the silt pinch-out have low SC values. These low SC values are identified in the HZ at an approximate distance of 70 to 85 m from the HWM. They represent a continuous, discharging band of fresh groundwater from the top portion of the aquifer.   Between approximately 85 and 100 m from the HWM, a rapid increase in SC is noted in HZ samples. SC is found to increase by up to an order of magnitude as the nature of discharging groundwater shifts from fresh to saline. This continuous band is where the onshore groundwater halocline intersects the river bed. 05101520250 2000 4000 6000 8000 10000 12000Depth (m) Specific Conductance (µS/cm) Groundwater Conductivity Below Intertidal Zone MW14-02MW14-0626     Figure 2.9 – Specific conductance of all HZ groundwater samples is shown. Regular river flow is from right to left, with the shoreline located just north of the frame. The increase in specific conductance with distance from the shoreline supports the conceptual model. Data were collected between July 2014 and January 2015 and thus represents a time-averaged distribution. 27   At distances beyond 100 m from the HWM, the SC of HZ samples is 5000 µS/cm and above. All HZ samples from beyond this point have similar SC, with a maximum of 7060 µS/cm. This suggests that the origin of all discharging saline groundwater in the central channel is the same, and comes from deeper portions of the lower aquifer. The farthest sample taken from the HWM is at a distance of 122 m, and has an SC value of 5890 µS/cm. At the River District site, the total width of the Fraser River North Arm is approximately 300 m.  Since the SC of HZ samples from the saline groundwater discharge zone is lower than the SC of deep groundwater, dilution by low-SC river water is evident. To verify that SC is a valid indicator for dilution, all samples from the monitoring wells and the HZ are plotted to compare conductivity with chloride and sodium concentrations (see Figures 2.10 and 2.11). The resulting correlation coefficients are very high, with R2 values ranging from 0.954 to 0.983. Chloride, which acts as a conservative tracer, correlates highest with SC.    Figure 2.10 – Effect of chloride (left) and sodium (right) concentrations on specific conductance are shown for all samples taken from the intertidal monitoring wells. Correlation with chloride is strongest.     R² = 0.976 020004000600080001000012000140000 1000 2000 3000 4000Specific conductance (µS/cm) [Cl] (aq) (mg/L)   MW14-02  MW14-06R² = 0.954 020004000600080001000012000140000 500 1000 1500 2000 2500[Na] (aq) (mg/L)   MW14-02  MW14-06Monitoring Wells 28     Figure 2.11 – Effect of chloride (left) and sodium (right) concentrations on specific conductance are shown for all samples taken from the hyporheic zone. Correlation with chloride is strongest.  Water Type Characterization  The major ion chemistry of the samples from the intertidal monitoring wells and the HZ can be compared to confirm the understanding of groundwater flow paths and discharge to the Fraser. Samples are plotted on a Piper plot (see Figure 2.12). This type of diagram plots samples based on the relative charge equivalent concentrations of major ions – one triangle for cations (Ca2+, Na+, Mg2+), and one for anions (Cl- , HCO3-, SO42-). The position of a sample in both triangles then determines where it plots overall on the upper diamond. Because samples are plotted based on relative charge equivalence, two samples with different absolute concentrations may still have similar relative concentrations and thus be plotted in the same spot. Finally, the placement of a sample in a certain area of the diamond can be used to describe its water type. For example, a fresh groundwater sample with a charge equivalence concentration dominated by Ca2+ and HCO3- would plot on the left-most side of the diamond, and would be termed a Ca-HCO3 water type (Appelo and Postma 2005, p. 244) [37]. If more than one ion makes up the dominant cationic or anionic charge, that ion can be included in the water type name, in descending order of proportional ionic charge (e.g. Ca·Mg-HCO3).  R² = 0.983 0100020003000400050006000700080000 500 1000 1500 2000Specific conductance (µS/cm) [Cl] (aq) (mg/L) R² = 0.9635 0100020003000400050006000700080000 200 400 600 800 1000[Na] (aq) (mg/L) Hyporheic Zone 29     Figure 2.12 – A Piper plot, or trilinear diagram, is used to plot the geochemistry of all water samples. Samples are plotted based on the relative charge equivalent concentrations of major ions. In general, the water type of deep monitoring well samples is similar to HZ samples farthest from the HWM. Similarly, shallow monitoring well samples are similar to HZ samples closest to the HWM. 30  Figure 2.12 shows that in general, the water type of deep monitoring well samples (depths greater than 13 m) are similar to HZ samples farthest from the HWM. Similarly, shallow samples from the monitoring well (depths up to 10 m) are plotted in the same region as HZ samples closest to the HWM. This suggests that shallow groundwater discharges into the river between 70 and 85 m from the HWM, while deep groundwater discharges at distances greater than 100 m from the HWM.   The major endmembers in this system are fresh groundwater, saline groundwater, and river water. In Figure 2.12, fresh groundwater is denoted by triangles with depth labels less than 10 m, saline groundwater is denoted by triangles with depth labels greater than 13 m, and river water is represented by an 'X'. Many samples plot between these major groupings, indicating mixing. When conservative mixing occurs (i.e. non-reactive mixing), the resultant point is found along a straight line between its two endmembers.  In Figure 2.12, however, there are indications of non-conservative mixing. One example is the location of HZ samples taken from the saline groundwater discharge zone (i.e. distances greater than 105 m from the HWM, denoted by red circles). These points do not plot in exactly the same position as their source groundwater samples, but rather show enrichment along the "Ca + Mg" axis (depths greater than 13 m, denoted by triangles). This suggests that non-conservative mixing is occurring as saline groundwater migrates into the HZ.  Another example of non-conservative mixing is the location of the "11.9 m" sample taken from MW14-02 – it also shows enrichment in Ca and Mg compared to the sample from 13.7 m. Here, non-conservative effects are likely a result of transverse mixing between fresh and saline groundwater along the horizontal flow path towards the river.  31  Cation exchange reactions provide the most likely explanation for the non-conservative effects observed. These reactions are common at the interface separating fresh and saline groundwater. When saline groundwater migrates into an area previously occupied by fresh water, the process is referred to as intrusion. The reverse process is called freshening. Together, the two serve as dominant geochemical processes in coastal aquifers (Appelo and Postma 2005, p. 242) [37]. From an anthropogenic perspective, intrusion is common where over-extraction of fresh groundwater near the coast causes ingression of saline groundwater.  The compositions of seawater and river water during the low-flow season are shown for reference in Figure 2.12 (see legend). Conservative mixing between these two compositions is indicated by a dotted line. This line represents the expected charge equivalent proportion of Ca2+ + Mg2+ and Na+ for a sample mixed with no reaction. When a sample plots far away from this line, it indicates that non-conservative reactions, such as cation exchange, are occurring. A sample plotted downward from this line indicates freshening, as there is more Na+ in solution than expected from conservative mixing. This is most likely a function of exchange sites on sediment surfaces equilibrating with the ions in solution adjacent to them. So if freshwater moves towards sediments previously equilibrated with saline water, the sediments will uptake Ca2+and release Na+ until equilibrium is re-established (see Reaction 2.1). A sample plotted upward from the dotted line in Figure 2.12 represents intrusion, for the contrary reason. A sample indicative of freshening or intrusion can be traced back to an estimated conservative mixing ratio by travelling up or down along the "Ca + Mg" and "Na" axes (Appelo and Postma 2005, p. 246) [37]. This is indicated by the straight-lined arrows in Figure 2.12.  ½ Ca2+(aq) + Na-X  ⇌  ½ Ca-X2 + Na+(aq)  In Figure 2.12, the shallowest ports (5.9 to 10 m) demonstrate an upward freshening sequence.  (2.1) 32  This suggests that groundwater migrating along shallower flow paths (i.e. 5.9 m) on its route to the river experiences more freshening than groundwater migrating along slightly deeper flow paths (i.e. 9.2 m). This may be a result of recharge from precipitation, or induced river recharge during high tides. Similarly, the deepest ports (17.3 to 19.9 m) show evidence of freshening, as they plot slightly downward from the other ports (15.5, 15.6 m) on the "Ca + Mg" and "Na" axes. This is attributed to the fact that the underlying aquitard is slowly releasing fresh groundwater into the deepest portion of the aquifer. Similar findings have been found by Neilson-Welch & Smith (2001) and Jia (2015).  Cation exchange affecting geochemistry in the halocline is further exemplified in Figure 2.13. Although Na+ and Mg2+ concentrations increase in unison with the halocline, and remain very high at the bottom of the aquifer, Ca2+ concentrations peak at the halocline, but then decrease significantly with depth. Similar findings are described by Jia (2015) at a nearby site. This is a result of saline groundwater interacting with overlying fresh groundwater through transverse mixing along the horizontal flow path towards the river. Pumping of groundwater at the River District site may also be contributing to transverse mixing. At this depth, cation exchange reactions are exemplified by the reverse of Reaction 2.1.   Where saline groundwater intrudes an originally fresh aquifer, "Ca-Cl2" water types are often created (Appelo and Postma 2005, p. 244) [37]. This is the case for the water sample collected from a depth of 11.9 m in MW14-02, which plots in the upper part of the diamond in Figure 2.12. The release of Ca2+ from intrusion-related exchange reactions is indeed more evident in MW14-02 than MW14-06, as peak concentrations are 42% higher (see Figure 2.13). When high Ca2+ concentrations are produced from exchange reactions, this can sometimes generate calcite precipitation [65]. The resulting reaction produces acidity (see Reaction 2.2). This process is possibly occurring below the intertidal zone at the River District site. Figure 2.14 shows that at the same depth of peak Ca2+ concentration in monitoring well MW14-02, there is a noticeable drop in pH.  Ca2+(aq) + HCO3-(aq) + OH-(aq)  ⇌  CaCO3 (S) + H2O (2.2) 33   Figure 2.13 – Dissolved Mg2+, Ca2+, and Na+ concentrations are shown for monitoring well samples from MW14-02 and MW14-06, which are 30 m apart, and equidistant from the HWM. Peak Ca2+ concentrations occur at depths 11.9 and 11.4 m, respectively, coinciding with the middle of the halocline. With increasing depth, Ca2+ concentrations rise in unison with Na+ and Mg2+, but then drop off below the halocline.   Figure 2.14 – pH measurements are shown from multi-level monitoring wells MW14-02 and MW14-06. The drop in pH at a depth of 11.9 m in MW14-02, shown by the arrow, is possibly a result of acidity generation due to calcite precipitation [65]. High Ca2+ concentrations are a result of cation exchange reactions associated with "intrusion", or fresh-saline groundwater interaction at the halocline. 051015201 10 100 1000Depth (m) mg/L Cation Concentrations Below Intertidal Zone MW14-02 MW14-0651015206 6.5 7 7.5 8 8.5Depth (m) pH MW14-02MW14-06Mg2+ Ca2+ Na+ 34  Conceptual Model   Analysis of data from the intertidal monitoring wells and the hyporheic zone support the conceptual model through two main lines of evidence – trends in specific conductance and water type characterization. Figure 2.15 compares the normalized specific conductance (NSC) of all samples, together with the water type as determined by the plotted position on the Piper plot in Figure 2.12. NSC is calculated by feature scaling the HZ and monitoring well datasets separately. Feature scaling is completed by subtracting the minimum value in a dataset from a given datum, and then dividing by the range of the dataset. The dividend then represents NSC, and can take a value from 0 to 1 (see Equation 2.1). Values used for this operation are found in Table 2.1.  𝑁𝑁𝑁 =  𝑁𝑁 −  𝑚𝑚𝑚𝑚𝑚𝑚𝑚𝑟𝑟𝑚𝑟𝑟  Table 2.1 – Values used to calculate NSC are shown below, in units of microSiemens per centimeter (µS/cm).  HZ Dataset Monitoring Well Dataset Minimum 148 437 Maximum 7060 11 840 Range 6912 11 403   In Figure 2.15, groupings A and C represent shallow and deep samples from monitoring wells, respectively. Groupings B and D represent near-shore and central-channel HZ samples, respectively. Figure 2.15 shows that A and B have low NSC values, and that the majority of samples are of the CaMg-HCO3 or CaMgNa-HCO3 types, suggesting they are on the same flow path. Likewise, C and D have high NSC values, and most samples are of the Na-Cl water type, suggesting that they are on another flow path. This supports the idea that groundwater flow towards the river is relatively horizontal, with low transverse vertical dispersivity. As it nears the discharge zone, groundwater turns upward and mixes with river water in the HZ.  (2.1) 35   Figure 2.15 – Normalized specific conductance for HZ and intertidal zone (IZ) samples is plotted together with water type (top). A labeled conceptual model is also displayed to indicate location of samples by grouping letter (below). Shallow groundwater (A) flows to the near-shore HZ (B) while deep groundwater (C) flows to the central-channel HZ (D). Dashed lines with arrows represent flow paths. 36  The highest SC value (7060 µS/cm) from the HZ is only 60% of the highest SC value from the monitoring wells (11840 µS/cm) – this is demonstrative of the dilution which groundwater experiences as it flows through the HZ (i.e. at the end of flow path C-D in Figure 2.15). Using NSC to compare the range of SC values between the datasets is valid assuming that dilution of discharging groundwater is uniform at different horizontal locations on the river bottom. Since the depth of the Fraser River North Arm is relatively constant for the majority of its width, this assumption holds. If discharge rates and dilution were to vary as a result of heterogeneities or dune formation, this assumption would be less valid.  Manganese Reduction in the Aquifer  Manganese concentrations from the monitoring wells are plotted in Figure 2.16. Most evident is the peak in Mn at the same depth as the halocline. Peak Mn concentrations for wells MW14-02 and MW14-06 are 3.1 and 3.95 mg/L, at depths of 13.7 and 11.4 m, respectively. In comparison, Mn concentrations at the shallowest ports are in the range of 1 mg/L, and they fall below 1 mg/L at the bottom of the aquifer. Similar Mn peaks are identified by Jia (2015). Jia hypothesizes that these peaks are a result of manganese reduction, driven by the oxidation of organic matter (OM). Possible energy sources for microbes at this location include OM from silt lenses and redox microzones, or advection of dissolved organic contaminants from the up-gradient direction. Since Jia finds that Mn reduction is occurring at a site which is not contaminated, however, the process should persist independently of dissolved anthropogenic contaminants. It is also possible that chemolithoautotrophic metabolisms, such as iron, sulfur, or methane oxidation, are driving Mn reduction.  37   Figure 2.16 – Manganese concentrations for the intertidal zone monitoring wells are shown. Peak concentrations in wells MW14-02 and MW14-06 are 3.10 and 3.95 mg/L at depths of 13.7 and 11.4 m, respectively. These Mn peaks occur at the same depth as the halocline.  In comparing Figures 2.13 and 2.16, it is evident that Ca and Mn peaks coincide with the halocline. These peaks are an outcome of fresh and saline groundwater interaction, but are likely caused by separate mechanisms – Ca by cation exchange, and Mn by reduction. As shown in Figure 2.17, Ca and Mn concentrations show strong linear association, indicating that the extent of both Ca exchange reactions and Mn reduction are correlated. This is observed in the monitoring wells where fresh and saline groundwater mix, and also in the HZ, where groundwater mixes with river water (Figure 2.17).   One possible explanation for this strong correlation is that exchange reactions and Mn reduction are limited equally by the extent of transverse mixing. Another possibility, which may not be reliant on mixing, is that Ca release occurs in parallel with Mn reduction. For example, the same process that reduces Mn may also be responsible for releasing CO2, which then causes an increase in the concentration of carbonic acid, ultimately dissolving Ca minerals.   051015200 1 2 3 4Depth (m) [Mn](aq) (mg/L) Manganese Concentrations in Monitoring Wells MW14-02MW14-0638   Figure 2.17 – Correlation between Mn and Ca concentrations is shown for all samples from the hyporheic zone (left) and the monitoring wells (right). Although aqueous Mn and Ca are generated via different mechanisms, they are both dependent on mixing. Dilution is also expected to play a role in the HZ.  In many environments, it is conceivable that a strong correlation in the concentration of two elements is simply a result of conservative dilution. This is unlikely the case here, however, since Ca and Mn concentrations both increase with distance from the HWM in the HZ (see Figure 2.18). Their concentrations do not peak where the halocline intersects the discharge zone (85-100 m from HWM), as would be expected if monitoring well data (i.e. Figures 2.13 and 2.16) were extrapolated out to the HZ using the conceptual model. Instead, Ca and Mn concentrations peak at the greatest distance from the HWM, suggesting that conservative dilution is not the major control of their correlation. 0501001502002503003504000 10 20 30 40[Ca] (mg/L) [Mn](aq) (mg/L) Hyporheic Zone 0501001502002503003504004505000 2 4[Ca] (mg/L) [Mn](aq) (mg/L) Monitoring Wells MW14-02MW14-0639   Figure 2.18 – Ca and Mn concentrations as measured in the hyporheic zone are plotted against their location relative to the high water mark (HWM). With increasing distance, concentrations increase.   Iron Reduction in the Aquifer  Data show that Fe reduction in the halocline is coincident with Mn reduction. Figure 2.19 shows that peak Fe concentrations for MW14-02 and MW14-06 are 45.1 and 48.0 mg/L at depths of 11.9 and 9.2 m, respectively. These values are lower than those identified by Jia (2015), which were in the range of 300 mg/L. These peaks take a similar shape as the Mn peaks, but the Fe peaks are vertically located just above the Mn peaks in both monitoring wells. At these locations, it is probable that Fe reduction is being driven by a similar mechanism as Mn reduction. Jia (2015) also finds Fe peaks to be coincident with Mn peaks at her site.   In well MW14-02, a secondary Fe peak occurs above the primary peak. Between depths of 7 and 9 m, Fe concentrations are in the range of 36 mg/L. Since this peak is found entirely within fresh groundwater at the top of the aquifer, Fe generation here is likely a result of a mixing-independent mechanism. Nearby at well MW14-03, dissolved naphthalene concentrations are measured to be on the order of 1 mg/L, exactly one year prior, at a depth of 5.2 m (Figure 2.3). Although naphthalene concentrations are found to be 05010015020025030035060 80 100 120[Ca] (mg/L) Distance from HWM (m) 051015202530354060 80 100 120[Mn] (mg/L) Distance from HWM (m) 0 0 40  negligible at shallow depths in MW14-02, it is possible that naphthalene degradation is driving reduction and mobilization of Fe up-gradient at MW14-03. Supporting this idea is the fact Roschinski (2007) finds strong correlation between ferrous iron and indane concentrations in an aquifer at a similar site. This suggests that naphthalene degradation may be driven by Fe reduction.   Figure 2.19 – Iron concentrations below the intertidal zone monitoring wells are shown. Peak concentrations in wells MW14-02 and MW14-06 are 45.1 and 48.0 mg/L at depths of 11.9 and 9.2 m, respectively. These Fe peaks occur just above the Mn peaks – they are approximately 2 m shallower. A secondary Fe peak for well MW14-02 is noticeable at depths of 7-9 m.  Iron and Manganese Reduction in the Hyporheic Zone  As groundwater discharges into the Fraser, it is significantly diluted. For essentially all elements measured, concentrations in the HZ are lower than in the monitoring wells. For Fe and Mn, however, concentrations in the HZ are actually higher. The maximum concentration of Fe in the HZ is 138 mg/L (Figure 2.20), which is about 3 times greater than the peak Fe of 48 mg/L in the monitoring wells (Figure 051015200 10 20 30 40 50Depth (m) [Fe] (aq) (mg/L) Iron Concentrations in Monitoring Wells MW14-02MW14-0641  2.19). For Mn, the highest measured concentration increases from 4.0 mg/L (Figure 2.16) in the monitoring wells to 34.7 mg/L in the HZ (Figure 2.20) – an increase factor of about 9.    Figure 2.20 – Mn and Fe concentrations from all samples in the HZ are shown. Although Mn concentrations increase with distance from the high water mark (HWM) for Mn (left), no noticeable trend exists for Fe (right).  This apparent increase in Fe and Mn despite dilution is likely a result of either ion-exchange effects, or biogeochemical activity. Since the supply of OM from the Fraser River to the HZ may be high, it is possible that Fe and Mn undergo reductive dissolution via OM oxidation in the HZ, or along the flow path to the HZ. Another potential explanation supported by previous research is that H2S oxidation is driving reduction of Fe and Mn in the aquifer and/or the HZ [66, 67]. Herszage and dos Santos Afonso (2003) state that in an anoxic environment, Fe or Mn oxides can oxidize H2S, either biotically or abiotically. Furthermore, Yao and Millero (1996) demonstrate that in marine environments, rates of H2S oxidation increase with decreasing ionic strength. As saline groundwater becomes increasingly diluted by river water in the HZ, ionic strength is reduced, thus potentially making the anaerobic portion of the HZ an ideal environment for H2S oxidation.  051015202530354060 80 100 120[Mn] (mg/L) Distance from HWM (m) 0 02040608010012014016060 80 100 120[Fe] (mg/L) Distance from HWM (m) 0 42  Although H2S oxidation may provide a potential explanation for Fe and Mn reduction, H2S is generally a product of sulfate reduction driven by OM oxidation in the first place (Appelo and Postma 2005, p. 247). Additionally, some research invites the possibility that H2S concentrations can be sustained through sulfate reduction driven by anaerobic oxidation of CH4 [68-70]. Although H2S oxidation may be contributing to some Fe and Mn reduction, it is highly unlikely that H2S oxidation alone is generating all of the reduced Fe and Mn in the HZ. In all likelihood, high Fe and Mn concentrations in the HZ are driven by a combination of interdependent geochemical processes supported by the combined oxidation of OM, H2S, and CH4.   In the HZ, there is no correlation between Fe concentrations and Mn or Ca. Additionally, there is no noticeable trend in the HZ between Fe and distance from the HWM (see Figure 2.20). This implies that Fe mobilization in the HZ is controlled by factors which are different from those governing the mobilization of Mn.  2.6 Conclusion  This chapter presents data on groundwater sampled from the HZ between July 2014 and January 2015, and from intertidal monitoring wells in August 2015. Specific conductance is found to increase with depth in the monitoring wells, and with distance from the HWM in the HZ. A conceptual model is developed to assist in the understanding of groundwater discharge patterns. Data reveal that fresh groundwater discharges to the HZ at distances between 70 and 85 m from the HWM, while saline groundwater discharges at distances greater than 100 m. A halocline separates the fresh and saline bodies of groundwater, and its intersection with the HZ is estimated to be between 85 and 100 m from the HWM. Water type characterizations for both fresh and saline groundwater support this understanding. Furthermore, conductivity and solute data have the potential to be used for numerical model benchmarking purposes by hydrogeologists working at the River District site.  43   This chapter provides an improved understanding of flow and geochemistry to better define principal groundwater discharge zones. There exist potential plans to shut off the pumping wells and allow the natural groundwater flow to re-establish itself, as part of a monitored natural attenuation strategy. If this is to proceed, monitoring may increase in the HZ, and predictable contaminant discharge points will provide direction to this process. These predictions are made by comparing the depth of peak dissolved contaminant concentrations (Figure 2.3) with NSC (Figure 2.15). For example, if peak contaminant concentrations in the intertidal monitoring wells are found at a depth of 16 m, the corresponding NSC value from Figure 2.15 (trend line at top-right) would be 0.85. When interpolated on the HZ line of best fit (trend line at top-left of Figure 2.15), this would correspond with a distance of 105-110 m from the HWM. A conservative tracer test carried out from the monitoring wells with an ensuing HZ survey could confirm this assessment.  Cation exchange effects are generated by transverse mixing along the halocline, and are responsible for high dissolved Ca concentrations. Peaks in Fe and Mn are also found in the halocline, although Fe peaks occur at slightly shallower depths than Mn peaks. The reduction of Fe and Mn are hypothesized to be driven by the oxidation of organic matter or reduced dissolved gases, which themselves are fundamentally generated by organic matter. The Fe and Mn peaks may also be an additional effect of cation exchange. A secondary peak in Fe concentrations is found in shallow, fresh groundwater from MW14-02. This peak may be a result of nearby naphthalene degradation. From the monitoring wells to the HZ, Fe and Mn concentrations are found to increase. This observation is believed to be a result of biogeochemical reactions occurring either along the flow path, or in the HZ. Microbial oxidation of OM, H2S, and CH4 serve as potential explanations for these observations.   44  Chapter 3 – Design and Development of a Sediment-Freezing Sampler  3.1 Previous Approaches  Previous efforts at collecting sediment cores from below the Fraser River have been successful. A freeze-shoe corer design by Bianchin et al. (2015) modified a drive-point piston sampler by Starr and Ingleton (1992) by adding a sample-freezing drive shoe and driving it into the sediments with a pneumatic hammering system [71-73]. This design was effectively a traditional core sampler with the added ability of freezing the lowest 10 cm of the core, thus securing the remaining overlying sample and sealing it from the atmosphere [73]. One drawback of this method was that it was less effective in environments with soft sediments, and this provided motivation to develop a new sampling approach.  Roschinski (2007) developed a hollow freeze-pipe sampler which used liquid N2 (boiling point -196°C) instead of CO2, (boiling point -78°C) with some similarities to soil-sampling equipment developed by Hofmann et al. (2000). Using N2 increased freezing speed and made the sampling process more efficient [34, 74]. Roschinski's sampler was fitted to the bottom of a sequence of drill rods, guided through a steel bracket on the side of the work vessel, and advanced through the sediments using a pneumatic hammer (see Figure 3.1). Liquid N2 was injected at 230 psi and it expanded within the freeze-pipe chamber. A 60 cm-long core was frozen in about 25 minutes. Sampler retrieval was estimated to take 10 minutes, as the drill rods connected to the sampler had to be consecutively unthreaded and removed.  45      An efficient and commonly-used design for freeze-sampling sediments involves inserting a N2 or CO2 distribution pipe with evenly-spaced small orifices into a pointed hollow pipe, such that a "nested pipe" configuration is created [75-78]. In every instance that this design is used, the objective is to sample a creek or stream. These water bodies are much smaller and shallower than the Fraser River, and can be accessed only by foot. In the case of Moser et al. (2003), freeze-coring was done to examine the HZ of the Columbia River at the Hanford site, but these samples were taken from shore-accessible stations on the edge of the river [79]. To the best of our knowledge, this type of design has not been used to collect frozen sediment cores from larger bodies of water. Other manifestations of sediment-freezing samplers require pouring cryogens directly into the freeze pipe [80], or pre-loading the sampler with crushed dry ice and alcohol prior to deployment [81].    Figure 3.1 – Side-view diagram showing Roschinski's method. Adapted from Roschinski (2007). 46  3.2 Design Considerations  Motivation  The cost-efficiency and smaller scale of the nested pipe design are desirable for this investigation since samples are collected from UBC's small (22-foot) research boat. A few major challenges of implementing this design include ensuring that the sampler can be reliably operated at depths of 10-20 m below the water surface, in a river with significant current, and that it can be deployed on a motorized winch line. Selection of a sampling location and operation of the sampler are also challenging at these depths, so a design that allows for an underwater camera is considered advantageous.  Objectives  Our objective is to collect a representative sample of porewater and adjacent sediment that preserves structure and does not introduce contamination. Sediment freezing provides an ideal collection and preservation method for this objective. By freezing the sediment sample prior to removal, the chemical and microbiological activity of the system is effectively reduced to near zero, thus maintaining in-situ representativeness. As long as the sample remains frozen, DNA from microorganisms is preserved at a high quality. When unfrozen sediments are collected by conventional coring from their stable physical environment, significant deformation may occur. As a result, sub-samples cannot be reliably collected at small intervals. By freezing the sediments, however, sediment structure is preserved without vertical deformation, allowing for analysis of sediment depositional patterns and grain size distribution at a finer scale. Furthermore, coincident porewater and sediment can be collected, allowing for the analysis of the relationships between solid phases and adjacent solutes in the HZ, where geochemical gradients are hypothesized to be strong.   47  Constraints  A major concern associated with collecting a sediment core external to the sampling device is potential contamination during removal. After removing the frozen sample from the river sediments, the core is exposed to river water on its journey to the surface for up to 30 seconds. Additionally, there is the possibility of river water migrating to the interior of the core during insertion of the sampler. It is expected that these concerns can be controlled by ensuring both inner and outer surfaces of the frozen core are removed during subsampling. For this thesis, an additional small core is taken so that potential differences in porewater chemistry due to river water contamination can be assessed on the inner and outer surfaces.  A major constraint associated with sampling sediment and porewater in a tidally-affected system is the oscillatory nature of groundwater flow. In this environment, sediment freezing has to be done as quickly as possible to ensure that an accurate temporal snapshot of geochemical conditions is captured. If freezing takes too long, the continually growing freezing front will start to assimilate porewater compositions which differ from those at the innermost portions of the core. Thus, the faster the freezing process, the more representative the porewater in the sample.   Brine exclusion is another concern when freezing porewater for subsequent analysis. Brine exclusion is characterized by ion concentrations in an ice sample being lower than the actual source water prior to freezing. This occurs because dissolved ions of high concentrations may separate out from solution as water turns into a solid phase. The phenomenon becomes more apparent with an increasing amount of dissolved solutes. Toran et al. 2013 investigates brine exclusion during freeze coring and determines that, although it is difficult to quantify the extent of brine exclusion for a given sample, the issue is more apparent for larger core sizes [80]. The extent of brine exclusion is thought to increase non-linearly with 48  core size, and this is hypothesized to be a function of the decreasing radial freezing rate. So as core volume increases, the rate of freezing slows, and solutes are more prone to brine exclusion.  Important environmental concerns exist when using CO2 as a cryogen. When CO2 dissolves in water, carbonic acid forms (see Reactions 3.1 and 3.2), and this poses a hazard to aquatic organisms. To avoid acidification of the water column, CO2(g) must be expelled from the sampler to the atmosphere with hosing that has high flexibility, high durability, and a low temperature rating. CO2(g) ⇌  CO2(aq) CO2(aq) + H2O ⇌  H2CO3   3.3 Production and Testing  The design of the sediment-freezing sampler was finalized after a systematic review of field objectives and practical constraints. A sampler length of one metre was chosen based on previous conclusions on the extent of mixing in HZ. This sizing was thought to be adequate for capturing the sharp geochemical gradients occurring in the first metre of sediments. Stainless steel piping with an outer diameter of 48 mm (1.9 ") was used for the main freeze pipe. A detachable pointed tip was also machined out of stainless steel. To minimize leaks and withstand high operating pressures, all major components of the sampler were designed with national pipe thread tapered (NPT) connections.   For the research presented in this thesis, the sediment-freezing sampler was combined with components of the KIST sampler (see Figure 3.2). A coupling was machined to connect the upper portion of the freeze pipe to the KIST sampler's slide hammer, thus giving it a means of penetrating the sediment. Future tests will likely evaluate the idea of replacing the hammer with a sonic vibration attachment, which could reduce the effort required to collect a sample. (3.1) (3.2) 49       slide hammer cord winch line tube exit port slide hammer percussion pad additional weight slide hammer stem CO2 (liq.) feed line temperature probe CO2 expansion in freeze pipe underwater camera CO2 (gas) vent line sediment freezes to outside of pipe  Figure 3.2 – Diagram showing unification of sediment freezing sampler (bottom) and KIST sampler (top).  50  The CO2 distribution manifold inside of the freeze pipe was made out of copper tubing (see Figure 3.3). Six very small orifices were drilled onto the tubing with a #79 (0.37 mm) drill bit. These were evenly spaced, radially and lengthwise. Although the orifices were initially planned to be smaller, drill bit availability was limited. The 0.37 mm size was found to be adequate, however, for maintaining back pressure on the liquid CO2 feed line. An even distribution of the orifices was considered critical for ensuring uniform release of CO2 within the freeze pipe. Homogeneous cooling along the entire pipe surface was sought to generate an evenly-formed sediment core.  Large 50-litre CO2 tanks fitted with siphon tubes were supplied by Praxair in Vancouver, BC. Less than one cylinder was required to freeze a full-size core of approximately 4 cm thickness (i.e. outer radius – inner radius = 4 cm). Initial CO2 pressure in the tanks was approximately 830 psi at 25°C, but since field work was completed at ambient temperatures between 8 and 15°C, operating pressure was slightly lower. Liquid CO2 was injected through stainless steel-braided Teflon hosing provided by New-Line Hose and Fittings in Burnaby, BC. Following expansion in the freeze pipe, gaseous CO2 was expelled up to the atmosphere through Tygothane polyurethane hosing suited for temperatures down to -75°C. This hosing was obtained from McMaster-Carr in Elmhurst, IL. Swagelok tube fittings and adapters for hose connections were provided by Columbia Valve and Fitting Limited in Burnaby, BC.   The CO2 cylinder valve was opened for only 5 seconds out of every 2 minutes, corresponding to a duty cycle of about 4%. After shutting the valve, residual liquid CO2 in the feed line continued to enter the freeze pipe at a decreasing rate for nearly 2 minutes. Keeping a low duty cycle was critical for minimizing dry ice clogs and managing the lag in energy transfer between materials. Since the rates of energy transfer between sediment, steel, and the atmosphere in the freeze pipe are much lower than the CO2's potential rate of energy removal, a low duty cycle is more efficient.    51  1.0 m Figure 3.3 – Diagram showing the inner workings of the freeze pipe. Six small orifices are drilled into the copper distribution manifold. These are evenly distributed along the length of the manifold, as well as azimuthally. freeze pipe distribution manifold 52  Repeated testing demonstrated that the sampler was capable of obtaining a uniformly-frozen sample. The sampler was suspended in a column of water and thermocouples were fixed to the outside of the freeze pipe to monitor the rate of temperature drop (see Figure 3.4). Figure 3.5 shows that during the 40-minute freezing test, four different points of measurement on the freeze pipe experienced comparable cooling rates. The measurement locations were evenly-spaced, at 8, 34, 60, and 86 cm from the tip of the sampler. The CO2 valve was opened for the first time at t = 7 minutes. Frequent temperature fluctuations on the order of 2 minutes were believed to be a result of the on/off duty cycle. At t = 28 minutes, a slight warming event in the upper half of the sampler coincided with an amplified cooling event in the lower half of the sampler. This was hypothesized to be a result of dry ice sloughing off from the upper portions of the inner freeze pipe and accumulating at the bottom. Figure 3.6 shows the resulting ice core taken from the water column after 35 minutes of freezing.                 Figure 3.4 – The laboratory setup for sampler freezing tests is shown above. Tests were conducted in a water column inside of a tall PVC pipe sealed at the bottom. Thermocouples fixed to the outside surface of the freeze pipe provided live temperature readings which were recorded with a data logger. 53           data logger thermocouples x = 8cm x = 60cm x = 86cm -40-30-20-100100 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34 36 38 40 42Temperature (°C) Time (minutes) 8cm34cm60cm86cmFigure 3.5 – Temperature data from a freezing test are shown above. Evenly-spaced thermocouples along the freeze pipe surface demonstrate uniform cooling. The CO2 valve is opened for the first time at t = 7 minutes, and is kept open for only 5 seconds out every 2 minutes. Deviances starting at t = 28 minutes are believed to be a result of dry ice sloughing off from the upper portions of the inner freeze pipe and accumulating at the bottom. Figure 3.6 – The resulting ice core from a laboratory freezing test is shown above. The maximum core circumference of 12" occurs the bottom, while the minimum core circumference of 10" occurs at the top. Midpoint circumference is 11.5". 54  3.4 Sampling Methodology  Core sampling was conducted near high tide on UBC's 22-foot aluminum work boat in November 2015. The freeze sampler was lowered slowly to the bottom of the river using the vessel's winch line. While maintaining tension in the winch line to achieve a near-vertical orientation, the sampler was gradually hammered into the hyporheic sediments using the slide hammer. A live feed of the hammering at the sediment surface was provided by a high-luminosity flashlight and a GoPro camera in underwater housings (see Figure 3.7). The camera was linked to a monitor onboard via HDMI cable and powered with a USB cable.     When the sampler was fully inserted, freezing began. Liquid CO2 was injected into the sampler, where it was forced through orifices on the distribution manifold. Expanding CO2 was propelled onto the inner Figure 3.8 – After retrieving the sampler, the freeze pipe is immediately unscrewed from the rest of the assembly. Figure 3.7 – Above, a sequence of video frames shows progress of sampler insertion into the riverbed during hammering. 55  walls of the freeze pipe, forming a thin layer of dry ice. CO2 sublimating back into the gas phase was vented to the surface through polyurethane tubing. The freezing process was monitored until sufficient core diameter was achieved. Core diameter was estimated using a temperature probe at a fixed radius from the freeze pipe that was wired to a data logger and laptop on board, thus providing live temperature readings. As the probe temperature approached 0°C, CO2 injection was discontinued and the sampler was removed. Upward hammering was initially used as a removal method, but was later deemed ineffective as the freeze pipe was unable to maintain full "grip" on the sediment core during upward acceleration. It was noted that a smaller upward force, produced over a longer time period, was more effective in removing the frozen sediment core. This was accomplished by applying continual tension with a high-power winch, accompanied by a slight rocking of the boat to yield small, repetitive increases in the applied force.   Once elevated to the surface, the sampler was placed on the boat deck. The freeze pipe and attached sediment core were unscrewed from the rest of the assembly using pipe wrenches (see Figure 3.8). Figure 3.9 shows the size and uniformity of the core, with notable sediment deposition sequences in the lower half. The outer core surface was immediately covered with plastic wrap (Figure 3.10) and placed in a 6" PVC storage tube seated within a custom dry-ice cooler at a temperature of -35°C (Figure 3.11). With the freeze pipe still open, boiling water was carefully poured into the pipe to promote melting of the inner core surface off from the outer pipe wall. Once free, the freeze pipe slid upwards and out of the sediment core. The storage tube was then purged with inert N2 gas and sealed shut. Once off the boat, the storage tube was kept in a -20°C freezer until sectioning. Sectioning and subsampling of the core are outlined in Chapter 4. 56   Figure 3.9 – A full view of the sediment core is shown above. Total length is approximately 95 cm. Silt lenses and woody debris in the lower half of the core are shown in the inset.     57                     Figure 3.11 (left): The plastic-wrapped core is carefully placed into the dry ice cooler with a temperature of  -35°C. With the top of the freeze pipe still open, boiling water is cautiously poured inside. After a few minutes of gentle rotation and lifting, the freeze pipe is withdrawn from the core.  Figure 3.10 (right): After being removed from the rest of the sampler, the core is covered in thin plastic  wrap.  58  3.5 Assessing Contamination from River Water  To assess potential contamination of the sediment core with river water, a smaller secondary core was sampled from the HZ. This core was only 14 cm long, consisting of sediments from 16 cm to 30 cm below the riverbed, for a midpoint depth of 23 cm. Instead of examining porewater geochemistry trends with depth, however, a radial analysis was completed. This involved dividing the core into five concentric layers (see Figure 3.12).  Successively smaller circular forms were held on top of the core to guide the scouring of each layer with a steel putty knife. Although the inner and outer surfaces of the primary core (Chapter 4) were removed prior to analysis, the same was not done for the secondary core. This ensured that potential river water contamination had the greatest chance of being detected.  Porewater was anaerobically extracted with the same method as the primary core. Major ion chemistry of these secondary core samples was compared to those of the primary core at the same depth (interval midpoints 17.5 cm, 22.5 cm, 27.5 cm below river bed). The resulting geochemical signatures were plotted in a Piper plot, shown in Figure 3.12.  Although the secondary core was taken from a relatively shallow depth, where groundwater had already experienced some dilution by river water, its porewater chemistry did not prominently coincide with the major ion signature of river water (Figure 3.12). Major ion chemistries of the radially-segmented samples had similar signatures as samples from the primary core at the same depth, which had its inner and outer surfaces removed. The innermost layers did, however, plot closer to the signature of river water, with successive outer layers plotting farther away. If contamination had occurred, it appears to have been more likely for the inner layers during insertion than for the outer layers during retrieval. These data suggest that even if river water contaminated the inner layers of the core during insertion, there was no major effect on porewater geochemistry. To eliminate this risk entirely, removal of the inner and outer frozen surfaces is almost certainly sufficient. 59    Figure 3.12 – A Piper plot showing major ion chemistry of extracted porewater samples is shown above. Blue data points represent the various layers of the half-size "secondary" core used for radial segmentation, as outlined by the legend in the upper left corner. This secondary core was 14 cm in length, representing depths from 16 to 30 cm below river bottom. The average depth of samples A-E is thus 23 cm. Black data points represent the major ion chemistry of porewater samples extracted from three intervals of the primary core (midpoint depths = 17.5, 22.5, and 27.5 cm below river bottom). The major ion chemistry of river water is shown by a red 'X'. The radially-segmented samples appear to have the same signatures as samples from the primary core, which had its inner and outer surfaces removed. The blue data points do not prominently coincide with the major ion signature of river water. This infers that contamination of the core by river water is not substantial. Inner layers, however, appear to have compositions closer to that of river water than the outer layers.     60  3.6 Challenges in the Field  A major challenge with working on the Fraser River is reduced underwater visibility as a result of high turbidity. Figure 1.1 in Chapter 1 shows that in April and May, turbidity spikes on the Fraser just prior to freshet, as sediments are released at the onset of snowmelt. At times outside this range, turbidity may peak suddenly as well. During these events, visibility with the underwater camera is zero. Although in some cases, sampling may be carried out blindly, this substantially increases the difficulty and chance of failure. As a result, it is important to proceed with sampling when turbidity on the river is adequately low. A suggested maximum turbidity value for ensuring marginal underwater visibility at a distance of two meters, as determined by camera tests in the field, is 40 NTU.  When visibility is sufficient, an underwater camera and spotlight vastly aid in site selection along the river bottom. The Fraser River sediments are littered with logs, organic matter, and metal debris from industrial operations. Being able to identify these obstacles and stay clear of them prior to sampler insertion is critical to actually obtaining a core, and ensuring that it remains intact during removal.  Perfect vertical insertion of the sampler is generally quite difficult. At slack tide, deployment conditions are ideal since there is no current to force the suspended sampler into an angled orientation. For the vast majority of situations, however, strong currents persist. This means that sampler insertion commonly takes place at a slight angle. Based on estimations of the winch line orientation at the water surface, this angle has never exceeded 20 degrees from the normal.  The "freeze anchoring" effect is a major challenge encountered during sample collection. Prior to removing the sampler, strong frictional forces between the frozen core and adjoining saturated sediments must be overcome. This presents large challenges, even with the use of a motorized winch. Future tests 61  may assess the effectiveness of a sonic vibration attachment to momentarily reduce this friction and aid in sampler retrieval.   62  Chapter 4 – Biogeochemical Characterization of Hyporheic Zone Sediments  4.1 Introduction  Abrupt physical, chemical, and biological gradients are found in the HZ as the loading of nutrients, organic matter, and other chemical species vary in space and time [5]. Versatile communities of microorganisms in the HZ both drive and take advantage of this variability by mediating numerous geochemical cycles [4]. These microbial metabolisms are responsible for a number of reactions, given appropriate conditions and a sufficient concentration of required chemical species [4]. Redox-sensitive elements such as O, N, Mn, Fe, S, and C are expected to be utilized most in these geochemical cycles. This chapter reviews microbial metabolisms studied in other environments and presents both geochemical and microbiological data from HZ sediment. Inferences on relevant metabolisms in the HZ are then proposed and discussed.  Aerobic respiration, which represents the oxidation of organic matter through O2 reduction, occurs in the shallowest of river sediments [4]. Oxygenated river water entering the HZ stimulates heterotrophic processes which rapidly modify the form of carbon. As a result, surface water mixing in the HZ is associated with a greater abundance of  microbial taxa that can degrade a wide range of organic compounds [82]. Aerobic respiration is thus often the most effective metabolism for the natural attenuation of organic contaminants in the HZ.   As the residence time of river water increases in the HZ, aerobic respiration progresses, and O2 becomes depleted. When this occurs, the denitrification of nitrate (NO3-) to molecular nitrogen (N2) becomes more prominent [35, 83, 84]. Denitrification is a known metabolism for many facultative anaerobic bacteria that degrade organic matter [85, 86]. Depending on the time of year and associated flow conditions, these bacteria may utilise either aerobic (i.e. O2 reduction) or anaerobic (i.e. NO3- reduction) pathways [87]. 63  Several researchers have also demonstrated the occurrence of nitrification in fluvial environments, which is exemplified by the oxidation of ammonia (NH3) or ammonium (NH4+) to nitrite (NO2-), and then nitrate via O2 reduction [88, 89].  Both Fe and Mn cycling are expected to be prominent in HZs with sharp redox gradients, but Fe is expected to play a greater biogeochemical role due to its higher environmental concentrations. Although Fe- and Mn-oxidizing bacteria may be populous enough in some environments to produce visible deposits of metal hydroxides, both processes yield relatively low amounts of energy compared to aerobic processes or denitrification [6]. In some sedimentary environments, however, Fe and Mn are cycled hundreds of times, and can be primary drivers of productivity [90, 91]. At sites similar to the River District, reductive dissolution of Fe and Mn were identified along the anaerobic flow path towards the river [60, 92]. In the HZ, oxidation of this dissolved Fe and Mn is expected.  Although Fe oxidation does occur abiotically when groundwater rich in ferrous iron encounters oxygen [6, 27], microbial oxidation of Fe is still expected. Microbially-mediated Fe oxidation also prevails  in anaerobic environments [93]. The biogeochemistry of iron and nitrogen, in some of these cases, are intimately linked. Coby et al. (2011) identified anaerobic laboratory cultures of β-proteobacteria (Dechloromonas) and δ-proteobacteria (Geobacter) which oxidized Fe2+ via NO3-reduction [94]. Straub et al. (2004) also identified three taxa of Fe2+-oxidizing NO3-reducers from freshwater sediments – these were in the genera of Acidovorax, Aquabacterium, and Thermomonas [95]. In both studies, these taxa and ferric iron-reducing bacteria were found to be interdependent, indicating that Fe and NO3 cycles are strongly related in many environments.  Gault et al. (2011) identified bacteriogenic iron oxides, composed mainly of ferrihydrite, which formed at a circumneutral groundwater seep. Scanning electron microscopy revealed iron sheath structures characteristic of a known Fe2+-oxidizing genus, Leptothrix. Other Fe2+-oxidizing taxa identified included 64  Gallionella spp. and Sideroxydans spp. At greater depths, higher concentrations of Fe2+ coincided with indicators of biological iron reduction. Here, Fe-reducing bacteria such as Rhodoferax ferrireducens and Geothrix fermentans were identified, along with many other δ-Proteobacteria [96]. In environments where Fe concentrations are high and steep redox gradients exist, it is very likely that microbially-mediated Fe cycling persists.  Manganese reduction has previously been identified during surface water infiltration into an alluvial aquifer in France. Along the flow path, Mn oxides were reduced after O2 and NO3 became depleted. Then, the flow path encountered a new aerobic zone, and Mn was precipitated out of solution [97]. Bacterially-mediated manganese oxidation has been identified in mining-affected streams, and is well-replicated in bacterial cultures [98]. At the pH and redox conditions typically encountered in freshwater HZs, Mn oxidation is favoured over reductive dissolution [10, 26]. One study found that approximately 20% of dissolved Mn discharging from groundwater into a stream was removed via microbially-mediated Mn oxidation, but this figure was believed to vary based on factors such as local reaction rates and residence times [26]. Although pH and redox conditions generally favour Mn oxidation in the HZ, sluggish reaction kinetics may keep Mn oxides from forming [26].  Sulfur is expected to play an important geochemical role in marine and estuarine environments, due to the large concentration of sulfate in seawater. When seawater intrudes an anoxic coastal aquifer, sulfate reduction occurs via oxidation of organic matter, as shown in Reaction 4.1 [37]. Sulfur has lower concentrations in freshwater systems, but has been investigated for its connection to other biogeochemical cycles. Wielinga et al. (1999) examined the biogeochemistry of sulfur in sediments adjacent to a creek affected by mine tailings. Sulfate-reducing bacteria (SRB) were distributed both at and below the oxic-anoxic interface and overlapped with both iron and sulfur oxidizers [99]. Chapelle and Lovely (1992) examined a scenario, in contrast, where SRB were competitively excluded by Fe-reducing bacteria in part of an aquifer with high concentrations of dissolved iron [100]. 65   SO42- + 2CH2O  ⇌  H2S + 2HCO3-  In the HZ of a river with riffle and pool sequences, sulfate reduction was found to be occurring where anaerobic groundwater discharged up to the surface, while sulfide oxidation was identified where aerobic freshwater entered the HZ [101]. Sulfur-oxidizing bacteria have been previously associated with environments low in photosynthetic activity, and in situations where both H2S and O2 were accessible nearby [102].   Methanogenesis has been found to coexist with sulfate reduction in organic-rich sediments [103]. These processes may or may not be competitive, depending on the organic substrates being utilized [104]. In the HZ of a desert stream, methanogenesis was identified not only in anoxic sediments, but also in oxic sediments, where it accounted for less than 1% of total respiration [105, 106]. The observation of methanogenesis in the aerobic zone of the HZ was reproduced by other researchers in another first-order stream [107]. At a site upstream from the River District, methanogenesis was identified along the anaerobic flow path towards the river, likely a result of organic contaminant degradation [92]. At an another site which also had high background concentrations of CH4 in groundwater, methane oxidation was identified in the HZ [108]. Elsewhere, Kohzu et al. (2004) used isotope ratios to demonstrate that aquatic macroinvertebrates in a HZ ecosystem utilized carbon which was first respired by methane-oxidizing bacteria [109]. In a study examining the biogeochemistry of a river with riffle and pool sequences, methanogenesis was associated with upwelling anaerobic groundwater, whereas methane oxidation was identified at locations of oxygenated river water recharge [101].  Motivation and Research Objectives  Since the cycling of redox-sensitive elements is examined at such a large variety of sites, our HZ investigation has only a broad basis for comparison. The HZ studied at the River District has no (4.1) 66  documented analogues in previous research, and thus the results discussed here represent a valuable addition to the field of HZ biogeochemistry. Uniqueness of the HZ at our site is characterized by highly variable oxygen concentrations as a function of freshet and tidal pumping, as well as saline groundwater discharge into a freshwater system. These factors are expected to strongly impact the distribution of various taxa within the microbial community. These conditions may also present unique niches for microorganisms with a high tolerance for challenging geochemical environments.    By examining the geochemistry of porewater and sediments in the HZ, as well as the microbiology, we seek to gain a better understanding of how microorganisms are affected by, and how they may control, their geochemical environment. This, in turn, allows us to compare our findings to conditions considered ideal for the natural attenuation of groundwater contaminants. Ultimately, contaminant fate in the HZ can be predicted, and subsequently assessed in a controlled setting. This research strives for a more process-based understanding of contaminant breakdown in the HZ.  4.2 Methodology  Core Sectioning  The frozen sediment core was sectioned following the protocol of the USGS Reston, Virginia Environmental Organic Geochemistry Laboratory [110]. This protocol was modified to accommodate the core's hollow centre and a different set of intended analyses. The frozen core, still in plastic wrap, was placed on a Styrofoam platform in a fumehood and covered with dry ice pellets during sectioning.  The core was measured and cut lines were marked on the plastic wrap at 5 cm intervals, generating 16 individual samples. All tools used for sectioning were washed with detergent and rinsed in distilled de-ionized water (DDIW) prior to use. Sectioning tools included a stainless steel "keyhole" style saw (see 67  Appendix H), and stainless steel putty knives. Significant effort was required to segment the core using this method, and thus for future attempts, mechanical cutting tools with diamond-bit blades are advised.  Once divided into 5 cm intervals, the samples were further sectioned for the various planned analyses. Approximately 5 mm of both the inner and outer surfaces of the core, which were most likely to experience contamination from river water, were scraped off.  Surfaces which melted due to friction from the saw were also removed. Each sediment interval was then broken in half, and approximately 10 g of frozen sediment was subsampled into 50 mL HDPE centrifuge tubes and immediately transferred to a  -20°C freezer for future DNA extraction and sequential Fe/Mn extraction. During the subsampling process for DNA extraction, all tools, work surfaces, and gloves were sterilized with ethanol to minimize contamination. Sterilized HDPE tubes were used to store the samples which would undergo DNA extraction. The remainder of the frozen sediment intervals, to be used for porewater extraction, were then placed in Ziploc bags and stored in a -20°C freezer.  Porewater Extraction  All porewater extraction work was completed in a Vinyl Type A anaerobic chamber by Coy Laboratory Products Inc. (Grass Lake, MI) filled with an anaerobic H2/N2 gas mixture supplied by Praxair (Vancouver, BC). All solutions used in the chamber were de-aired prior to use by bubbling N2 gas through them. A palladium catalyst fan box, also supplied by Coy, was used to circulate air within the chamber and scrub out low levels of O2. Equipment and reagents were transferred into the chamber through the attached vacuum airlock. This process consisted of a 3-cycle vacuum-and-purge procedure. After all materials were loaded into the airlock, the first cycle began by applying a vacuum, followed by a purge of inert N2 gas. This was then repeated for the second cycle. On the third cycle, a 10% H2 in N2 gas mix was used for purging, and the process was completed by applying a slight vacuum to the airlock.  68  The chamber O2 concentration was checked prior to, during, and following porewater extractions to ensure anoxic conditions. This was done with a reservoir of de-aired DDIW and a small fan overhead, thus maintaining equilibrium conditions between the reservoir and chamber atmosphere. By measuring the reservoir's dissolved oxygen (DO) content with a HACH DR/2400 portable spectrophotometer (method #8316), Henry's Law could be used to convert the aqueous oxygen concentration (µg/L) to an approximate atmospheric concentration (% v/v) in the chamber. Throughout the duration of the porewater extractions, the chamber's atmospheric O2 concentration was kept below 0.2% v/v.   Once all materials and equipment were loaded into the chamber (see Appendix H), the frozen sediment samples were transferred into the work area through the airlock. After being broken up into smaller portions, the samples were placed inside Nalgene filter holders with receivers (Part # 300-4000) and allowed to melt. While melting, the samples were compacted in the filtering units to increase connectivity of saturated soil pores. A Nalgene hand-operated vacuum pump was used to commence extraction, and porewater was drawn through Whatman ashless grade 41 filter paper (pore diameters of 20-25 µm) into the receivers below. To promote further extraction of porewater, a feed line N2 gas at 20 psi was connected to the upper chamber via a cover port, thus forcing out the remainder of the porewater sample into the receiver (see Appendix H). On average, porewater volume equivalent to 25% of the bulk volume was extracted from each frozen sample.  After porewater was extracted, the filtrate was drawn into a syringe and filtered once again through cellulose acetate syringe filters with a pore diameter of 0.45µm. This secondary filtrate was then used for analysis. Aliquots were assigned for analysis of pH, conductivity (specific conductance), cations, anions, dissolved ferrous iron (Fe2+), alkalinity, and dissolved oxygen (DO). Cation samples were acidified to 2% HNO3 and preserved until analysis by ICP-OES. Anion samples were analyzed via ion chromatography (AGAT Labs, Burnaby BC). pH was measured with a Thermo Orion 250A+ meter and a Fisher Scientific accumet® probe calibrated with pH 4, 7, and 10 buffers. Fe2+ concentrations were measured using the 69  1,10 Phenanthroline method (#8146) on a HACH DR/2400 portable spectrophotometer and were evaluated as a proxy for redox potential. Alkalinity was immediately determined following porewater extraction using the Gran titration method. DO was measured using the Indigo Carmine method (#8316) on the HACH DR/2400 spectrophotometer. The remaining sediment samples, having been anaerobically dried, were then subsampled into 50 mL HDPE centrifuge tubes for future mineralogical analysis.  Sequential Extractions  Sequential extractions were completed on samples from the sediment core to investigate partitioning of Fe and Mn on grain surfaces. The extractions were designed to consecutively target the exchangeable phase, the amorphous phase, the crystalline phase, and residual phase of Fe and Mn. These extractions were strictly operationally defined, since target phases were not exclusive to each step and results were dependent on factors such as exposure time and grain size [111]. Existing protocols found to be representative of these target phases were selected for this work. Table 4.1 describes the extractants used for each target phase in these experiments, and the original authors of the methods. The table of methods is derived from Jia (2015), who used a similar set of methods on comparable sediments taken from a nearby Fraser River Sands aquifer [60].  Table 4.1 – Sequential extraction steps used to target various phases of Fe and Mn are shown below. Adapted from Jia (2015). Step Targeted Phase Reagents Solution-solid ratio (mL/g) Extraction Mechanism 1 Exchangeable and adsorbed ions [1] 1M CaCl2 80 Cation exchange 2 Amorphous oxides [2] 0.5M HCl 80 Proton dissolution, Fe-Cl complexation 3 Oxides with intermediate crystallinity [3] 1M NH2OH-HCl in 25% CH3COOH 80 Reductive dissolution of Fe(III) to Fe(II) 4 Residual extractable fraction, or "near-total" digestion [3] HNO3-HClO4-HF-HCl, full strength 50 Near-total dissolution [1] Heron et al. (1994), [2] Lovley and Phillips (1986), [3] Hall et al. (1996) [111] [112][113] 70  After being sectioned and subsampled, the sediments were weighed out into 60 mL HDPE centrifuge tubes. Approximately 0.5 g of sediment was transferred into the HDPE tube, and this was done in duplicate for each of the 16 core intervals. An additional subsample from each interval was also taken to measure water content and porosity by heating the sediment to 100°C for 24 hours. Steps 1 and 2 were completed in the anaerobic chamber to avoid oxygen contamination of the sediment samples. Step 1 was initiated by adding 40 mL of 1M CaCl2 to the samples. The samples were vortexed and then shaken for 24 hours in a Julabo SW22 shaking water bath. Next, the tubes were removed from the shaker, placed in a centrifuge, and centrifuged at 2500 rpm for 30 minutes. The supernatant was decanted into a 50 mL HDPE syringe and filtered through 25mm-diameter 0.45µm pore size cellulose acetate lockable filters. The filtrate was then spiked with HNO3 to an approximate pH of 2 for preservation and stored at 4°C until analysis. Prior to beginning Step 2, the sediment remaining in the centrifuge tube was rinsed. This was done by adding 10 mL of DDIW, vortexing the sample, and centrifuging the sample at 2500 rpm for 10 minutes. The rinsing process was always done twice.  For Step 2, 40 mL of 0.5M HCl was added to each rinsed sample. Vortexing, shaking, centrifugation, and filtration were performed identically to Step 1. Since the 0.5M HCl step was expected to maintain the redox state of extracted Fe [112], a speciation experiment was completed on each aliquot. This was done with the 1,10 Phenanthroline method (#8146) on a HACH DR/2400 portable spectrophotometer. The remaining filtrate was then spiked with HNO3 to pH < 2 and stored at 4°C until analysis.  The samples were rinsed once again, and Step 3 was carried out. This involved first adding 25 mL of 1M NH2OH-HCl in 25% CH3COOH to each sample. The samples were then placed in a 90°C water bath for 3 hours, with samples being removed once every 20 minutes to be vortexed. Next, the samples were centrifuged at 2500 rpm for 10 minutes, and the supernatant was decanted into a syringe. Subsequently, 7.5 mL of 25% CH3COOH was added to the sample tube, which was vortexed and centrifuged again at 2500 rpm for 10 minutes. The supernatant was again decanted, another 7.5 mL of 25% CH3COOH was 71  added, and centrifugation was repeated. In total 40 mL of extractant was collected in the syringe, filtered, preserved with HNO3 to pH < 2, and refrigerated. Prior to Step 4, the remaining sediment was rinsed with DDIW and dried.  All extracts from Steps 1 through 3 were analyzed simultaneously for Fe and Mn with ICP-OES. Reagent blanks (DDIW) were also performed for each step, and produced concentrations below the detection limit. Reported concentrations were converted to mg/g of dry sediment weight, or ‰ (m/m). Step 4 was completed by the ALS Laboratory Group in North Vancouver, BC. Dried samples were submitted for manual pulverization and a four-acid "near-total" digestion, which used a mixture of HNO3, HClO4, HF, and HCl. The digestion residue was then topped up with dilute HCl and the resulting solution was analyzed by ICP-AES (ALS method ME-ICP61).  Four of the 16 sediment intervals were subsampled in triplicate as opposed to duplicate to assess mass loss during Steps 1-3. Triplicate samples were submitted to ALS for a four-acid digestion, without having been exposed to Steps 1 through 3. Differences between these triplicates (Step 4 only) and singles/duplicates (Steps 1-4) were calculated to compare the net masses of extracted Fe and Mn – a low value obtained here implied minimal unaccounted mass. For Fe, the average difference was 7.5%, while for Mn, the average difference was only 4.7% (see Appendix E). Mass conservation through each extraction step was thus considered acceptable.  DNA Extraction and Sequencing  For living organisms, deoxyribonucleic acid (DNA) serves as a stable, life-long medium for storing and transmitting genetic information. Ribonucleic acid (RNA), however, allows for this genetic information to be accessed by the organism on a regular basis. RNA transfers genetic instructions from an organism's nucleus to the ribosome, where proteins can then be generated for routine functions. While DNA is 72  relatively more stable and holds information for longer periods of time, RNA is more versatile, since it allows for the utilization of important genetic information.  Although DNA sampled from the environment serves as an indication of which organisms are present, it cannot be used to determine which organisms are active. Organisms identified in a community based on DNA alone may thus be dormant. RNA, conversely, can be used to determine which organisms are actively manufacturing proteins in the environment, thus giving an indication of which ones are the most metabolically active.  To investigate the composition of the microbial community in the HZ, ribosomal DNA (rDNA) was studied. The community was examined through a 16S rRNA gene tag-sequencing survey. Since DNA yields no information on the relative activity of the identified taxa, results only indicate which organisms are present.   DNA was extracted from subsamples of frozen sediment core using the PowerSoil® DNA Isolation Kit from Mo Bio Laboratories (Carlsbad, CA) according to the manufacturer's protocol. Purity and quantity of DNA were assessed using Picogreen® and gel electrophoresis on a 0.8% agarose gel containing 0.5 µg/mL ethidium bromide.   Purified genomic DNA was submitted to Microbiome Insights Inc. at the University of British Columbia in Vancouver, BC. Samples were amplified by PCR, in triplicate, using barcoded primer pairs adjoining the V3 region of the 16S rRNA gene [114]. The PCR program consisted of an initial DNA denaturation step at 95°C (5 min.), 25 cycles of DNA denaturation at 95°C (1 min.), an annealing step at 50°C (1 min.), an elongation step at 72°C (1 min.), and a final elongation step at 72°C (7 min.). To ensure that contamination did not occur, controls devoid of template DNA were also included.   73  Sequence amplicons were run on a 2% agarose gel to ensure amplification was adequate. Those that displayed bands at a length of approximately 160 base pairs were purified using the illustra GFX PCR DNA purification kit. Purified samples were diluted 50 times and quantified using PicoGreen® in a TECAN M200. Pooled PCR amplicons were diluted down to 20 ng/µL and sequenced at the V3 hypervariable region using Hi-Seq 2000 bidirectional Illumina sequencing and a Macrogen Cluster Kit v4. Library preparation was completed using a TruSeq DNA Sample Prep v2 Kit (Illumina) with 100 ng of DNA and a QC library provided by an Agilent DNA 1000 Bioanalyzer.  Sequences were processed using Mothur [115] and custom PERL scripts. When sequences contained ambiguous characters, had homopolymers longer than 8 bp, or did not match a reference alignment in the correct sequencing region, they were removed from the analysis. Sequences were checked for chimeras using UCHIME [116]. Reads were subsequently clustered into operational taxonomic units (OTUs) at a similarity threshold of 97% based on uncorrected pairwise distance matrices. OTUs were classified using the SILVA reference taxonomy database (release 123).  Sediment Mineralogy  Rietveld X-ray diffraction (XRD) was completed for 4 separate intervals of the sediment core. These intervals had midpoint depths of 22.5, 37.5, 47.5, and 67.5 cm below the river bottom. The samples were first manually ground with a mortar and pestle, and then reduced to the optimum grain-size range for quantitative X-ray analysis (<10 µm). Grinding of each sample completed under ethanol in a vibratory McCrone Micronizing Mill for 10 minutes. The samples were set to dry in a fumehood for 24 hours and then mounted in a back-loading cavity using a sheet of sandpaper to randomize grain orientation. X-ray powder diffraction data were collected continuously over a range 3-80°2θ with CoKα radiation on a Bruker D8 Advance Bragg-Brentano diffractometer equipped with a Fe monochromator foil, 0.6 mm (0.3°) divergence slit, incident- and diffracted-beam Soller slits and a LynxEye-XE detector. The long 74  fine-focus Co X-ray tube was operated at 35 kV and 40 mA, using a take-off angle of 6°. X-ray diffractograms were analyzed using the International Centre for Diffraction Database PDF-4 and Search-Match software by Bruker.  X-ray powder-diffraction data were refined with the Rietveld program Topas 4.2 (Bruker AXS).  Samples were examined by scanning electron microscopy (SEM) to qualitatively inspect for iron and manganese precipitation on sediment grains. The higher mass number for Fe and Mn make these elements easy to identify amongst the lower mass number of other elements which make up the majority of the mineral's structure (Al, Si, O for samples analyzed here). Sediments were dried in the anaerobic chamber, mounted onto sample stubs, and coated with graphite in an Edwards Auto306 carbon evaporation system. The samples were then transferred to the vacuum chamber and examined on a Philips XL-30 SEM, equipped with a Gamma-Tech energy dispersive x-ray spectrometer and Bruker Quantax EDX system.  4.3 Results and Discussion  Core Description  After removing small portions of the base of the core which was slightly deformed during storage, the length of the core was measured to be 85 cm. In the upper half, the core was composed mainly of uniformly-graded coarse sands, with a few thin lenses of finer sands (see Figure 3.8 in Chapter 3). These fine-grained lenses appeared to have the same orientation, likely representative of sediment deposition processes. Approximately four silt lenses with a thickness of less than 1 cm were noted between the depths of 55 and 65 cm, all of which had different orientations. From a depth of 65 to 70 cm, inclusions of woody debris were observed. Below the depth of 70 cm, the sediment composition shifted to coarser sand and included some pebbles with diameters up to 2 cm. 75  Porewater Geochemistry  All geochemical parameters measured from porewater were plotted with depth. Figures 4.1, 4.2, 4.3, and 4.6 show results in mg/L – identical graphs showing molar concentrations are available in Appendix C. Ca, Mg, Na and K concentrations all increase with depth (Figure 4.1). Both Mg and K concentrations increase by about an order of magnitude with depth, whereas Ca concentrations only increase by a factor of 5. The concentration of Na increases at a much greater rate than the other solutes, especially below the depth of 32.5 cm. Na concentrations increase from 2.1 mg/L at the top to 354 mg/L bottom – a difference of more than two orders of magnitude. In the upper 45 cm, Ca concentrations are higher, while in the lower 40 cm, Na concentrations are higher.      01020304050607080900.1 1 10 100Depth (cm) Concentration (mg/L) NaCaMgKFigure 4.1 – Dissolved concentrations of Na, Ca, Mg, and K are shown. The horizontal axis is logarithmic. Mg and K concentrations increase by about an order of magnitude with depth, whereas Ca concentrations only increase by a factor of 5. The concentration of Na increases much faster than other solutes, especially below the depth of 32.5 cm. Na concentrations increase from 2.1 mg/L at the top to 354 mg/L bottom – an increase factor of 170. Although Ca concentrations increase with depth, they do so in a sinuous manner, inferring tidally-induced ion exchange. 76  Porewater concentrations of Fe, Fe(II), and Mn are plotted in Figure 4.2. In the top-most sediments, Fe concentrations increase slowly with depth from 0.08 mg/L at 12.5 cm to 0.33 mg/L at 37.5 cm. Below this, Fe levels increase rapidly with depth, reaching 22 mg/L at 82.5 cm. Field work completed for Chapter 2 indicated that Fe concentrations in saline upwelling groundwater ranged from 3.2 to 138 mg/L. It is thus quite possible that Fe concentrations continue to increase with depth below 85 cm.  Fe(II) is only first detectable at a depth of 32.5 cm. Beginning at a depth of 42.5 cm, Fe(II) concentrations are nearly identical to total Fe concentrations, and this trend continues with increasing depth. This infers that redox conditions at 42.5 cm and below are sufficiently reducing that essentially all dissolved Fe exists as ferrous iron (Fe2+).      01020304050607080900.01 0.1 1 10Depth (cm) Concentration (mg/L) FeFe(II)MnFigure 4.2 – Total Fe, Fe(II), and Mn concentrations are shown. The horizontal axis is logarithmic. Fe concentrations increase slowly with depth from 0.08 mg/L at 12.5 cm to 0.33 mg/L at 37.5 cm. Below this, Fe levels increase rapidly with depth, reaching 22 mg/L at 82.5 cm. Fe(II) is only first detectable at a depth of 32.5 cm . Beginning at a depth of 42.5 cm, Fe(II) concentrations are nearly identical to total Fe concentrations, and this trend continues with depth. Since Fe and Fe(II) do not appear to reach a maximum, it is possible that their concentrations continue to increase with depth. 77  In all but the shallowest portions of the HZ investigated here, measureable Mn is expected to represent Mn(II), a reduced species. This is because the redox potential at which Mn(II) becomes the dominant Mn species is quite high (see Figure 1.3, Chapter 1). In the top-most sediments, the porewater concentration of Mn is only 0.1 mg/L, but it increases with depth to 2.5 mg/L at 22.5 cm. At depths below 25 cm, Mn concentrations remain stable, ranging only between 2.4 and 3.7 mg/L down to a depth of 85 cm. The fact that Mn concentrations remain constant in the 25-40 cm depth range despite dilution of conservative species (e.g. Cl, discussed below) suggests that reduced Mn is accumulating at this location.      Porewater concentrations of SO4, NO3, and Cl are plotted in Figure 4.3. NO3 concentrations decrease with depth, from 3.5 mg/L in the top-most sediments, to 0.7 mg/L at a depth of 32.5 cm. This suggests that NO3 is supplied mainly from the river, although these results could also be explained by oxidation of 01020304050607080900.1 1 10 100 1000Depth (cm) Concentration (mg/L) ClNO₃ SO₄ Figure 4.3 – Cl, NO3, and SO4 concentrations are shown. The horizontal axis is logarithmic. NO3 concentrations initially decrease with depth. From 50 cm downward, NO3 concentrations increase, and then remain stable. This may be a result of sample oxidation. SO4 concentrations are highest in the shallowest interval. With depth, SO4 concentrations decrease. A local peak occurs at 52.5 cm, and this may be associated with sulfide oxidation. From the top to the bottom, Cl increases by a factor of 170. Dilution in the HZ appears to persist down to 85 cm, and possibly further. 78  ammonia and nitrite to nitrate. Between the depths of 32.5 and 52.5 cm, NO3 concentrations are stable, not exceeding 1.5 mg/L. Between 52.5 cm and 62.5 cm in depth, NO3 concentrations actually increase to 2.5 mg/L. From here to the bottom of the core, concentrations remain in the 2.0 mg/L range.  Sulfate concentrations are highest in the shallowest sediment interval, at 8.0 mg/L. This is comparable to SO4 levels in the Fraser, measured to be 6.8 mg/L. With depth, SO4 concentrations decrease, with an especially rapid decline between the depths of 20 and 45 cm. The lowest SO4 concentration of 0.6 mg/L is found at the bottom of the core. At a depth of 52.5 cm, there is an unexpected large peak in SO4 at 6.6 mg/L. This peak may be associated with sulfide oxidation, or it may be a sampling artifact. Although H2S concentrations were not measured, discharging groundwater was noted to smell strongly of rotten eggs, indicative of sulfide.   At a depth of 12.5 cm, the Cl concentration is 3.9 mg/L, whereas at 82.5 cm, it is 661 mg/L. From the top to the bottom of the core, the porewater concentration of Cl increases by a factor of about 170 – the same as for Na. Cl is generally unreactive and can be treated as a conservative tracer. Since the core was sampled in an area of saline groundwater discharge, the extent of groundwater dilution can be characterized via Cl concentrations and conductivity. This is shown by the specific conductance of porewater, which increases with depth, in comparable manner to Cl concentrations (Figure 4.4). In the HZ characterized here, dilution appears to persist down to 85 cm (the bottom of the core), and likely even further. The inferred dilution rate for HZ porewater migrating from this depth to the surface, based on Cl and Na concentration gradients, is apparently more than two orders of magnitude.  With depth, porewater pH increases (see Figure 4.5). In the top-most sediments, pH is measured to be 7.07, whereas at 52.5 cm, pH is 7.76. Superimposed on this downward increasing trend, a local peak occurs at a depth of 32.5 cm (pH = 7.76) and a local depression occurs at a depth of 42.5 cm (pH = 7.36). Below the depth of 55 cm, pH remains stable, varying only between 7.80 and 7.87. During groundwater 79  discharge characterization (see Chapter 2), the highest measured pH of any HZ sample from a depth of 1 m was 7.74. The slightly higher pH of core porewater samples may indicate that some degassing of CO2 occurred during analysis in the anaerobic chamber. This is because degassing of CO2 following exposure to an open atmosphere causes a shift in the equilibrium of carbonic acid, resulting in a reduction of its aqueous concentration, thus lowering acidity.      Dissolved O2 (DO) concentrations, as expected, decrease with depth (see Figure 4.6). The highest DO measurement of 0.15 mg/L is observed in the top-most sediments (4-10 cm interval). This value is more than an order of magnitude lower than the anticipated concentration of DO in water which is in equilibrium with Earth's atmosphere (8-9 mg/L). A DO test blank yielded a value of 0.012 mg/L, and the error for each measurement was given as +/- 0.003 mg/L. DO measurements which fall within the shaded box of Figure 4.6 thus effectively represent zero DO values (52.5 cm and 82.5 cm). Within the overall 01020304050607080900 500 1000 1500 2000Depth (cm) µS/cm Specific Conductance 01020304050607080906.5 7 7.5 8 8.5Depth (cm) pH Figure 4.4 – Specific conductance is shown with depth. The downward increase is similar to that of Cl.   Figure 4.5 – pH is shown with depth. pH increases with depth initially, but then remains stable. A local peak (7.76) occurs at 32.5 cm while a local depression (7.36) occurs at 42.5 cm.  80  decreasing trend of DO with depth, two significant depressions in DO occur – one between the depths of 15 and 30 cm, and another between the depths of 45 and 55 cm.  A surprising observation from these measurements is that DO is still detectable in the deepest porewater intervals, which are expected to be anaerobic. Work done for this analysis was conducted in an anaerobic chamber with gaseous O2 concentrations kept below 0.2% v/v. This corresponds to a solubility of 0.08 mg/L (Henry's Law), which is higher than most measurements made here. Although porewater was analyzed immediately following extraction, it is possible that samples were contaminated by rapid dissolution of O2. As a result of this possibility, DO measurements likely include some degree of contamination or sampling artifacts.  Porewater alkalinity was found to increase with depth (see Figure 4.7). The minimum alkalinity value, observed in the top-most sediment interval, was 0.58 mEq/L. Within the overall trend of increasing alkalinity with depth, three depressions were noted at the depths of 32.5 cm, 47.5 cm, and 72.5 cm. Moving downward from these depressions, alkalinity was found to resume its steady increase with depth once again. Peak alkalinity, measured to be 2.96 mEq/L, was observed in the deepest interval.  Major ion chemistries of each porewater interval were graphed on a Piper plot (see Figure 4.8) to examine vertical trends in water type. Sample depth (in cm) is indicated next to each point plotted in the central field.  Water types for the 5 upper-most intervals (4 to 30 cm), in addition to river water, all plotted in the Ca-HCO3 region, on the left side of the diamond plot. The bottom six intervals (55 to 85 cm) conversely, all plotted in the Na-Cl region, on the far right side of the diamond plot. Water types for the middle 5 intervals showed a smooth transition between these extremes, plotting as follows; 32.5 cm in the Ca-HCO3-Cl region, 37.5 and 42.5 cm in the Ca-Na-HCO3-Cl region, 47.5 and 52.5 cm in the Ca-Na-Cl region. The depths at which the transition in water type occurs (30 to 55 cm) coincides with the sharpest 81  change in concentrations of both Na and Cl, as well as Fe(II)/Fe. The greatest variability in both pH and alkalinity is also noted to occur at these depths.          01020304050607080900 0.05 0.1 0.15Depth (cm) mg/L Dissolved O₂ 01020304050607080900 1 2 3 4Depth (cm) meq/L Alkalinity Figure 4.6 – Dissolved O2 (DO) decreases with depth. Peak DO occurs at the top, as expected. A DO test blank yielded a value of 0.012 mg/L, and the error for each measurement was given as +/- 0.003 mg/L. DO measurements which fall within the shaded box thus effectively represent zero DO values (52.5 cm and 82.5 cm). Within the overall decreasing trend of DO with depth, two significant depressions in DO occur – one between the depths of 15 and 30 cm, and another between the depths of 45 and 55 cm.  Figure 4.7 – Porewater alkalinity increases with depth. Minimum alkalinity is observed at the top. Within the overall trend of increasing alkalinity with depth, three depressions are noted at the depths of 32.5 cm, 47.5 cm, and 72.5 cm. Alkalinity was determined using the Gran titration method.  82          Figure 4.8 – A Piper plot demonstrating major ion chemistry of porewater extracted from all sediment intervals is shown. Numbered labels represent the depth interval midpoint for each sample in centimetres. Water types for the 5 upper-most intervals (7.0 to 27.5 cm), all plot in the Ca-HCO3 region, on the left side of the diamond plot. The bottom six intervals (57.5 to 82.5 cm) all plot in the Na-Cl region, on the far right side. Water types for the middle 5 intervals show a smooth transition between these end-members.   62.5 67.5 77.5 River 7.0 12.5 17.5 27.5 32.5 37.5 42.5 47.5 52.5 57.5 72.5 82.5 22.5 83  Sequential Extractions  Results from sequential extractions of Fe and Mn are displayed in Figures 4.9 and 4.10, respectively. Fractions of Fe and Mn extracted during the experiments are operationally divided into exchangeable, amorphous, crystalline, and residual portions. Values for all fractions are shown in mg of Fe or Mn per gram of dry sediment.  Figure 4.9 shows that the exchangeable Fe fraction increases from 0.0026 mg/g (2.6 ppm) in the shallowest interval to 0.16 mg/g (160 ppm) in the deepest interval. The sharpest increase occurs between the depths of 40 and 60 cm. This increase in exchangeable Fe mirrors the rapid increase in porewater concentrations of Fe at the same depth. The amorphous Fe fraction increases modestly with depth, from 3.2 mg/g (0.32 %) in the top-most sediments to 4.1 mg/g (0.41%) in the deepest sediments. At the depths of 42.5 and 72.5 cm, minor drops in amorphous Fe content are noted, followed by steady resumed increases with depth. The crystalline fraction of Fe has no significant variability with depth. Similarly, although the residual Fe fraction demonstrates some variability with depth, no significant trend is noticeable, and the average value of residual Fe fraction is about 17 mg/g (1.7%). Assuming minimal participation in biogeochemical reactions, the lack of increase or decrease in the crystalline and residual fractions is expected, as the nature of the sand was observed to be consistent with depth in the core sample.  84      0204060800.001 0.01 0.1 1 10 100Depth (cm) mg Fe / g sediment 1M CaCl20.5M HCl1M NH2OH-HCl in 25% AcOHHNO3-HClO4-HF-HCl digestionSequential Extractions Step 1: 1M CaCl2 - targeting adsorbed/exchangeable ions Step 2: 0.5M HCl - targeting reactive amorphous oxides Step 3: 1M NH2OH-HCl in 25% AcOH - targeting crystalline oxides Step 4: Total digestion in HNO3-HClO4-HF-HCl - targetting total residual portion exchangeable Fe 0204060800 1 2 3 4 5Depth (cm) mg Fe / g sediment amorphous Fe crystalline Fe 0204060800 5 10 15 20 25Depth (cm) mg Fe / g sediment residual Fe 0% 10% 20% 30% 40% 50% 60% 70% 80% 90% 100%712.517.522.527.532.537.542.547.552.557.562.567.572.577.582.5mg Fe / g sediment Depth interval midpoint (cm) amorphouscrystallineresidualFigure 4.9 – Results from sequential Fe extractions are shown above. Data are given in mg of Fe per gram of sediment. Step 1 targeted the exchangeable fraction, Step 2 targeted the amorphous fraction, Step 3 targeted the crystalline fraction, and Step 4 targeted the residual fraction. 85      0204060800.01 0.1 1Depth (cm) mg Mn / g sediment 1M CaCl20.5M HCl1M NH2OH-HCl in 25% AcOHHNO3-HClO4-HF-HCl digestionSequential Extractions Step 1: 1M CaCl2 - targeting adsorbed/exchangeable ions Step 2: 0.5M HCl - targeting reactive amorphous oxides Step 3: 1M NH2OH-HCl in 25% AcOH - targeting crystalline oxides Step 4: Total digestion in HNO3-HClO4-HF-HCl - targetting total residual portion exchangeable Mn 0204060800 0.05 0.1 0.15 0.2 0.25 0.3 0.35 0.4 0.45Depth (cm) mg Mn / g sediment amorphous Mn residual Mn crystalline Mn 0% 10% 20% 30% 40% 50% 60% 70% 80% 90% 100%712.517.522.527.532.537.542.547.552.557.562.567.572.577.582.5mg Mn / g sediment Depth interval midpoint (cm) exchageableamorphouscrystallineresidualFigure 4.10 – Results from sequential Mn extractions are shown above. Data are given in mg of Mn per gram of sediment. Step 1 targeted the exchangeable fraction, Step 2 targeted the amorphous fraction, Step 3 targeted the crystalline fraction, and Step 4 targeted the residual fraction. 86  In Figure 4.10 the exchangeable fraction of Mn is found to increase from 0.02 mg/g (20 ppm) in the shallowest interval to a peak of 0.05 mg/g (50 ppm) at a depth of 32.5 cm. Exchangeable Mn drops back down to below 0.02 mg/g (20 ppm) at the base of the core. Compared to Fe, the exchangeable content of Mn contributes a noticeably higher proportion of total Mn, peaking at 10%. In contrast to Fe, the content of amorphous Mn actually decreases with depth. Peak amorphous Mn (0.13 mg/g, 130 ppm) occurs at a depth of 17.5 cm, which coincides with the depth where porewater Mn concentrations decrease rapidly in the upward direction. This suggests that Mn oxidation is occurring between the depths of 10 and 25 cm, immediately above the region of suspected Mn reduction. As observed with Fe, the crystalline fraction of Mn displays little variation with depth.  There are strong correlations between exchangeable sediment fractions of Fe and Mn and porewater concentrations of Fe and Mn, (Figure 4.11). The top-most graph in Figure 4.11 displays the porewater charge-equivalent concentrations of Fe and Mn with depth – that is, the proportion of total cationic charge provided by both elements. When plotted against the exchangeable sediment fractions of Fe and Mn at the same depth, R2 values of 0.897 and 0.723 are obtained, respectively. This strong relationship may be a function of coulombic attraction of cations to negatively-charged sites on sediment surfaces. For Fe and Mn in porewater, specific adsorption to sediment surfaces would mostly likely occur via inner-sphere complexation.   Amorphous iron extracted during Step 2 was speciated.  Figure 4.12 (bottom) displays the variation in Fe speciation for amorphous minerals formed on sediment surfaces. At the shallowest depths, Fe(II) makes up approximately 1% of all Fe bound in the amorphous fraction. At the bottom of the core, this number increases to approximately 2%. This is anticipated, as anaerobic metabolisms are expected to dominate at greater depths, where higher sulfide concentrations would likely scavenge out ferrous iron to form iron sulfides. In some ways, it is surprising that this percentage is not higher. If only 2% of all iron in the amorphous fraction is indeed ferrous, anaerobic iron oxidation must be occurring at greater depths.  87      01020304050607080900 0.005 0.01 0.015 0.02 0.025 0.03 0.035Depth (cm) Proportion of  total cationic charge FeMnporewater concentration 01020304050607080900 0.05 0.1 0.15 0.2Depth (cm) mg Fe / g sediment exchangeable Fe on sediment 01020304050607080900 0.02 0.04 0.06Depth (cm) mg Mn / g sediment exchangeable Mn on sediment 00.040.080.120.160.20 0.01 0.02 0.03 0.04Exchangeable Fe (mg/g) Fe proportion of total cationic charge 00.010.020.030.040.050 0.01 0.02Exchangeable Mn (mg/g) Mn proportion of total cationic charge Figure 4.11 – Comparisons between porewater concentrations and exchangeable fractions of Fe and Mn are shown. The top figure shows porewater charge-equivalent concentrations, the middle figures show the exchangeable fractions extracted from sediment, and the bottom figures show correlations for these data sets (0.897 for Fe and 0.723 for Mn). These strong relationships may be a function of coulombic attraction of cations to negatively-charged sites on sediment surfaces. 88          01020304050607080900 2 4 6Depth (cm) [Fe] mg/g 01020304050607080900 0.05 0.1Depth (cm) [Fe2+] mg/g 01020304050607080900 0.005 0.01 0.015 0.02Depth (cm) [Fe2+]/[Fe(tot)] Figure 4.12 – Results from Fe speciation during extraction of amorphous sediment fractions are shown above. Total amorphous Fe is shown at top left, amorphous Fe(II) at top right, and Fe(II) as a fraction of total Fe at the bottom. At the shallowest depths, Fe(II) makes up approximately 1% of all Fe bound in the amorphous fraction. At the bottom of the core, this number increases to approximately 2%. amorphous Fe(tot) on sediment amorphous Fe2+ on sediment 89  DNA Sequencing  From all depth intervals combined, the two dominant bacterial phyla, making up more than half of all identified sequences, were the well-documented Proteobacteria (44.1% of all sequences) and the less understood Acidobacteria (13.8%). Amongst the Proteobacteria, the Deltaproteobacteria had the greatest abundance (11.6%), followed by the Betaproteobacteria (10.1%), the Gammaproteobacteria (8.3%), the Alphaproteobacteria (6.5%), and the Epsilonproteobacteria (0.2%). Unclassified Proteobacteria made up 7.4% of all sequences. The remaining bacterial phyla in decreasing order were as follows; Chloroflexi (6.8%), Bacteriodetes (6.3%), Gemmatimonadetes (4.8%), Chlorobi (4.6%), Actinobacteria (4.6%), Nitrospirae (4.3%), Planctomycetes (3.9%), Verrucomicrobia (2.3%), and Spirochaetae (0.6%). The domain of Archaea represented 3.9% of all sequences, with the main phylum being Euryarchaea (3.0%). Figure 4.13 shows the distribution of these phyla with depth, indicating that most families and genera within these phyla were specialized towards a shallow aerobic niche, or conversely, a deeper anaerobic niche. Figure 4.14, additionally, shows the distribution of major classes within the above phyla.  The total number of sequences obtained per depth interval varied significantly, as shown in Figure 4.15. Maxima were noted at depths of 32.5 cm (24 974 sequences) and 72.5 cm (23 541), while minima were identified at 12.5 cm (10 324) and 52.5 cm (11 695). This significant variation was likely a result of bias during sampling or processing. To account for this, taxa were plotted in Figures 4.13 and 4.14 with depth by percentage of total DNA sequences, and not their absolute number.   Seven classes were found to demonstrate higher proportional abundances in the upper section of the core, suggesting they were composed predominantly of species with aerobic metabolisms (see Figure 4.14). These were the Acidobacteria (peak of 23.8% at 27.5 cm), the Sphingobacteriia (11.5% at 7 cm), the Betaproteobacteria (11% through top 50 cm), Planctomycetacia (5.2% at 27.5 cm), the Spartobacteria (4.6% at 7 cm), Verrucomicrobiae (1.2% at 22.5 cm), and the Flavobacteriia (0.7% at 7 cm). 90      0% 20% 40% 60% 80% 100%712.517.522.227.532.537.542.547.552.557.562.567.572.577.582.5Depth interval midpoint (cm) Percent of Total Reads AcidobacteriaActinobacteriaBacteroidetesChlorobiChloroflexiEuryarchaeotaGemmatimonadetesNitrospiraePlanctomycetesProteobacteriaSpirochaetaeVerrucomicrobia0% 10% 20% 30% 40% 50% 60% 70% 80% 90% 100%7.012.517.522.227.532.537.542.547.552.557.562.567.572.577.582.5Percent of Total Reads Depth interval midpoint (cm) AcidimicrobiiaAcidobacteriaAlphaproteobacteriaAnaerolineaeBetaproteobacteriaDeltaproteobacteriaGammaproteobacteriaGemmatimonadetesHolophagaeIgnavibacteriaMethanomicrobiaNitrospiraPhycisphaeraePlanctomycetaciaSpartobacteriaSphingobacteriiaSpirochaetesThermoleophiliaThermoplasmataotherunclassifiedFigure 4.13 – The distribution of major phyla with depth is shown above. Results are shown in percent of total sequences. For many phyla, distribution is weighted towards either the upper aerobic zone, or the lower anaerobic zone. All phyla listed are in the domain of bacteria, except for Euryarchaeota, which are part of the domain of archaea. Figure 4.14 – The distribution of major classes with depth is shown above. Results are shown in percent of total sequences. A large portion of taxa remain unidentified at the class rank. Classes which made up a minor proportion of total reads were grouped into the "other" category. Dominant Phyla 91      Another seven classes were found to demonstrate higher proportional abundances in the lower section of the core, suggesting a predominant composition of anaerobic species (see Figure 4.14). These were the Deltaproteobacteria (19.9% at 67.5 cm), Anaerolineae (16.0% at 67.5 cm), Gammaproteobacteria (10.8% at 72.5 cm), Nitrospira (5.0% at 57.5 cm), Thermoplasmata (3.7% at 67.5 cm), Spirochaetes (1.8% at 67.5 cm), and the Epsilonproteobacteria (1.3% at 72.5 cm).  Three classes also demonstrated their highest proportional abundances in the zone between the aerobic and anaerobic zones. These were the Ignavibacteria (8.4% at 52.5 cm), Thermoleophilia (4.1% at 52.5 cm), and Acidimicrobiia (2.1% at 47.5 cm). Although few taxa reached their peak abundance in this depth range (30-55 cm), this transition zone could theoretically offer a competitive advantage. This portion of the core was the location of the strongest gradients in porewater geochemistry and thus may offer a wider 01020304050607080900 5000 10000 15000 20000 25000 30000Depth (cm) Total Count Figure 4.15 – The total number of sequences obtained per depth interval is shown above. Maxima are noted at depths of 32.5 cm (24 974 sequences) and 72.5 cm (23 541), while minima are identified at 12.5 cm (10 324) and 52.5 cm (11 695). Since the number of sequences varies significantly, distribution of all taxa is plotted with depth by percentage of total sequence reads, and not their absolute number.  92  variety of redox species to bacteria and archaea. Consequently, this depth range is hereby referred to as the "transition zone".  Within the above classes, taxa identified at the genus and family level were investigated for potential metabolisms relevant to the geochemistry of the HZ.  When a specific metabolism (e.g. iron reduction) was identified in a species from the literature, it was assumed that this metabolism could be held by other species in the overarching genus. This was considered acceptable because, although there can be exceptions, genetic relatedness tends to imply a level of metabolic similarity [117].  Many taxa from the sediment samples remained unclassified at the genus and family levels. As a result, the metabolism of many dominant species could not be assessed, and thus the metabolisms discussed below reflect only the most-studied, culturable bacteria. For example, the Acidobacteria are the second-most abundant phylum identified in this study, but little information could be found regarding many of the dominant genera, so few metabolic processes can be surmised.  Identifiable Microbial Metabolisms  A summary of all microorganisms identified in this thesis with metabolisms recognized by prior researchers is listed at the end of this section in Table 4.2. Taxa are characterized based on the depth range at which their highest proportional abundance was found. These ranges consisted of the aerobic zone (0-30 cm depth), the transition zone (30-55 cm), and the anaerobic zone (55-85 cm).  The class of Alphaproteobacteria did not display a prominent trend with depth, although slightly higher proportional abundances were noted in the top half of the core, as shown in the Figure 4.16. In this class, three genera were found to have species with documented metabolisms. Figure 4.16 shows the distribution of Pedomicrobium, Hyphomicrobium, and Novosphingobium with depth. One species from 93  Pedomicrobium, whose abundance ranged between 0.2% and 0.4%, was found to be capable of oxidizing Fe, while two were capable of oxidizing Mn [118]. Multiple species from Hyphomicrobium, which had consistent abundance of about 1% throughout the core, were found to carry out denitrification via methanol oxidation [119]. Of pertinence to the contaminated nature of the River District site was the identification of Novosphingobium, whose abundance peaked at 0.33% in the shallowest interval. One species from this genus discovered in a South Korea estuary was found to be capable of degrading high-molecular-mass polycyclic aromatic hydrocarbons of two to five rings [120].     Proportional abundances of Betaproteobacteria were found to be constant and elevated in the upper-most 50 cm (see Figure 4.17). The genus Methylibium, part of the Burkholderiales order, had a peak abundance of 0.46%, in the upper part of the transition zone (37.5 cm). The only species known in this genus is a facultative anaerobe capable of completely degrading the gasoline additive MTBE, a persistent environment pollutant [121].  01020304050607080900% 2% 4% 6% 8% 10%α-proteobacteria RhizobialesRhodospirillalesSphingomonadalestotal 0.0% 1.0% 2.0%Hyphomicrobium 0.0% 0.5%Pedomicrobium 0.0% 0.2% 0.4%Novosphingobium Figure 4.16 – Shown on the left are the total distribution of α-proteobacteria, and the 3 most abundant classes – Rhizobiales, Rhodospirillales, and Sphingomonadales. To the right, Hyphomicrobium, Pedomicrobium, and Novosphingobium represent the 3 most abundant genera of the α-proteobacteria containing species with known metabolisms. 94      Figure 4.18 shows the distribution of Gammaproteobacteria, whose abundance increased with depth, attaining 10.8% proportional abundance at a depth of 72.5 cm. In the order Methylococcales, the genus Crenothrix was identified, and its abundance increased with depth, reaching 2.9% abundance at the deepest interval. Crenothrix is a well-known Fe and Mn oxidizer commonly responsible for well and pipe clogging [122]. Some species of Crenothrix are also believed to be able to oxidize methane [123]. Also in the order Methylococcales, the family Methylococcaceae was identified. Its abundance increased with depth, reaching 3.5% between 70 and 80 cm (see Figure 4.18). Methylococcaceae are well-known by their ability to obtain their energy from the oxidation of methane [124].  Chromatiales and Oceanospirillales, which are additional orders of Gammaproteobacteria, yielded two other genera with known metabolisms – Acidiferrobacter and Halomonas. Abundance of the genus Acidiferrobacter was highest at depths between 45 and 55 cm, at the bottom of the transition zone, reaching a proportion of 1.7%. Species of Acidiferrobacter are known to be facultatively anaerobic iron- and sulfur-oxidizers [125]. The abundance of Halomonas was found to increase with depth, but it peaked 01020304050607080900% 2% 4% 6% 8% 10% 12%β-proteobacteria BurkholderialesMethylophilalesNitrosomonadalesTRA3-20total 0.0% 0.3% 0.6%Methylibium Figure 4.17 – Shown on the left are the total distribution of β-proteobacteria, and the 4 most abundant classes – Burkholderiales, Methylophilales, Nitrosomonadales, and the candidate TRA3-20. To the right, Methylibium, represents the most abundant genus of the β-proteobacteria containing species with known metabolisms. 95  at 67.5 cm with 0.29%. Species from Halomonas have been characterized as halophilic (salt-loving) bacteria capable of reducing NO3 to NO2 [126]. The distribution of halophilic species such as those from Halomonas may be useful in determining the extent of freshwater ingression during freshet due to their predisposition to avoid such environments.      The Deltaproteobacteria increased in relative abundance with depth, reaching an abundance of 19.9% at 67.5 cm (see Figure 4.19). The class is known to contain many anaerobic sulfate- and sulfur-reducing bacteria [127]. Desulfobacterales was a major order in this class, present mainly at depth, with abundance reaching 4.9% at 72.5 cm.  This order is composed of strictly anaerobic sulfate-reducing bacteria [128].     01020304050607080900% 5% 10% 15%γ-proteobacteria ChromatialesMethylococcalesOceanospirillalestotal 0.0% 2.0% 4.0%CrenothrixMethylococcaceae0.0% 1.0% 2.0%Acidiferrobacter 0.0% 0.3%Halomonas Figure 4.18 – Shown on the left are the total distribution of γ-proteobacteria, and the 3 most abundant classes – Chromatiales, Methylococcales, and Oceanospirillales. To the right, Crenothrix, Methylococcaceae, Acidiferrobacter, and Halomonas represent the 4 most abundant genera of the γ-proteobacteria containing species with known metabolisms. 96          In the order Syntrophobacterales, two notable genera were identified – Desulfobacca and Syntrophus. Desulfobacca, an acetate-utilizing sulfate-reducer [129], had an abundance which increased with depth, peaking at 67.5 cm with 0.56%. As shown in Figure 4.19, Syntrophus was also present mainly at depth, with a peak abundance of 1.9% at 62.5 cm. Species in this genus are anaerobic, and are known to produce methane and acetate syntrophically (i.e. by utilizing products from other species) [130]. In the order 01020304050607080900.01% 0.10% 1.00% 10.00% 0% 5% 10% 15% 20% 25%DesulfarculalesDesulfobacteralesDesulfuromonadalesMyxococcalesSyntrophobacteralestotal 01020304050607080900.0% 1.0% 2.0%Syntrophus 0.0% 0.5% 1.0%Desulfobacca 0.0% 0.2% 0.4%Deferrisoma 0.0% 1.0% 2.0%Anaeromyxobacter Figure 4.19 – Shown in the top half are the total distribution of δ-proteobacteria, and the 5 most abundant classes – Desulfarculales, Desulfobacterales, Desulfuromonadales, Myxococcales, and Syntrophobacterales. On the bottom, Syntrophus, Desulfobacca, Deferrisoma, and Anaeromyxobacter represent the 4 most abundant genera of the δ-proteobacteria containing species with known metabolisms. δ-proteobacteria 97  Bdellovibrionales, the genus Deferrisoma was identified. Its abundance was higher at depth, reaching 0.2% at 77.5 cm. Deferrisoma is a dissimilatory Fe reducer which can also reduce sulfur [131]. In the order Myxococcales, the genus of Anaeromyxobacter was identified. Anaeromyxobacter's abundance increased with depth, peaking at 82.5 cm with 1.6%. One species in this genus was previously found to be a facultative anaerobe capable of reducing both Fe and NO3 [132].  In the class of Epsilonproteobacteria, the major identifiable genus was that of Sulfuricurvum, in the order of Campylobacterales. Sulfuricurvum was present only at depth, with its abundance peaking at 72.5 cm with 1.3% (see Figure 4.20). One species from this genus, isolated from an underground crude-oil storage cavity, was found to be a facultatively anaerobic sulfur-oxidizing, nitrate-reducing bacterium [133].     The genus Blastocatella was a dominant taxa from the Acidobacteria phylum. Blastocatella, an aerobic chemoorganotroph [134], had its abundance decrease with depth, peaking at 12.5 cm with 9.0% (Figure 4.21). The genus Thermoanaerobaculum, also from the phylum of Acidobacteria, showed increasing 01020304050607080900.0% 0.5% 1.0% 1.5%Sulfuricurvum Figure 4.20 – Shown above is the proportional distribution of Sulfuricurvum, a genus comprising a species found to be a facultatively anaerobic sulfur-oxidizing, nitrate-reducer. Sulfuricurvum represents the dominant genus identified from the ε-proteobacteria. 98  abundance with depth, peaking at 72.5 cm with 0.7% (Figure 4.21). Although much less dominant, this genus is comprised of species which are capable of fermentative growth and both Fe and Mn reduction [135].      In the phylum Nitrospirae, a dominant genus was Nitrospira. The proportional abundance of Nitrospira was highest in the top 40 cm, and then decreased with depth (Figure 4.22). Peak abundance was 2.4% at the shallowest sediment interval. Nitrospira contain species which are capable of oxidizing NO2 to NO3 [136].     01020304050607080900% 5% 10% 15% 20% 25% 30%Acidobacteria Subgroup_17Subgroup_3Subgroup_4Subgroup_6total 0.0% 5.0% 10.0%Blastocatella 0.0% 0.5% 1.0%Thermoanaerobaculum Figure 4.21 – Shown on the left are the total distribution of Acidobacteria, and the 4 most abundant Subgroups – 3, 4, 6, and 17. To the right, Blastocatella and Thermoanaerobaculum represent the 2 most abundant genera of the Acidobacteria containing species with known metabolisms. 99          01020304050607080900% 5% 10%Nitrospira (class) 0.0% 1.0% 2.0% 3.0%Nitrospira (genus) 01020304050607080900% 2% 4% 6%Planctomycetacia 0.0% 0.5% 1.0% 1.5%Planctomyces 0.0% 0.5% 1.0% 1.5%Pirellula 0.0% 0.5%Blastopirellula Figure 4.22 – Shown on the left is the proportional distribution of the class Nitrospira. On the right is proportional distribution of the genus Nitrospira, which is known to contain species capable of oxidizing NO2 to NO3. Figure 4.23 – Shown on the left is the proportional distribution of the phylum Planctomycetacia. To the right, Planctomyces, Pirellula, and Blastopirellula represent the 3 most abundant genera of Planctomycetacia containing species with confirmed aerobic metabolisms. 100  Three aerobic genera of the phylum Planctomycetes were identified – Blastopirellula, Pirellula, and Planctomyces [137-139]. All three had decreasing abundance with depth (Figure 4.23), confirming their aerobic nature. Blastopirellula's abundance peaked at 17.5 cm with 0.47%, Pirellula's at 17.5 cm with 1.0%, and Planctomyces' at 27.5 cm with 1.4%.  The genus Chthoniobacter was dominant in the phylum of Verrucomicrobia. Chthoniobacter was confirmed to be an aerobic heterotroph [140], and its decreasing proportional abundance confirmed this. Peak abundance for Chthoniobacter was 4.4% in the shallowest sediment interval (Figure 4.24).     In the domain of Archaea, two genera were dominant. From the class of Methanomicrobia, the genus Candidatus Methanoperedens was identified with a depth-independent abundance distribution, ranging from 1.4% to 3.4% (Figure 4.25). One species in Candidatus Methanoperedens is capable of anaerobic methane oxidation coupled to nitrate reduction [141]. Its persistence in the upper, aerobic portion of the core is thus somewhat counterintuitive. The second genus, from the class Thermoplasmata, is an 01020304050607080900.0% 1.0% 2.0% 3.0% 4.0% 5.0%Chthoniobacter Figure 4.24 – Shown above is the proportional distribution of Chthoniobacter, a dominant genus in the phylum of Verrucomicrobia. The genus Chthoniobacter contains confirmed aerobic species. 101  unknown genus of Marine Benthic Group D. Its abundance peaks at 67.5 cm with 2.6% (Figure 4.25), and is also expected to take part in the cycling of CH4.                         01020304050607080900.0% 1.0% 2.0% 3.0% 4.0%Candidatus Methanoperedens 0.0% 0.5% 1.0% 1.5% 2.0% 2.5% 3.0%Marine Benthic Group D Figure 4.25 – To the left, the proportional distribution of the genus Candidatus Methanoperedens, containing known anaerobic CH4 oxidizers, is shown. To the right, the proportional distribution of "Marine Benthic Group D" is shown. The above genera are part of the Methanomicrobia and Thermoplasmata classes, respectively, in the domain of Archaea. 102  Table 4.2 – Summary of microorganisms with known metabolisms are listed below. Taxa are categorized based on where their proportional abundance is highest; in the aerobic zone (0-30 cm depth), the transition zone (30-55 cm), or the anaerobic zone (55-85 cm). Some taxa show depth-independent abundance. Taxa listed twice are believed to be capable of multiple metabolisms. All listed taxa represent microbial genera, except for denotations (F) and (O), which represent families and orders, respectively. The abundance profiles of many taxa do not have prominent maxima. For better detail on abundance variation with depth, see Figures 4.16 to 4.25.   Aerobic Zone (0-30 cm) Transition Zone (30-55 cm) Anaerobic Zone (55-85 cm) Depth-independent Microbial Metabolism Aerobic Chthoniobacter[1], Blastocatella[2], Blastopirellula[3], Pirellula[4], Planctomyces[5]       Denitrification     Halomonas[6], Sulfuricurvum[7] Hyphomicrobium[8], Candidatus Methanoperedens[9] Nitrification Nitrospira[10]   Anaeromyxobacter[11]   Mn oxidation     Crenothrix[12] Pedomicrobium[13] Mn reduction     Thermoanaerobaculum[14]   Fe oxidation   Acidiferrobacter[15]  Crenothrix[12] Pedomicrobium[13] Fe reduction     Thermoanaerobaculum[14], Deferrisoma[16], Anaeromyxobacter[11]   H₂S or S oxidation   Acidiferrobacter[15] Sulfuricurvum[7]   SO₄ or S reduction     Desulfobacterales[17] (O), Desulfobacca[18], Deferrisoma[16]   CH₄ oxidation     Methylococcaceae[19] (F), Crenothrix[12] Candidatus Methanoperedens[9] Methanogenesis     Syntrophus[20]   PAH degradation Novosphingobium[21]       MTBE degradation   Methylibium[22]     [1]Sangwan et al. (2004), [2]Foesel et al. (2013), [3]Lee et al. (2013), [4] Glöckner et al. (2003), [5]Bauld and Staley (1976), [6]Vreeland et al. (1980), [7]Kodama and Watanabe (1994), [8]Timmermans and Van Haute (1983), [9]Haroon et al. (2013), [10]Ehrich et al. (1995), [11]Treude et al. (2003), [12]Stoecker et al. (2006), [13]Cox and Sly (1997), [14]Losey et al. (2013), [15]Hallberg et al. (2011), [16]Slobodkina et al. (2012), [17]Garrity et al. (2006), [18]Elferink et al. (1999), [19]Bowman (2006), [20]Jackson et al. (1999), [21]Sohn et al. (2004), [22]Nakatsu et al. (2006)   Sediment Mineralogy  The results of quantitative phase analysis by Rietveld refinements are given in Table 4.3. These figures represent the relative amounts of crystalline phase minerals normalized to 100% for sediment samples 103  corresponding to depths of 22.5, 37.5, 47.5, and 67.5 cm. One Rietveld refinement plot, which is representative of all four samples due to their high degree of similarity, is shown in Figure 4.26.   Table 4.3 – Results of quantitative phase analysis by Rietveld refinements are shown below. These figures represent the relative amounts of crystalline phase minerals normalized to 100%. Mineral composition throughout the core is nearly identical, confirming that sediment composition is uniform with depth. Mineral Ideal Formula 22.5cm 37.5cm 47.5cm 67.5cm Actinolite Ca2(Mg,Fe)5Si8O22(OH)2 2.5 2.4 2.7 2.0 Ankerite-Dolomite Ca(Fe2+,Mg,Mn)(CO3)2 - CaMg(CO3)2 0.7 0.6 0.6 0.5 Chamosite (Fe,Al,Mg)6(Si,Al)4O10(OH)8 4.9 4.8 4.9 4.5 Illite-Muscovite 1M K0.65Al2.0Al0.65Si3.35O10(OH)2 - KAl2(AlSi3O10)(OH)2 3.7 3.7 3.7 3.4 Illite-Muscovite 2M1 K0.65Al2.0Al0.65Si3.35O10(OH)2 - KAl2(AlSi3O10)(OH)2 3.5 3.2 3.7 3.1 K-feldspar  KAlSi3O8 4.5 4.4 4.4 4.6 Lizardite* Mg3Si2O5(OH)4 0.6 0.5 0.6 0.6 Plagioclase NaAlSi3O8 – CaAl2Si2O8 30.2 29.9 30.8 30.2 Quartz SiO2 49.4 50.4 48.6 50.9 Total  100.0 100.0 100.0 100.0    Table 4.3 shows that the composition of sand taken from throughout the core is nearly identical. Quartz and plagioclase make up the vast majority of the mineral composition, averaging 49.8% and 30.3%, respectively. Of particular interest are the compositions of actinolite (2.4% average) and chamosite (4.8%), which contain some degree of Fe, as well as ankerite-dolomite (0.6%), which can contain both Fe and Mn. These minerals are expected to provide some fraction of the Fe and Mn used by dissimilatory metal-reducing bacteria. The mineral compositions provide an explanation for the high dissolved concentrations of Fe and Mn in the HZ and offshore.  104    2Th Degrees75706560555045403530252015105Sqrt(Counts)200150100500MZ_4.raw Quartz low 49.42 %Albite low 17.01 %Albite low, calcian 13.18 %Actinolite 2.51 %Orthoclase 4.53 %Ankerite 0.71 %Illite/Muscovite 2M1 3.65 %Illite/Muscovite 1M 3.52 %Chamosite 1MIIb 4.91 %Lizardite 1T ? 0.58 %Figure 4.26 – Shown above is a Rietveld refinement plot, which is representative of the sample from a depth of 22.5 cm. Samples from depths of 37.5, 47.5, and 67.5 cm are found in Appendix G, although their spectra have a high degree of similarity to the one presented here. Results from this refinement yield mineralogical compositions listed in Table 4.3. Figures represent the relative amounts of crystalline phase normalized to 100%. The presence of lizardite, although suggested by the refinement plot, is not confirmed to a high degree of certainty. 105   Images from the scanning electron microscope (SEM) provide a qualitative assessment of potential precipitated minerals on the surface of sediment grains. A subsample of the sediment interval from a depth of 62.5 cm was examined with the SEM. This depth interval was selected because it had the highest content of amorphous Fe during sequential extraction experiments. Figure 4.27 shows an image obtained from the SEM, with its accompanying spectra (Figure 4.28) obtained from the energy-dispersive x-ray (EDX).      Figure 4.27 shows small white particles on the exterior surface of a sediment grain (area A). The lightest colour represents the highest atomic numbers visible in the spectrum which, as shown in Figure 4.28, represent iron. The estimated size of white particles in area A is in the 1-6 µm range. Further to the right Figure 4.27 – An image of a sample from the scanning electron microscope (SEM) is shown above. The sample originates from a depth of 62.5 cm, which was chosen because it was observed to have the highest amorphous content of extractable iron. Small white particles on the exterior surface of a sediment grain (area A) are visible. Light shades (i.e. white) represent the highest atomic numbers visible in the spectrum which, as shown in Figure 4.28, represent iron. The estimated size of white particles in area A is in the 1-6 µm range. Further to the right in area B, even smaller particles at similar levels of brightness can be seen, but their spatial distribution is far less dense. 106  in area B, even smaller particles at similar levels of brightness can be seen, but their spatial distribution is much less dense.   Magnetotactic bacteria are known to precipitate magnetite in round, oval formations in the size range of 35 to 120 nm [142], which is much smaller than the apparent size of Fe accumulations here. In groundwater samples rich in ferrous iron from a site in Michigan, two strains of iron-oxidizing bacteria were found to generate Fe oxides along the outside of their cell wall. The diameters of these strains and their outer iron accumulations were 0.32 µm and 0.73 µm [143] – these are much closer to the size range observed in samples here. Although it is possible that these iron accumulations are bacteriogenic, the SEM images do not seem to offer sufficient detail for this type of investigation, and are thus inconclusive.        Figure 4.28 – Shown above is a qualitative evaluation of elemental composition from "area A" in Figure 4.27. Higher peaks represent higher abundance of the given element. Since Fe is identified as the element with the highest value on the x-axis, it represents the most likely elemental candidate for the white accumulations visible in Figure 4.27. The spectrum is obtained from an energy-dispersive x-ray (EDX) installed in tandem with the SEM. 107  Interpretation  Results from porewater, sediment, and microbiological analyses suggest dynamic biogeochemistry in the HZ. Although in some cases, the biogeochemical processes implied by each method agree with each other, it remains difficult to determine whether they are intimately linked. This is because porewater is much more mobile than sediment. Since the HZ represents a flow-through regime affected by the tidal cycle, porewater is not necessarily representative of the biogeochemical processes occurring in the exact location from which it is sampled. Nonetheless, Table 4.4, located at the end of this section, provides a summary of inferred geochemical processes in the HZ based on interpretation of porewater analyses and sediment extractions.   Chloride serves as a conservative tracer to determine the amount of dilution of groundwater by river water. The concentration increase factor of 170 from 12.5 cm to 82.5 cm for both Cl and Na indicate that groundwater from the depth of 85 cm is diluted by at least two orders of magnitude prior to discharging. This falls within the range of modelling estimates by Bianchin et al. (2010), who quantified dilution of groundwater from a depth of 1.05 m to be 84% (factor of 6.25) during low tide and 99.75% (factor of 400) during high tide [2]. Since the core collected for this research was sampled just prior to high tide, these results agree with each other.  Previous sampling in the saline groundwater discharge zone at a depth of 1 m (Chapter 2) shows that specific conductance values vary between 5000 and 7000 µS/cm, and Cl concentrations vary between 1400 and 1800 mg/L. Porewater extracted from the deepest interval of the sediment core (80-85 cm) has specific conductance and chloride concentration values which are only 41% and 37%, respectively, of the above-mentioned range averages. This infers that dilution likely persists beyond a depth of 85 cm in the HZ. Furthermore, if brine exclusion were occurring during sediment freezing, it would have a greater effect on deeper porewater, which has a higher content of dissolved solutes. In this case, the true solute 108  content would be higher than what was actually measured, inferring greater dilution than estimated. For either (or both) of these reasons, the apparent groundwater dilution factor of 170 obtained from the Cl concentration gradient is a conservative estimate.     Results from porewater geochemical analyses infer oxidation of both Fe and Mn. The profiles of both Fe and Mn are a consequence of dilution as well as the low solubility of the reduced forms of these elements in the more oxygen-rich portion of the HZ. Their concentrations decrease significantly in the upwards direction. From the depth of 82.5 cm to 37.5 cm, total Fe decreases at a rate which is 9 times faster than Cl (Figure 4.29), while between 82.5 cm and 32.5 cm, Fe(II) concentrations decrease 18 times faster than Cl. The same is true for Mn, from the depth of 22.5 cm up to 7.0 cm, where concentrations decrease 21 times faster than Cl (Figure 4.29). These results infer that Fe and Mn oxidation are occurring, and this is likely to be most intensive at the depths of sharpest concentration decline. For Mn, this includes depths down to 30 cm, while for Fe, this is in the depth range of 35-50 cm. The shallower depth obtained for Mn 01020304050607080900 0.01 0.02 0.03 0.04 0.05Depth (cm) Fe / Cl mass ratio 01020304050607080900 0.1 0.2 0.3 0.4Mn / Cl mass ratio Figure 4.29 – Iron-to-chloride and manganese-to-chloride ratios (by mass) for porewater are plotted with depth. Highlighted areas represent the depths at which Fe and Mn decline at the greatest rate compared to Cl, suggesting oxidation. Reduced Mn also appears to be accumulating at depths below 25 cm despite dilution. 109  is indicative of the fact that Mn precipitation occurs at a redox value that is more positive than Fe (Figure 1.3, Chapter 1).  Although amorphous sediment fractions of Mn and porewater Mn concentrations together indicate that aerobic Mn oxidation is probable in the 10-25 cm depth range, there is less evidence to show that the same is true for Fe. At depths where oxidation of Fe is suggested by upward decreasing porewater concentrations (50-35 cm depth range), there is no significant peak in the amorphous sediment fractions of Fe. Amorphous sediment fractions of Fe gradually increase downward, however, implying that more intense Fe oxidation may be occurring at a greater depth. Furthermore, the speciation of Fe bound in amorphous minerals shows that it is essentially all Fe(III). This infers that anaerobic iron oxidation is probable at depth. A potential electron acceptor for anaerobic iron oxidation is NO3.  In previous HZ investigations below the Fraser, mineralogical evidence for oxidative iron precipitation has been elusive [34]. Prior hypotheses for the lack of an amorphous Fe peak exist. The first is that continual sediment scour removes sediments with iron precipitates before they can be sampled. The second is that oxygen is available only under complex non-equilibrium conditions, and it is reduced before it can be utilized by iron oxidation [34]. A third explanation is that large quantities of iron are anaerobically oxidized at depth before encountering significant concentrations of oxygen in shallower zones.  Crenothrix, a dominant genus whose abundance increases with depth, is a well-known Fe and Mn oxidizer. The abundance of Crenothrix follows a similar trend to the content of amorphous iron on sediment, which increases slightly in the downward direction. Crenothrix is thus a highly plausible candidate for Fe oxidation in the HZ studied here. Acidiferrobacter, furthermore, is known to be a facultatively anaerobic iron and sulfur oxidizer. Its abundance peaks in the 45-55 cm depth range, matching the zone of sharp decline in dissolved Fe concentrations. Pedomicrobium, another genus with 110  Fe and Mn oxidizers, is distributed evenly within the top 70 cm. Genera which may contribute to Fe reduction include Deferrisoma, whose abundance peaks at 77.5 cm, and Anaeromyxobacter, which peaks at a depth of 82.5 cm.  Manganese reduction also appears to be occurring in the HZ, with greatest intensity immediately below the zone of suspected Mn oxidation. Between the depths of 22.5 and 82.5 cm, Mn concentrations remain stable, varying by no more than 20%. In the same depth range, Cl concentrations decrease by an approximate factor of 90. This suggests that reductive dissolution of Mn is occurring, either in-situ or upstream of this location, possibly through the oxidation of dissolved organic matter. The only genus identified with the ability to reduce Mn is Thermoanaerobaculum, but its peak abundance occurs at a depth of 72.5 cm. As a result, it is unlikely to be responsible for the Mn reduction identified here.  Reduction and oxidation of nitrogen compounds is expected in the HZ, and data support this notion. NO3 concentrations decrease with depth in the upper 35 cm by a factor of 5, inferring NO3 reduction (i.e. denitrification). At depths below 55 cm, however, NO3 concentrations are higher, indicating that sample oxidation may have occurred in the time prior to analysis. Although NO3 cannot be generated anaerobically, anaerobic oxidation of ammonia to N2 (i.e. anammox) via nitrate reduction is possible [144]. Nitrospira, a dominant genus with species that oxidize NO2 to NO3, are found to be quite abundant in the top 40 cm. Nitrospira is thus expected to play an important role in the nitrogen cycle with other facultative anaerobes in the upper HZ. Other identified genera known to have some capacity for denitrification include Hyphomicrobium (uniform distribution with depth), Halomonas (peak abundance at 67.5 cm), and Sulfuricurvum (peak abundance at 72.5 cm).  Since SO4 is not detected in saline groundwater from the monitoring wells, and since H2S is qualitatively detectable in upwelling groundwater via smell, sulfide oxidation must be occurring in the HZ. 111  Furthermore, sequencing data strongly support the notion that both sulfate reducers and sulfide oxidizers are present in significant numbers. Desulfobacterales, a class of strictly anaerobic sulfate-reducers, has an increasing abundance with depth, peaking at 72.5 cm. Other genera with similar abundance profiles include Desulfobacca, a sulfate-reducer, and Deferrisoma, a metal and sulfur reducer.  At the depth of 52.5 cm, a large peak in SO4 is noted, and this is attributed to sulfide oxidation. Acidiferrobacter, which comprises species that can oxidize Fe as well as S, has a peak abundance in the depth range of 45-55 cm, which encompasses the aforementioned SO4 peak. Sulfuricurvum, a genus of the Epsilonproteobacteria known to oxidize sulfide via NO3 reduction, has an increasing abundance with depth, and likely takes part in the oxidation of upwelling H2S.  Although methane concentrations are not measured, sequencing results also support the presence of both methanogenic and methane-oxidizing microorganisms. Candidatus Methanoperedens is identified with depth-independent abundance. This genus couples CH4 oxidation to NO3 reduction. The family of Methylococcaceae, furthermore, which consists of many CH4 oxidizers, has high proportional abundance at depth, especially in the range of 70-80 cm. In addition to oxidizing Fe and Mn, some species of Crenothrix are also believed to be capable of oxidizing CH4. The genus Syntrophus is a primary candidate at depth to take part in methanogenesis. Species in this genus are known to have methanogenic metabolisms which depend on the products of other species with which they cohabit.   The sediment interval between the depths of 50 and 55 cm demonstrates the greatest amount of deviations from vertical trends in geochemistry. This may represent a hotspot of (bio)geochemical activity. At the interval midpoint of 52.5 cm, SO4 concentrations peak notably, while a decline in NO3 concentrations is apparent (Figure 4.3). Additionally, the proportion of total Fe contributed by Fe(II) is lower at the depth of 52.5 cm (50%), compared to the intervals just above (87%) and below (80%). Lastly, DO is measured to be zero (Figure 4.6), and a local drop in pH is also observed (Figure 4.5). 112  These localized deviations may be explained by any combination of sulfide oxidation, iron oxidation, nitrate reduction, and ion exchange-driven calcite precipitation. The fact that more than one of these processes may be occurring at close proximity seems to imply that redox microzone theory is more relevant than simple redox zonation with depth.  Table 4.4 – Summary of inferred geochemical processes in the HZ are identified based on results from porewater analyses and sediment extractions. Processes are listed in the depth range at which they are believed to occur; in the aerobic zone (0-30 cm depth), the transition zone (30-55 cm), or the anaerobic zone (55-85 cm). References to figures showing relevant data are included.  Aerobic Zone (0-30 cm) Transition Zone (30-55 cm) Anaerobic Zone (55-85 cm) Denitrification  Downward decrease in NO₃ concentrations from river bed (Figure 4.3)   Mn oxidation Upward decline in porewater Mn concentrations (Figure 4.2, Figure 4.29), peak in amorphous Mn at 17.5 cm (Figure 4.10)     Mn reduction   Mn concentrations not varying with depth despite dilution (Figure 4.2, Figure 4.29)   Fe oxidation   Upward decline in porewater concentrations (Figure 4.2, Figure 4.29) Speciation of amorphous Fe on sediment grains is 98% Fe(III) (Figure 4.12) H₂S oxidation   Localized peak in SO₄ at 52.5 cm (Figure 4.3)   SO₄ reduction    Downward decrease in SO₄ concentrations from river bed (Figure 4.3)   4.4 Conclusion  Major Microbial Metabolisms and Geochemical Processes  Table 4.2 presents a summary of potential microbial metabolisms in the HZ, while Table 4.4 shows a summary of inferred geochemical processes. Results suggest that denitrification is occurring in both the aerobic and transitions zones of the HZ. Data strongly support the notion that Mn oxidation and reduction are occurring as well. Both aerobic and anaerobic Fe oxidation are implied, and this is supported by the identification of Fe-oxidizing microorganisms. Sulfide oxidation is suggested in the transition and 113  anaerobic zones, while sulfate reduction is most apparent in the anaerobic zone. Methane oxidation is also inferred in the anaerobic zone.  Defining the Aerobic Portion of the Hyporheic Zone  In examining the abundance profiles of known aerobic genera, part the HZ could be delineated as aerobic itself. Seven classes of bacteria demonstrated peak proportional abundance above the depth of 30 cm – these were the Acidobacteria, the Sphingobacteriia, the Betaproteobacteria, Planctomycetacia, the Spartobacteria, Verrucomicrobiae, and the Flavobacteriia. Within classes, however, metabolisms can vary widely, and thus it was more useful to examine genera with known aerobic species. Blastocatella was the most abundant identified genus known to be aerobic, with a peak abundance of 9.0% at a depth of 12.5 cm. Chthoniobacter, another aerobic heterotroph, had peak abundance at a depth of 7.0 cm. Three other genera comprising known aerobic species included Blastopirellula (peak abundance at 17.5 cm), Pirellula (17.5 cm), and Planctomyces (27.5 cm).  This suggested that the vast majority of oxygen-utilizing species at the time of sampling were found in the top 30 cm of sediment. In order to delineate the aerobic portion of the HZ, geochemical data were also considered. Since Fe(II) was non-detectable above the depth of 30 cm, this depth was confirmed to serve as an adequate delineation of the aerobic HZ. Despite this delineation, it is possible that some aerobes are still present at depths greater than 30 cm. Furthermore, oxygen dependence is not a strictly dualistic trait. Across a range of microorganisms, oxygen dependence manifests itself in a continuum, as can been seen with facultative anaerobes and microaerophiles. Since some of the aerobic microorganisms identified in the top 30 cm have the potential to be facultatively anaerobic, and since Fe(II) may also undergo oxidative precipitation via NO3 reduction, the delineation of the aerobic HZ is more likely to be an overestimation than an underestimation. 114  Between the depths of 30 and 55 cm, a transition zone was observed. In this zone, there was a considerable shift in major ion chemistry, as well a rapid increase in Fe(II) concentrations. Increased variability in both pH and alkalinity were also observed here, which may have been indicative of oxidative iron precipitation or ion exchange-driven calcite precipitation. By hosting such rapid changes in porewater geochemistry, this transition zone offers the widest variety of redox species to microorganisms, and this may serve as a competitive advantage. A potential challenge of inhabiting the transition zone, however, would be greater seasonal variations in salinity due to freshet-driven dilution.  Implications for Discharging Contaminants  Aerobic environments yield the highest rates of biodegradation for the majority of organic contaminants. Since the top 30 cm of the HZ was identified as aerobic at the time of sampling, this depth range would provide the greatest capacity for biodegradation of upwelling contaminants. Natural attenuation, however, also includes physical factors such as dilution. From a depth of 85 cm to the surface, the distribution of chloride concentrations infer a dilution rate greater than two orders of magnitude.  One particular genus of interest identified in the aerobic zone was Novosphingobium. A species from this genus, isolated from an estuarine environment, was found to be capable of degrading PAHs with a size range of 2 to 5 rings. The metabolic potential of Novosphingobium could be of substantial importance if contaminated groundwater from the River District site is allowed to flow towards the river following a planned shut-off of on-site monitoring wells.   115  Chapter 5 – Summary and Recommendations  5.1 The River District Site  At the River District site, groundwater flows horizontally towards the river at a velocity of 0.1 to 0.2 m/day [55]. In shallow portions of the aquifer, fresh groundwater is found down to a depth of approximately 10 m, below which a transition to saline groundwater is observed. At depths below 13 m, groundwater is entirely saline. As groundwater approaches the river, it turns upward and discharges into the HZ.   A dominant source of PAH contamination exists at the base of the aquifer, but shallow, localized contamination is also present. Pumping wells are installed to generate capture zones, which mitigate the offsite migration of dissolved PAHs. It is expected that the operation of these pumping wells affects the natural groundwater flow regime to some extent. For the majority of findings presented in this thesis, however, these effects are not considered to be significant, since geochemical datasets from the River District show similarity to a nearby site not under the influence of pumping wells.  5.2 Characterization of Groundwater Discharge to the Fraser River  In Chapter 2, a conceptual model is established to help characterize groundwater discharge to the Fraser River. In the river channel, fresh groundwater discharges to the HZ at distances between 70 and 85 m from the shoreline's high-water mark (HWM), while the deeper, saline groundwater discharges at distances greater than 100 m (Figure 2.9). The interface between the fresh and saline bodies of groundwater, considered a halocline, is found to intersect the HZ between the distances of 85 and 100 m from the HWM. The chemistry of shallow onshore groundwater and nearshore HZ samples is similar, 116  while deep onshore groundwater and central-channel HZ samples are also similar. Figure 2.15 in Chapter 2 shows the anticipated flow paths that conform to these findings.  A proposed monitored natural attenuation (MNA) strategy at the River District site calls for the shutdown of capture wells, which would allow natural groundwater flow to re-establish itself. Contaminants dissolved in groundwater would once again flow towards the river. Central to the MNA strategy would be to track the fate of contaminants as they flow towards the river. The discharge pattern characterized in this thesis will thus be of great value in guiding future monitoring efforts.   At the onshore halocline separating fresh and saline groundwater, the exchange of sorbed Ca by Na is most likely responsible for Ca peaks in porewater (Figure 2.13). These cation exchange effects occur because of transverse mixing along the horizontal flow path towards the river. Peaks for Fe and Mn found at similar depths (Figures 2.16 and 2.19) are believed to be driven by either the anaerobic oxidation of organic matter (OM) or reduced gases, which themselves are likely generated by OM oxidation in silt lenses or redox microzones.  From the intertidal monitoring wells out to the HZ, Fe and Mn concentrations are found to increase despite dilution in the HZ. This is believed to be a result of either desorption of Fe and Mn due to cation exchange, or oxidation of OM, H2S, or CH4. Although one or more of these processes likely take place along the flow path, some of them may also be occurring in the HZ.  5.3 Design and Development of a Sediment-Freezing Sampler  Chapter 3 presents a summary of the design and development of a new sediment-freezing sampler. Our objective is to collect representative, high-quality sediment samples from the HZ for biogeochemical characterization. Samples are collected with a custom-made cryogenic probe, or "popsicle sampler", 117  which is driven into the sediments with a slide hammer. The controlled expansion of liquid CO2 is used to rapidly cool the steel sampling pipe, to which sediments gradually freeze.  Preservation quality is excellent (Figure 3.9), and there is a high level of confidence that samples are representative of in-situ conditions. Contamination of HZ porewater by river water during sample retrieval through the surface water column is not observed to be significant (Figure 3.12). The "freeze anchoring" effect is a major challenge encountered during sample collection, even with the use of a motorized winch. Future tests are planned to assess the effectiveness of a sonic vibration attachment to reduce friction and aid in sampler insertion and retrieval.  The research potential of the sampler is significant. The sampler could improve the sediment sampling process for many fields, including hydrology, sedimentology, and benthic/hyporheic ecology. Hydrologists would be interested in the sampler's ability to preserve a snapshot of porewater chemistry in time, thus aiding in determining dilution based on the concentration of conservative species. Multiple core samples taken over a given period would allow for a time-lapse analysis of chemical changes in dynamic fluvial or tidal environments. Sedimentologists, additionally, would be interested the sampler's ability to preserve the sediment structure as laid out during deposition at a reliably fine scale. Aquatic ecologists, furthermore, would be interested in the sampler's ability to capture and preserve invertebrates and microbes. This method of sample collection would directly improve the characterizations of biodiversity and preserve DNA for sequencing. A key advantage of the sampler over other methods is the relative ease of collection in deep water from a relatively small boat. Samples analyzed here are collected from a depth of about 10 m, but sampling depth is only theoretically limited by the length of utility lines and cables.    118  5.4 Biogeochemical Characterization of Hyporheic Zone Sediments  Chapter 4 consists of a comprehensive vertical characterization of biogeochemistry in the HZ offshore of the River District site. Results suggest dynamic biogeochemistry in the HZ, as evidenced by the identification of many redox-sensitive species. Of great importance in this investigation is the recognition that microbiology serves as a useful indicator for chemical and physical processes in the HZ. Although a sediment core captures just a snapshot of temporally varying geochemistry, microbiology helps to constrain variability over time. Due to the largely sessile nature of DNA in the HZ, microbiology effectively represents a long-term average of environmental conditions.  Chloride is used as a conservative tracer to determine the amount of dilution of groundwater by river water. A conservation dilution factor of 170 is obtained by comparing Cl concentrations between the top and bottom of the core (Figure 4.1). The lack of a concentration plateau at the bottom of the core suggests that dilution persists beyond the depth of 85 cm in the HZ. This notion is supported by modelling estimates by Bianchin et al. (2010), who quantify dilution of groundwater from a depth of 1.05 m to be 84% (factor of 6.25) during low tide and 99.75% (factor of 400) during high tide. Since the 0.85 m core collected for this research is sampled just prior to high tide, and a dilution factor of 170 is obtained, these results are in agreement.  Sharply decreasing porewater concentrations of Fe and Mn in the upward direction, even after accounting for dilution, suggest that oxidation of these metals occurs to some extent (Figures 4.2 and 4.29). Mn oxidation is believed to be occurring between the depths of 15 and 25 cm, while Fe oxidation is probable between the depths of 35 and 50 cm.  The abundance of the genus Acidiferrobacter, a facultatively anaerobic iron oxidizer, peaks in the 45-55 cm depth range, and the genus is thus considered a plausible candidate for Fe oxidation at this depth 119  (Figure 4.18). The genus Crenothrix, a well-known Fe-oxidizer, is also considered to be responsible for Fe oxidation in the HZ studied here, since its abundance follows a similar vertical trend to the content of amorphous iron extracted from sediment samples (Figures 4.9 and 4.18).  Although geochemical evidence for Fe reduction in porewater extracted from the core is not resounding, the identification of Fe-reducers Deferrisoma and Anaeromyxobacter, whose abundance increase with depth (Figure 4.19), is consistent with Fe reduction in the HZ. Mn concentration patterns also suggest that it is being reduced in the HZ, with greatest intensity immediately below the zone of suspected Mn oxidation. Below the depths of 22.5 cm, Mn concentrations remain stable, varying by no more than 20%, while Cl concentrations decrease in the upward direction by an approximate factor of 90 (Figures 4.2 and 4.3). In this depth range, dissolved Mn appears to be produced at a rate sufficient enough to counteract the effects of dilution. No obvious taxa for Mn reduction are identified during the DNA sequencing process. Although these data do suggest Fe and Mn reduction, it is also possible that the concentrations trends of the elements are partially affected by ion exchange in the HZ.  NO3 concentrations decrease with depth in the uppermost 35 cm, suggesting that denitrification is occurring in the HZ (Figure 4.3). Additionally, Nitrospira is identified as a potential genus responsible for nitrification, and its abundance is proportionally highest in the upper half of the core (Figure 4.22).  Results from DNA sequencing suggest that both sulfate reduction and sulfide oxidation are occurring in the HZ. The class Desulfobacterales, as well as the genera Desulfobacca and Deferrisoma, are all identified in the HZ. With peak abundances at greater depths, these taxa are identified as prime candidates for the reduction of sulfate and sulfur (Figure 4.19). Acidiferrobacter, a genus also possibly responsible for Fe oxidation, is considered a candidate for sulfide oxidation at the depth of 52.5 cm, where a localized peak of SO4 is noted.  120  Although CH4 concentrations are not measured, methanogenic and methane-oxidizing microorganisms are identified in the HZ. The genus Candidatus Methanoperedens and the family of Methylococcaceae are identified as plausible CH4 oxidizers (Figure 4.25). The genus Syntrophus, moreover, is a possible candidate for methane generation in the deepest portions of the HZ.  The fact that many geochemical processes are implied to exist within close proximity in the transition zone and anaerobic zone seems to imply that the classical understanding of the redox ladder is less valid. Although some redox zonation is expected to occur with depth due to the downwelling of oxygen, redox microzone theory seems to be much more accurate in explaining how such overlap in biogeochemical processes can occur.  5.5 Delineation of the Aerobic Hyporheic Zone and its Implications  Since multiple known aerobic taxa are found to have highest proportional abundances in the top 30 cm of the HZ, this depth range is considered to represent the portion of the HZ which is continually exposed to significant amounts of oxygen from the river. Porewater geochemistry also supports this delineation, as Fe(II) is non-detectable above the depth of 30 cm.   The implications of this delineation are perhaps most noteworthy for the fate of discharging contaminants. Since aerobic environments tend to be most favourable for contaminant biodegradation, dissolved PAHs at a concentration above the environmental guideline have a good chance of undergoing some degree of biodegradation as they pass through the aerobic HZ. However, it is also important that the residence time of contaminants be sufficient such that degradation actually occurs.  The modelling efforts of Bianchin et al. (2010) yield a solute residence time of approximately 58 days for the top metre of the HZ [2]. For comparison, a rough estimation of residence time in the HZ can be 121  determined based on knowledge of the following parameters: hydraulic conductivity (K), hydraulic gradient (i), and porosity (n). For the Fraser River sands, K is estimated to be on the order of 1 x 10-4 m/s and n is estimated to be 0.30 [145]. The gradient, i, is much more difficult to estimate in the HZ due to tidal oscillations. Bianchin et al. (2010), however, determine that the average vertical gradient in the first 2.8 m of sediment below the river at a nearby site oscillated between 0.05 (discharge) and -0.10 (recharge) during the 2005 freshet [2]. An initial estimate of i = 0.01 (discharge) is thus used for an annual average, with values of 0.1 and 0.001 being used to check sensitivity. The specific discharge (q), vertical groundwater velocity (v), and vertical residence time (t) over a depth (d) of 2.8 m can be calculated with Equations 5.1, 5.2, and 5.3 respectively.   𝑞 = 𝐾𝑚 ?̅? = 𝑞 𝑚⁄  𝑡 =  𝑑 ?̅?⁄    For i = 0.01, vertical residence time t equals 10 days, while for i = 0.1 and 0.001, t = 23 hours and 98 days, respectively. Since the hydraulic conductivity of unconsolidated sand (i.e. river sediment) is likely higher than that of consolidated sand, actual residence times may be lower than the estimated values.  The estimation of residence time in the HZ, the delineation of the aerobic portion of the HZ, and the characterization of dilution rates in Chapter 4 provide valuable information on the biogeochemical and physical components of natural attenuation in the HZ.  5.6 Direction of Research and Recommendations  The work done in this thesis to characterize the biogeochemistry of the HZ serves as a valuable starting point for many other potential areas of research. For example, to test whether the hypothesized processes are consistent with data, a process-based HZ model that includes both physical and biogeochemical (5.1) (5.2) (5.3) 122  parameters could be developed. Reactive transport modelling could make use of physical and geochemical data, whereas the relative abundance of certain microbial taxa with known metabolisms could serve as a starting point for Monod or Michaelis-Menten kinetics.   Now that DNA from the HZ has been sampled and sequenced, the dynamics of the microbial community and its variability with depth are much better understood. From here, further extractions of RNA and proteins can be undertaken to provide insight into the most active metabolisms. It would be most valuable to perform such investigations before, during, and after freshet to see how active microbial metabolisms transform when exposure to oxygen changes significantly. Additionally, if such characterizations are completed prior to the shut-off of wells at the River District site, an environmental baseline serving as an "uncontaminated" control would be obtained. This would be essential if dissolved PAHs migrate all the way to the HZ at detectable levels. By comparing the microbial community and prominent metabolisms before and after exposure to dissolved PAHs, valuable information would be obtained about which species are indeed capable of biodegradation.  To further assess natural attenuation potential in the HZ, in- or ex-situ experiments could be implemented to assess the manner in which dissolved PAHs degrade. Although ex-situ experiments involving a core extracted from the HZ would allow for better monitoring and high-frequency data collection, controlling such experiments so that they are actually representative of in-situ conditions would present a major challenge. Of great interest would be monitoring the growth and adaption of species capable of degrading PAHs, such as the genus Novosphingobium, which is identified in the aerobic portion of the HZ offshore of the River District site.  An in-situ experiment, involving diffusion or peeper samplers, would likely provide data which are far less biased. These could include injecting small amounts of PAHs into the HZ, and then monitoring the rate at which they attenuate, all the while monitoring for signs of biodegradation. Logistically, this 123  approach would be a greater challenge. Deploying equipment below the Fraser River and retrieving it at a later time would be extremely difficult due to the dynamic nature of the Fraser, as well as the presence of organic and industrial debris on the riverbed.  Aside from the context of natural attenuation of contaminants, there is still more to be understood about the nature of the HZ in the Fraser. The biogeochemical characterization presented in this thesis only focuses on the zone of saline groundwater discharge, while fresh groundwater discharge is still occurring at distances between 70 and 85 m from the HWM. Since the geochemistry of fresh groundwater is entirely different, the biogeochemistry of the HZ where freshwater is discharging would also be expectedly different. For example, the much lower concentration of H2S in fresh groundwater would imply that sulfur oxidation in the fresh groundwater discharge zone would be severely inhibited. Additionally, the presence of halophilic microorganisms which require high-salinity environments would be suppressed. The outcome of these differences in terms of biogeochemistry has yet to be assessed.  Although our understanding of HZ biogeochemistry is still in its infancy, the HZ of the Fraser River has several unique qualities which implore an astounding number of research questions. The influences of freshet, tidal oscillations, sediment erosion and deposition, and variable groundwater geochemistry all contribute to a massively dynamic system. Studying the effects of all of these factors will only improve our understanding of biogeochemistry in the HZ, as we compare findings to other HZs being investigated, all the while elucidating important conclusions for the field of contaminant hydrogeology.   124  References  1. Bianchin, M., L. Smith, and R. Beckie, Defining the hyporheic zone in a large tidally influenced river. Journal of Hydrology, 2011. 406(1): p. 16-29. 2. Bianchin, M., L. Smith, and R. Beckie, Quantifying hyporheic exchange in a tidal river using temperature time series. Water Resources Research, 2010. 46(7). 3. Fischer, H., et al., A river's liver–microbial processes within the hyporheic zone of a large lowland river. Biogeochemistry, 2005. 76(2): p. 349-371. 4. Boano, F., et al., Hyporheic flow and transport processes: Mechanisms, models, and biogeochemical implications. Reviews of Geophysics, 2014. 52(4): p. 603-679. 5. Boulton, A.J., et al., The functional significance of the hyporheic zone in streams and rivers. Annual Review of Ecology and Systematics, 1998: p. 59-81. 6. Storey, R., R. Fulthorpe, and D. Williams, Perspectives and predictions on the microbial ecology of the hyporheic zone. Freshwater Biology, 1999. 41(1): p. 119-130. 7. Febria, C.M., et al., Bacterial community dynamics in the hyporheic zone of an intermittent stream. The ISME journal, 2012. 6(5): p. 1078-1088. 8. Hedin, L.O., et al., Thermodynamic Constraints On Nitrogentransformations And Other Biogeochemicalprocesses At Soil-Stream Interfaces. Ecology, 1998. 79(2): p. 684-703. 9. Nogaro, G., et al., Influence of hyporheic zone characteristics on the structure and activity of microbial assemblages. Freshwater Biology, 2013. 58(12): p. 2567-2583. 10. Gandy, C., J. Smith, and A. Jarvis, Attenuation of mining-derived pollutants in the hyporheic zone: a review. Science of the Total Environment, 2007. 373(2): p. 435-446. 11. Stanford, J. and J. Ward, The hyporheic habitat of river ecosystems. 1988. 12. Findlay, S.E., et al., Metabolic and structural response of hyporheic microbial communities to variations in supply of dissolved organic matter. Limnology and oceanography, 2003. 48(4): p. 1608-1617. 13. Lenting, N., D.D. Williams, and B.G. Fraser, Qualitative differences in interstitial organic matter and their effect on hyporheic colonisation. Hydrobiologia, 1997. 344(1-3): p. 19-26. 14. Jones, J.B. and R.M. Holmes, Surface-subsurface interactions in stream ecosystems. Trends in Ecology & Evolution, 1996. 11(6): p. 239-242. 15. Tesoriero, A.J., H. Liebscher, and S.E. Cox, Mechanism and rate of denitrification in an agricultural watershed: Electron and mass balance along groundwater flow paths. Water Resources Research, 2000. 36(6): p. 1545-1559. 16. Hill, A., The potential role of in-stream and hyporheic environments as buffer zones. Buffer zones, 1996: p. 115. 125  17. Landmeyer, J.E., et al., MTBE, TBA, and TAME attenuation in diverse hyporheic zones. Ground Water, 2010. 48(1): p. 30-41. 18. Fuller, C.C. and J.W. Harvey, Reactive uptake of trace metals in the hyporheic zone of a mining-contaminated stream, Pinal Creek, Arizona. Environmental science & technology, 2000. 34(7): p. 1150-1155. 19. Hester, E., K. Young, and M. Widdowson, Mixing of surface and groundwater induced by riverbed dunes: Implications for hyporheic zone definitions and pollutant reactions. Water Resources Research, 2013. 49(9): p. 5221-5237. 20. Käser, D., A. Binley, and A. Heathwaite, On the importance of considering channel microforms in groundwater models of hyporheic exchange. River Research and Applications, 2013. 29(4): p. 528-535. 21. Sawyer, A.H., M. Bayani Cardenas, and J. Buttles, Hyporheic exchange due to channel‐spanning logs. Water Resources Research, 2011. 47(8). 22. Bradley, R., et al., Flow and sediment suspension events over low‐angle dunes: Fraser Estuary, Canada. Journal of Geophysical Research: Earth Surface, 2013. 118(3): p. 1693-1709. 23. Villard, P. and M. Church, Dunes and associated sand transport in a tidally influenced sand-bed channel: Fraser River, British Columbia. Canadian Journal of Earth Sciences, 2003. 40(1): p. 115-130. 24. Ahmerkamp, S., et al., The impact of bedform migration on benthic oxygen fluxes. Journal of Geophysical Research: Biogeosciences, 2015. 120(11): p. 2229-2242. 25. Packman, A.I., et al., Development of layered sediment structure and its effects on pore water transport and hyporheic exchange. Water, Air, & Soil Pollution: Focus, 2006. 6(5-6): p. 433-442. 26. Harvey, J.W. and C.C. Fuller, Effect of enhanced manganese oxidation in the hyporheic zone on basin‐scale geochemical mass balance. Water Resources Research, 1998. 34(4): p. 623-636. 27. Charette, M.A. and E.R. Sholkovitz, Oxidative precipitation of groundwater‐derived ferrous iron in the subterranean estuary of a coastal bay. Geophysical Research Letters, 2002. 29(10): p. 85-1-85-4. 28. Gan, P., et al., Sampling methods to determine the spatial gradients and flux of arsenic at a groundwater seepage zone. Environmental toxicology and chemistry, 2006. 25(6): p. 1487-1495. 29. Benner, S.G., E.W. Smart, and J.N. Moore, Metal behavior during surface-groundwater interaction, Silver Bow Creek, Montana. Environmental science & technology, 1995. 29(7): p. 1789-1795. 30. Spiteri, C., et al., pH-Dependent iron oxide precipitation in a subterranean estuary. Journal of Geochemical Exploration, 2006. 88(1): p. 399-403. 31. Zobrist, J., et al., Mobilization of arsenite by dissimilatory reduction of adsorbed arsenate. Environmental science & technology, 2000. 34(22): p. 4747-4753. 126  32. Nagorski, S.A. and J.N. Moore, Arsenic mobilization in the hyporheic zone of a contaminated stream. Water Resources Research, 1999. 35(11): p. 3441-3450. 33. Bolton, M. and R. Beckie, Aqueous and mineralogical analysis of arsenic in the reduced, circumneutral groundwater and sediments of the lower Fraser River delta, British Columbia, Canada. Applied geochemistry, 2011. 26(4): p. 458-469. 34. Roschinski, T.G., Geochemistry in the hyporheic zone of the lower Fraser river. 2007. 35. Briggs, M.A., et al., A physical explanation for the development of redox microzones in hyporheic flow. Geophysical Research Letters, 2015. 36. Zarnetske, J.P., et al., Coupled transport and reaction kinetics control the nitrate source‐sink function of hyporheic zones. Water Resources Research, 2012. 48(11). 37. Appelo CAJ, P.D., Geochemistry, Groundwater and Pollution. Fifth ed2005, Amsterdam: A.A. Balkema Publishers. 38. Scott, M.J. and J.J. Morgan. Energetics and conservative properties of redox systems. in ACS symposium series. 1990. Oxford University Press. 39. Langmuir, D., Aqueous Environmental Geochemistry1997, New Jersey: Prentice-Hall. 40. Paerl, H. and J. Pinckney, A mini-review of microbial consortia: their roles in aquatic production and biogeochemical cycling. Microbial Ecology, 1996. 31(3): p. 225-247. 41. Davey, M.E. and G.A. O'toole, Microbial biofilms: from ecology to molecular genetics. Microbiology and molecular biology reviews, 2000. 64(4): p. 847-867. 42. Hall-Stoodley, L., J.W. Costerton, and P. Stoodley, Bacterial biofilms: from the natural environment to infectious diseases. Nature Reviews Microbiology, 2004. 2(2): p. 95-108. 43. Flemming, H.-C., Sorption sites in biofilms. Water Science and Technology, 1995. 32(8): p. 27-33. 44. Gadd, G.M., Biosorption: critical review of scientific rationale, environmental importance and significance for pollution treatment. Journal of Chemical Technology and Biotechnology, 2009. 84(1): p. 13-28. 45. Headley, J.V., et al., Rates of sorption and partitioning of contaminants in river biofilm. Environmental science & technology, 1998. 32(24): p. 3968-3973. 46. Gan, J., et al., Surface amendment of fertilizer ammonium thiosulfate to reduce methyl bromide emission from soil. Environmental science & technology, 1998. 32(16): p. 2438-2441. 47. Suarez, M.P. and H.S. Rifai, Biodegradation rates for fuel hydrocarbons and chlorinated solvents in groundwater. Bioremediation Journal, 1999. 3(4): p. 337-362. 48. Mihelcic, J.R. and R.G. Luthy, Degradation of polycyclic aromatic hydrocarbon compounds under various redox conditions in soil-water systems. Applied and Environmental Microbiology, 1988. 54(5): p. 1182-1187. 127  49. Rockne, K.J. and S.E. Strand, Biodegradation of bicyclic and polycyclic aromatic hydrocarbons in anaerobic enrichments. Environmental science & technology, 1998. 32(24): p. 3962-3967. 50. EPA., Use of monitored natural attenuation at superfund, RCRA corrective action, and underground storage tank sites. United States Environmental Protection Agency, Washington, 1999. 51. Jørgensen, K.S., J.M. Salminen, and K. Björklöf, Monitored natural attenuation. Bioremediation: Methods and Protocols, 2010: p. 217-233. 52. White, D.C., et al., In situ microbial ecology for quantitative appraisal, monitoring, and risk assessment of pollution remediation in soils, the subsurface, the rhizosphere and in biofilms. Journal of Microbiological Methods, 1998. 32(2): p. 93-105. 53. Venosa, A.D. and X. Zhu, Biodegradation of crude oil contaminating marine shorelines and freshwater wetlands. Spill Science & Technology Bulletin, 2003. 8(2): p. 163-178. 54. Contaminated Sites Regulation. Environmental Management Act, 1996, BC Ministry of Environment: Canada. 55. Golder Associates Limited, Detailed Site Investigation, Risk Assessment and Remedial Plan, 2003: Vancouver, BC. 56. Mueller, J.G., P. Chapman, and P. Pritchard, Action of a fluoranthene-utilizing bacterial community on polycyclic aromatic hydrocarbon components of creosote. Applied and Environmental Microbiology, 1989. 55(12): p. 3085-3090. 57. Zapf-Gilje, R., G.C. Patrick, and R. McLenehan, Overview of the remediation process at sites with creosote related contamination in soil, groundwater and river sediment. Canadian Journal of Civil Engineering, 2001. 28(S1): p. 141-154. 58. Patty, F.A., Industrial Hygiene and Toxicology. Vol. I. General Principles. Industrial Hygiene and Toxicology. 2nd Revised Edition. Vol. I. General Principles., 1958. 1(2nd Revised Edition). 59. Neilson-Welch, L. and L. Smith, Saline water intrusion adjacent to the Fraser River, Richmond, British Columbia. Canadian geotechnical journal, 2001. 38(1): p. 67-82. 60. Jia, K., Iron and Manganese Reduction Driven by Organic Matter and Mixing of Fresh and Saline Groundwater in the Fraser River Delta Aquifer, Vancouver, Canada. Retrospective Theses and Dissertations, 1919-2015, 2015. 61. Ages, A. and A. Woollard, The tides in the Fraser River estuary. Environment Canada, Institute of Ocean Sciences, Sidney, BC, Report, 1976: p. 76-5. 62. Simpson, G. and I. Hutcheon, Pore-water chemistry and diagenesis of the modern Fraser River delta. Journal of Sedimentary Research, 1995. 65(4a): p. 648-655. 63. Neilson-Welch, L.A., Saline water intrusion from the fraser river estuary. A hydrogeological investigation using field chemical data and a density-dependent groundwater flow model, 1999, University of British Columbia. 128  64. Bridger, D. and D. Allen, An investigation into the effects of diffusion on salinity distribution beneath the Fraser River Delta, Canada. Hydrogeology Journal, 2006. 14(8): p. 1423-1442. 65. Chapelle, F.H., Groundwater geochemistry and calcite cementation of the Aquia aquifer in southern Maryland. Water Resources Research, 1983. 19(2): p. 545-558. 66. Herszage, J. and M. dos Santos Afonso, Mechanism of hydrogen sulfide oxidation by manganese (IV) oxide in aqueous solutions. Langmuir, 2003. 19(23): p. 9684-9692. 67. Yao, W. and F.J. Millero, Oxidation of hydrogen sulfide by hydrous Fe (III) oxides in seawater. Marine Chemistry, 1996. 52(1): p. 1-16. 68. Devol, A.H., Methane oxidation rates in the anaerobic sediments of Saanich Inlet. Limnology and oceanography, 1983. 28(4): p. 738-742. 69. Devol, A., et al., A model for coupled sulfate reduction and methane oxidation in the sediments of Saanich Inlet. Geochimica et Cosmochimica Acta, 1984. 48(5): p. 993-1004. 70. Reeburgh, W.S., Oceanic methane biogeochemistry. Chemical Reviews, 2007. 107(2): p. 486-513. 71. Murphy, F. and W. Herkelrath, A sample‐freezing drive shoe for a wire line piston core sampler. Groundwater Monitoring & Remediation, 1996. 16(3): p. 86-90. 72. Starr, R.C. and R.A. Ingleton, A new method for collecting core samples without a drilling rig. Groundwater Monitoring & Remediation, 1992. 12(1): p. 91-95. 73. Bianchin, M., L. Smith, and R. Beckie, Freeze Shoe Sampler for the Collection of Hyporheic Zone Sediments and Porewater. Groundwater, 2014. 74. Hofmann, B., D. Sego, and P. Robertson, In situ ground freezing to obtain undisturbed samples of loose sand. Journal of geotechnical and geoenvironmental engineering, 2000. 126(11): p. 979-989. 75. Stocker, Z. and D.D. Williams, A freezing core method for describing the vertical distribution of sediments in a streambed. Limnology and oceanography, 1972. 17(1): p. 136-138. 76. Walkotten, W.J., An improved technique for freeze sampling streambed sediments. Vol. 281. 1976: US Dept. of Agriculture, Forest Service, Pacific Northwest Forest and Range Experiment Station. 77. Carling, P. and N. Reader, A freeze-sampling technique suitable for coarse river bed-material. Sedimentary Geology, 1981. 29(2): p. 233-239. 78. Petts, G.E., et al., A freeze-coring technique applied to pollution by fine sediments in gravel-bed rivers. Science of the Total Environment, 1989. 84: p. 259-272. 79. Moser, D.P., et al., Biogeochemical processes and microbial characteristics across groundwater-surface water boundaries of the Hanford Reach of the Columbia River. Environmental science & technology, 2003. 37(22): p. 5127-5134. 129  80. Toran, L., et al., Freeze Core Sampling to Validate Time‐Lapse Resistivity Monitoring of the Hyporheic Zone. Groundwater, 2013. 51(4): p. 635-640. 81. Miskimmin, B.M., et al., A new hammer-driven freeze corer. Journal of Paleolimnology, 1996. 15(3): p. 265-269. 82. Stegen, J.C., et al., Groundwater-surface water mixing shifts ecological assembly processes and stimulates organic carbon turnover. Nature communications, 2016. 7. 83. Zarnetske, J.P., et al., Dynamics of nitrate production and removal as a function of residence time in the hyporheic zone. Journal of Geophysical Research: Biogeosciences, 2011. 116(G1). 84. Parkin, T.B., Soil microsites as a source of denitrification variability. Soil Science Society of America Journal, 1987. 51(5): p. 1194-1199. 85. Grishchenkov, V., et al., Degradation of petroleum hydrocarbons by facultative anaerobic bacteria under aerobic and anaerobic conditions. Process Biochemistry, 2000. 35(9): p. 889-896. 86. Sawyer, A., Enhanced removal of groundwater‐borne nitrate in heterogeneous aquatic sediments. Geophysical Research Letters, 2015. 42(2): p. 403-410. 87. Arnon, S., K.A. Gray, and A.I. Packman, Biophysicochemical process coupling controls nitrogen use by benthic biofilms. Limnology and oceanography, 2007. 52(4): p. 1665-1671. 88. Holmes, R.M., S.G. Fisher, and N.B. Grimm, Parafluvial nitrogen dynamics in a desert stream ecosystem. Journal of the North American Benthological Society, 1994: p. 468-478. 89. Triska, F.J., J.H. Duff, and R.J. Avanzino, Influence of exchange flow between the channel and hyporheic zone on nitrate production in a small mountain stream. Canadian journal of fisheries and aquatic sciences, 1990. 47(11): p. 2099-2111. 90. Canfield, D.E., B. Thamdrup, and J.W. Hansen, The anaerobic degradation of organic matter in Danish coastal sediments: iron reduction, manganese reduction, and sulfate reduction. Geochimica et Cosmochimica Acta, 1993. 57(16): p. 3867-3883. 91. Aller, R.C., Bioturbation and remineralization of sedimentary organic matter: effects of redox oscillation. Chemical Geology, 1994. 114(3): p. 331-345. 92. Bieber, C., Field sampling and modelling of creosote-derived contamination in a tidally forced aquifer. 2003. 93. Hackett, G., J.H. Lehr, and N.W.W. Association, Iron bacteria occurrence, problems and control methods in water wells, in Iron bacteria occurrence, problems and control methods in water wells1985, National Water Well Association. 94. Coby, A.J., et al., Repeated anaerobic microbial redox cycling of iron. Applied and Environmental Microbiology, 2011. 77(17): p. 6036-6042. 95. Straub, K.L., et al., Diversity of ferrous iron-oxidizing, nitrate-reducing bacteria and their involvement in oxygen-independent iron cycling. Geomicrobiology Journal, 2004. 21(6): p. 371-378. 130  96. Gault, A.G., et al., Microbial and geochemical features suggest iron redox cycling within bacteriogenic iron oxide-rich sediments. Chemical Geology, 2011. 281(1): p. 41-51. 97. Bourg, A.C. and C. Bertin, Biogeochemical processes during the infiltration of river water into an alluvial aquifer. Environmental science & technology, 1993. 27(4): p. 661-666. 98. Wielinga, B., et al., Geomicrobial profile through the hyporheic zone of a historic mining flood plain, 1994, American Water Resources Association, Herndon, VA (United States). 99. Wielinga, B., et al., Microbiological and geochemical characterization of fluvially deposited sulfidic mine tailings. Applied and Environmental Microbiology, 1999. 65(4): p. 1548-1555. 100. Chapelle, F.H. and D.R. Lovley, Competitive exclusion of sulfate reduction by Fe (lll)‐reducing bacteria: a mechanism for producing discrete zones of high‐iron ground water. Ground Water, 1992. 30(1): p. 29-36. 101. Hendricks, S.P., Microbial ecology of the hyporheic zone: a perspective integrating hydrology and biology. Journal of the North American Benthological Society, 1993: p. 70-78. 102. Ruby, E.G., H.W. Jannasch, and W.G. Deuser, Fractionation of stable carbon isotopes during chemoautotrophic growth of sulfur-oxidizing bacteria. Applied and Environmental Microbiology, 1987. 53(8): p. 1940-1943. 103. Holmer, M. and E. Kristensen, Coexistence of sulfate reduction and methane production in an organic-rich sediment. Marine Ecology-Progress Series, 1994. 107: p. 177-177. 104. Oremland, R.S. and S. Polcin, Methanogenesis and sulfate reduction: competitive and noncompetitive substrates in estuarine sediments. Applied and Environmental Microbiology, 1982. 44(6): p. 1270-1276. 105. Jones Jr, J.B., et al., Chemoautotrophic production and respiration in the hyporheic zone of a Sonoran Desert stream, 1994, American Water Resources Association, Herndon, VA (United States). 106. Jones Jr, J.B., et al., Methanogenesis in Arizona, USA dryland streams. Biogeochemistry, 1995. 31(3): p. 155-173. 107. Dahm, C., D. Carr, and R. Coleman, Anaerobic carbon cycling in stream ecosystems. Internationale Vereinigung fuer Theoretische und Angewandte Limnologie. Verhandlungen IVTLAP, 1991. 24(3). 108. Trimmer, M., et al., Potential carbon fixation via methane oxidation in well-oxygenated riverbed gravels. Limnology and oceanography, 2010. 55(2): p. 560. 109. Kohzu, A., et al., Stream food web fueled by methane-derived carbon. Aquatic Microbial Ecology, 2004. 36(2): p. 189-194. 110. USGS, Standard Operating Procedure for Sectioning of Sediment Cores, 2007. 111. Hall, G., et al., Selective leaches revisited, with emphasis on the amorphous Fe oxyhydroxide phase extraction. Journal of Geochemical Exploration, 1996. 56(1): p. 59-78. 131  112. Heron, G., et al., Speciation of Fe (II) and Fe (III) in contaminated aquifer sediments using chemical extraction techniques. Environmental science & technology, 1994. 28(9): p. 1698-1705. 113. Lovley, D.R. and E.J. Phillips, Availability of ferric iron for microbial reduction in bottom sediments of the freshwater tidal Potomac River. Applied and Environmental Microbiology, 1986. 52(4): p. 751-757. 114. Bartram, A.K., et al., Generation of multimillion-sequence 16S rRNA gene libraries from complex microbial communities by assembling paired-end Illumina reads. Applied and Environmental Microbiology, 2011. 77(11): p. 3846-3852. 115. Schloss, P.D., et al., Introducing mothur: open-source, platform-independent, community-supported software for describing and comparing microbial communities. Applied and Environmental Microbiology, 2009. 75(23): p. 7537-7541. 116. Edgar, R.C., et al., UCHIME improves sensitivity and speed of chimera detection. Bioinformatics, 2011. 27(16): p. 2194-2200. 117. Kreimer, A., et al., The evolution of modularity in bacterial metabolic networks. Proceedings of the National Academy of Sciences, 2008. 105(19): p. 6976-6981. 118. Cox, T.L. and L. Sly, Phylogenetic relationships and uncertain taxonomy of Pedomicrobium species. International Journal of Systematic and Evolutionary Microbiology, 1997. 47(2): p. 377-380. 119. Timmermans, P. and A. Van Haute, Denitrification with methanol: fundamental study of the growth and denitrification capacity of Hyphomicrobium sp. Water Research, 1983. 17(10): p. 1249-1255. 120. Sohn, J.H., et al., Novosphingobium pentaromativorans sp. nov., a high-molecular-mass polycyclic aromatic hydrocarbon-degrading bacterium isolated from estuarine sediment. International Journal of Systematic and Evolutionary Microbiology, 2004. 54(5): p. 1483-1487. 121. Nakatsu, C.H., et al., Methylibium petroleiphilum gen. nov., sp. nov., a novel methyl tert-butyl ether-degrading methylotroph of the Betaproteobacteria. International Journal of Systematic and Evolutionary Microbiology, 2006. 56(5): p. 983-989. 122. Ghiorse, W., Biology of iron-and manganese-depositing bacteria. Annual reviews in microbiology, 1984. 38(1): p. 515-550. 123. Stoecker, K., et al., Cohn's Crenothrix is a filamentous methane oxidizer with an unusual methane monooxygenase. Proceedings of the National Academy of Sciences of the United States of America, 2006. 103(7): p. 2363-2367. 124. Bowman, J., The methanotrophs—the families Methylococcaceae and Methylocystaceae, in The prokaryotes2006, Springer. p. 266-289. 125. Hallberg, K.B., S. Hedrich, and D.B. Johnson, Acidiferrobacter thiooxydans, gen. nov. sp. nov.; an acidophilic, thermo-tolerant, facultatively anaerobic iron-and sulfur-oxidizer of the family Ectothiorhodospiraceae. Extremophiles, 2011. 15(2): p. 271-279. 132  126. Vreeland, R., et al., Halomonas elongata, a new genus and species of extremely salt-tolerant bacteria. International Journal of Systematic and Evolutionary Microbiology, 1980. 30(2): p. 485-495. 127. Kuever, J., F.A. Rainey, and F. Widdel, Class IV. Deltaproteobacteria class nov2005: Springer. 128. Garrity, G., et al., Bergey's Manual® of Systematic Bacteriology: Volume Two: The Proteobacteria2006: Springer Science & Business Media. 129. Elferink, S.J.O., et al., Desulfobacca acetoxidans gen. nov., sp. nov., a novel acetate-degrading sulfate reducer isolated from sulfidogenic granular sludge. International Journal of Systematic and Evolutionary Microbiology, 1999. 49(2): p. 345-350. 130. Jackson, B.E., et al., Syntrophus aciditrophicus sp. nov., a new anaerobic bacterium that degrades fatty acids and benzoate in syntrophic association with hydrogen-using microorganisms. Archives of microbiology, 1999. 171(2): p. 107-114. 131. Slobodkina, G.B., et al., Deferrisoma camini gen. nov., sp. nov., a moderately thermophilic, dissimilatory iron (III)-reducing bacterium from a deep-sea hydrothermal vent that forms a distinct phylogenetic branch in the Deltaproteobacteria. International Journal of Systematic and Evolutionary Microbiology, 2012. 62(10): p. 2463-2468. 132. Treude, N., et al., Strain FAc12, a dissimilatory iron-reducing member of the Anaeromyxobacter subgroup of Myxococcales. FEMS microbiology ecology, 2003. 44(2): p. 261-269. 133. Kodama, Y. and K. Watanabe, Sulfuricurvum kujiense gen. nov., sp. nov., a facultatively anaerobic, chemolithoautotrophic, sulfur-oxidizing bacterium isolated from an underground crude-oil storage cavity. International Journal of Systematic and Evolutionary Microbiology, 2004. 54(6): p. 2297-2300. 134. Foesel, B.U., M. Rohde, and J. Overmann, Blastocatella fastidiosa gen. nov., sp. nov., isolated from semiarid savanna soil–The first described species of Acidobacteria subdivision 4. Systematic and applied microbiology, 2013. 36(2): p. 82-89. 135. Losey, N.A., et al., Thermoanaerobaculum aquaticum gen. nov., sp. nov., the first cultivated member of Acidobacteria subdivision 23, isolated from a hot spring. International Journal of Systematic and Evolutionary Microbiology, 2013. 63(11): p. 4149-4157. 136. Ehrich, S., et al., A new obligately chemolithoautotrophic, nitrite-oxidizing bacterium, Nitrospira moscoviensis sp. nov. and its phylogenetic relationship. Archives of microbiology, 1995. 164(1): p. 16-23. 137. Lee, H.-W., et al., Blastopirellula cremea sp. nov., isolated from a dead ark clam. International Journal of Systematic and Evolutionary Microbiology, 2013. 63(6): p. 2314-2319. 138. Glöckner, F.O., et al., Complete genome sequence of the marine planctomycete Pirellula sp. strain 1. Proceedings of the National Academy of Sciences, 2003. 100(14): p. 8298-8303. 139. Bauld, J. and J.T. STALEY, Planctomyces maris sp. nov.: a marine isolate of the Planctomyces-Blastocaulis group of budding bacteria. Microbiology, 1976. 97(1): p. 45-55. 133  140. Sangwan, P., et al., Chthoniobacter flavus gen. nov., sp. nov., the first pure-culture representative of subdivision two, Spartobacteria classis nov., of the phylum Verrucomicrobia. Applied and Environmental Microbiology, 2004. 70(10): p. 5875-5881. 141. Haroon, M.F., et al., Anaerobic oxidation of methane coupled to nitrate reduction in a novel archaeal lineage. Nature, 2013. 500(7464): p. 567-570. 142. Konhauser, K.O., Diversity of bacterial iron mineralization. Earth-Science Reviews, 1998. 43(3): p. 91-121. 143. Emerson, D. and C. Moyer, Isolation and characterization of novel iron-oxidizing bacteria that grow at circumneutral pH. Applied and Environmental Microbiology, 1997. 63(12): p. 4784-4792. 144. Wang, Z., et al., Characteristics of aerobic and anaerobic ammonium-oxidizing bacteria in the hyporheic zone of a contaminated river. World Journal of Microbiology and Biotechnology, 2012. 28(9): p. 2801-2811. 145. Neilson-Welch, L.A., Saline water intrusion from the Fraser River estuary: A hydrogeological investigation using field chemical data and a density-dependent groundwater flow model, 1999, University of British Columbia.   Date Sampled 07/24/2014 07/24/2014 08/25/2014 08/25/2014 08/25/2014 08/26/2014 08/26/2014 08/26/2014 08/26/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014Time Sampled (time in + 30 min) 12:00 15:10 11:45 13:45 14:50 12:00 13:30 15:00 16:15 9:40 10:20 12:15 13:10 13:45 14:25 15:00Water Level (m, New West Encoder 1) 2.761 1.902 0.653 0.705 1.176 0.747 0.603 1.064 1.764 1.84 1.582 0.931 0.732 0.695 0.782 0.968Tide (rising/falling/H/L) ↗ ↘ ↘ ↘ ↘ ↗ ↗ ↘ L ↗ ↗ ↘ ↘ ↘ ↘ L ↗ ↗Electrical Conductivity (uS/cm)A04 A05 B01 B02 B03 B04 B05 B06 B07 B08 B09 B10 B11 B12 B13 B14941 2316 3000 1990 1714 952 404 672 1764 5910 5730 3970 1674 200 202 504pHA04 A05 B01 B02 B03 B04 B05 B06 B07 B08 B09 B10 B11 B12 B13 B146.86 6.68 7.2 6.67 6.77 6.77 6.77 6.79 6.58 7.26 7.09 6.5 6.58 7.17 7.14 6.7920°‐Distance (m) to HWMomitted (silt): B01, B02, B07, B10, B11, C07, C11, C12 68 73 81 75 73 75 99 105 79 83 85Anion ScanSample Description A04-FU A05-FU B01-FU B02-FU B03-FU B04-FU B05-FU B06-FU B07-FU B08-FU B09-FU B10-FU B11-FU B12-FU B13-FU B14-FUDate Sampled 07/24/2014 07/24/2014 08/25/2014 08/25/2014 08/25/2014 08/26/2014 08/26/2014 08/26/2014 08/26/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014Parameter Unit G / S RDL 5791378 5791379 5791380 5791381 5791382 5791383 5791384 5791385 5791386 5791387 5791388 5791389 5791390 5791391 5791392 5791393Chloride mg/L 1500 0.05 143 479 713 106 191 116 52.9 53.7 150 1750 1670 936 315 48.0 10.9 76.8Nitrate-N mg/L 400 0.005 0.244 <0.005 <0.005 0.106 <0.005 <0.005 0.034 <0.005 <0.005 <0.005 <0.005 0.036 <0.005 <0.005 0.013 0.010Nitrite-N mg/L 0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005Sulphate mg/L 1000 0.5 <0.5 <0.5 1.5 <0.5 <0.5 <0.5 5.8 <0.5 <0.5 87.2 59.8 <0.5 <0.5 6.7 6.6 <0.5Fluoride mg/L 0.02 0.04 0.04 0.08 0.11 0.06 0.09 0.04 0.08 0.03 0.11 0.11 0.04 0.02 0.09 0.12 0.13Bromide mg/L 0.05 0.13 0.73 1.48 0.11 0.26 0.41 0.23 0.35 0.16 2.81 2.68 1.49 0.55 0.26 0.06 0.48Dissolved Metals Low LevelSample Description A04-FA A05-FA B01-FA B02-FA B03-FA B04-FA B05-FA B06-FA B07-FA B08-FA B09-FA B10-FA B11-FA B12-FA B13-FA B14-FADate Sampled 07/24/2014 07/24/2014 08/25/2014 08/25/2014 08/25/2014 08/26/2014 08/26/2014 08/26/2014 08/26/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014 08/27/2014Parameter Unit G / S RDL 5791394 5791395 5791396 5791397 5791398 5791399 5791400 5791401 5791402 5791403 5791404 5791405 5791406 5791407 5791408 5791409Aluminum Dissolved µg/L 0.5 13.6 4.2 4.1 6.3 4.1 3.9 21.6 4.3 2.6 2.4 5.4 5.1 3.2 8.8 10.3 7.3Antimony Dissolved µg/L 200 0.01 0.02 0.03 0.03 0.05 0.02 0.01 0.04 0.02 0.01 0.03 0.03 0.02 0.01 0.01 0.02 <0.01Arsenic Dissolved µg/L 50 0.05 40.4 27.6 26.2 94.3 79.1 55.5 9.41 34.1 51.4 1.75 3.71 54.7 35.8 5.43 10.5 13.2Barium Dissolved µg/L 10000 0.05 147 526 472 518 316 233 51.5 204 316 505 539 558 319 34.5 39.1 119Beryllium Dissolved µg/L 53 0.005 0.007 0.010 0.012 0.012 0.009 0.006 <0.005 0.010 0.008 <0.005 <0.005 0.011 0.009 <0.005 0.008 0.012Bismuth Dissolved µg/L 0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 0.01 <0.01 0.03 <0.01 <0.01 <0.01 0.01Boron Dissolved µg/L 50000 1 35 252 336 38 239 84 30 50 112 176 118 338 177 45 55 83Cadmium Dissolved µg/L 0.005 <0.005 <0.005 0.007 0.007 0.005 0.008 0.011 <0.005 <0.005 0.008 <0.005 0.013 0.005 <0.005 0.006 <0.005Calcium Dissolved µg/L 50 140000 90900 31200 209000 49700 53400 22100 47900 122000 242000 209000 69700 79300 11200 9640 26900Chromium Dissolved µg/L 0.1 0.5 0.7 0.5 0.9 0.6 0.5 0.1 0.3 0.3 0.2 0.2 0.8 0.5 0.2 0.2 0.4Cobalt Dissolved µg/L 40 0.005 0.260 0.461 0.217 1.93 0.292 0.441 0.149 0.217 0.293 0.109 1.76 0.555 0.356 0.018 0.039 0.094Copper Dissolved µg/L 0.05 0.37 0.18 0.39 0.18 0.14 0.18 0.73 0.23 0.07 0.47 0.15 1.26 0.12 0.33 0.26 0.21Iron Dissolved µg/L 10 105000 97500 10900 126000 42900 35900 7610 24600 76400 3230 5780 102000 44300 4040 7520 29800Lead Dissolved µg/L 0.01 0.15 0.03 0.04 0.05 0.04 0.05 0.06 0.05 0.04 0.04 0.03 0.33 0.04 0.06 0.06 0.06Lithium Dissolved µg/L 0.1 4.7 9.0 3.5 4.8 6.6 4.8 2.1 3.9 7.6 6.0 5.1 12.0 9.3 2.2 2.5 4.1Magnesium Dissolved µg/L 50 33300 35700 30700 56400 18200 22700 9950 13100 48900 112000 97500 42400 34900 3290 3500 6770Manganese Dissolved µg/L 1 9810 5370 1830 11700 3100 3550 479 4060 12400 11200 10400 2620 3820 576 816 2020Mercury Dissolved µg/L 1 0.003 0.023 0.008 0.005 0.026 0.005 0.005 0.004 0.005 0.006 0.005 0.005 0.007 <0.003 <0.003 0.026 <0.003Molybdenum Dissolved µg/L 10000 0.01 0.36 0.14 1.18 2.15 1.08 1.31 0.87 1.78 0.33 0.85 0.93 0.97 1.46 1.35 1.03 0.29Nickel Dissolved µg/L 0.05 0.42 0.38 0.54 4.89 0.29 11.8 0.65 0.38 0.33 0.91 2.64 0.65 0.28 0.16 0.13 0.16Potassium Dissolved µg/L 50 6800 14300 15500 9900 14300 8200 2800 6900 13900 18200 17100 18500 15300 3700 3200 4800Selenium Dissolved µg/L 10 0.1 <0.1 0.2 0.2 0.2 0.1 <0.1 <0.1 <0.1 0.1 0.2 0.2 0.2 <0.1 <0.1 <0.1 <0.1Silicon Dissolved µg/L 50 13300 18200 8600 18000 14500 14000 4200 12600 14700 6400 6400 18300 18200 9100 10700 13700Silver Dissolved µg/L 0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005Sodium Dissolved µg/L 50 61900 285000 546000 84700 256000 79000 35100 46400 138000 730000 750000 617000 169000 21200 21400 43000Strontium Dissolved µg/L 0.05 748 698 342 1400 440 411 174 368 912 2070 1670 817 670 95.2 84.7 192Thallium Dissolved µg/L 3 0.002 <0.002 <0.002 <0.002 <0.002 <0.002 <0.002 <0.002 <0.002 <0.002 <0.002 <0.002 <0.002 <0.002 0.003 0.004 <0.002Tin Dissolved µg/L 0.01 0.04 0.02 0.04 0.02 0.01 0.01 0.02 0.04 0.02 0.07 0.05 0.15 0.02 0.05 0.02 0.04Titanium Dissolved µg/L 1000 0.1 5.4 7.9 10.8 6.7 8.3 5.5 1.9 5.1 7.4 3.8 4.5 11.5 7.3 3.9 3.7 5.5Uranium Dissolved µg/L 3000 0.005 0.009 0.008 0.037 0.061 0.005 <0.005 0.132 <0.005 <0.005 1.04 1.31 0.006 <0.005 <0.005 <0.005 0.006Vanadium Dissolved µg/L 0.1 1.0 1.0 3.0 2.7 1.7 0.7 0.4 1.3 1.1 0.3 0.5 1.6 1.2 0.7 0.9 1.2Zinc Dissolved µg/L 0.5 2.6 1.1 1.1 2.8 10.3 1.8 1.6 2.2 1.0 1.3 1.5 6.3 1.3 1.4 <0.5 1.4Hardness (calc) ug CaCO3/L 100 487000 374000 204000 754000 199000 227000 96200 174000 506000 1070000 923000 349000 342000 41500 38500 95100Comments: RDL - Reported Detection Limit;     G / S - Guideline / StandardAlkalinityA04-FU A05-FU B01-FU B02-FU B03-FU B05-FU B06-FU B07-FU B08-FU B09-FU B10-FU B11-FU B12-FU B13-FU B14-FUmeq/L 0.01998 11.76 11.2 7.52 17.05 11.2 1.92 4.88 15.2 5.44 5.36 11.6 7.28 1.2 1.28 2mg/L CaCO3 1 588.6 560.6 376.4 853.4 560.6 96.1 244.2 760.8 272.3 268.3 580.6 364.4 60.1 64.1 100.1mg/L HCO3- 1.2192148 717.6 683.4 458.9 1040.4 683.4 117.2 297.8 927.5 332.0 327.1 707.9 444.2 73.2 78.1 122.0Appendix A134Date Sampled 10/15/2014 10/15/2014 10/15/2014 10/16/2014 10/16/2014 10/16/2014 10/17/2014 10/17/2014 10/17/2014 10/17/2014 10/17/2014 12/18/2014 12/18/2014 12/18/2014 12/18/2014Time Sampled (time in + 30 min) 15:30 16:30 17:50 13:00 14:25 15:30 12:30 13:20 14:10 14:50 15:45 11:20 11:55 12:50 13:45Water Level (m, New West Encoder 1) 2.629 2.327 2.083 2.845 2.876 2.728 2.591 2.824 2.972 2.991 2.926 2.523 2.653 2.84 2.991Tide (rising/falling/H/L) ↗ ↘ ↘ ↘ ↘ ↗ ↘ ↘ ↗ ↗ ↗ H ↘ ↗ ↗ ↗ ↗Electrical Conductivity (uS/cm)C01 C02 C03 C04 C05 C06 C08 C09 C10 C11 C12 D01 D02 D03 D041107 1783 1564 910 148.4 5670 2150 2920 1060 2120 1340 480 1879 2530 7060pHC01 C02 C03 C04 C05 C06 C08 C09 C10 C11 C12 D01 D02 D03 D047.22 7.37 6.93 6.6 6.92 6.97 6.83 6.83 6.83 6.83 6.59 7.23 7.12 6.9 7.6220°‐Distance (m) to HWMomitted (silt): B01, B02, B07, B10, B11, C07, C11, C12 78 91 96 85 77 100 101 93 85 80 83 91 100Anion ScanSample Description C01-FU C02-FU C03-FU C04-FU C05-FU C06-FU C08-FU C09-FU C10-FU C11-FU C12-FU D01-FU D02-FU D03-FU D04-FUDate Sampled 10/15/2014 10/15/2014 10/15/2014 10/16/2014 10/16/2014 10/16/2014 10/17/2014 10/17/2014 10/17/2014 10/17/2014 10/17/2014 12/18/2014 12/18/2014 12/18/2014 12/18/2014Parameter Unit G / S RDL 5988761 5988768 5988770 5988772 5988774 5988775 5988777 5988778 5988779 5988780 5988781 6211756 6211758 6211759 6211760Chloride mg/L 1500 0.05 131 420 341 120 0.85 1630 549 795 198 497 188 83.4 372 636 2040Nitrate-N mg/L 400 0.005 <0.005 <0.005 <0.005 <0.005 0.016 <0.005 0.011 <0.005 0.006 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005Nitrite-N mg/L 0.005 <0.005 <0.005 <0.005 <0.005 0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005Sulphate mg/L 1000 0.5 <0.5 12.2 <0.5 <0.5 8.9 14 61.9 115 <0.5 13.9 <0.5 5.6 <0.5 88.4 <0.5Fluoride mg/L 0.02 0.12 <0.02 <0.02 0.04 0.04 0.19 0.04 0.07 0.06 0.02 0.08 0.07 <0.02 0.04 <0.02Bromide mg/L 0.05 0.46 1.39 1.12 0.46 <0.05 0.15 1.85 2.54 0.74 1.76 0.79 0.28 0.83 2.18 2.14Dissolved Metals Low LevelSample Description C01-FA C02-FA C03-FA C04-FA C05-FA C-06-FA C08-FA C09-FA C10-FA C11-FA C12-FA D01-FA D02-FA D03-FA D04-FADate Sampled 10/15/2014 10/15/2014 10/15/2014 10/16/2014 10/16/2014 10/16/2014 10/17/2014 10/17/2014 10/17/2014 10/17/2014 10/17/2014 12/18/2014 12/18/2014 12/18/2014 12/18/2014Parameter Unit G / S RDL 5988795 5988801 5988803 5988804 5988805 5988806 5988808 5988810 5988811 5988812 5988813 6211736 6211744 6211745 6211746Aluminum Dissolved µg/L 0.5 3.1 2.3 7.5 7.1 15.1 2.9 5.6 4.0 5.5 6.8 9.4 12.4 4.2 3.7 1.4Antimony Dissolved µg/L 200 0.01 0.02 0.02 0.03 0.03 0.04 0.03 0.02 0.02 0.02 0.02 0.03 0.38 0.55 0.15 0.12Arsenic Dissolved µg/L 50 0.05 56.0 4.54 11.7 44.3 24.3 3.22 4.25 12.3 20.7 44.4 60.3 14.7 41.0 4.34 1.18Barium Dissolved µg/L 10000 0.05 175 177 157 250 26.1 24.0 159 237 135 167 311 48.9 161 218 392Beryllium Dissolved µg/L 53 0.005 0.007 0.006 0.016 0.009 0.006 0.005 0.005 0.007 0.006 0.006 0.011 0.006 0.007 <0.005 <0.005Bismuth Dissolved µg/L 0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01Boron Dissolved µg/L 50000 1 145 107 77 108 27 157 85 188 143 352 376 71 135 84 256Cadmium Dissolved µg/L 0.005 <0.005 0.007 0.006 0.013 0.011 0.027 0.089 0.023 0.010 0.008 0.006 <0.005 <0.005 <0.005 <0.005Calcium Dissolved µg/L 50 37300 54400 50300 46700 6380 192000 53900 39600 35900 30100 81600 16300 59800 63400 262000Chromium Dissolved µg/L 0.1 0.3 0.1 0.4 0.4 0.2 0.1 0.1 0.2 0.3 0.3 0.7 0.2 0.2 0.1 <0.1Cobalt Dissolved µg/L 40 0.005 0.158 0.067 0.154 0.251 0.066 0.064 0.131 0.078 0.104 0.254 0.400 0.047 0.199 0.048 0.102Copper Dissolved µg/L 0.05 0.25 0.06 0.08 0.66 1.15 0.11 0.62 0.26 0.34 0.20 0.07 0.22 0.11 0.09 0.08Iron Dissolved µg/L 10 19700 20600 24300 81600 6900 42500 15600 37500 16600 17600 104000 11600 38600 59000 9010Lead Dissolved µg/L 0.01 0.03 0.02 0.02 0.08 0.15 0.02 0.09 0.05 0.04 0.04 0.02 0.05 0.02 <0.01 <0.01Lithium Dissolved µg/L 0.1 5.1 5.8 6.4 8.2 2.0 10.2 4.5 7.5 5.7 6.3 5.9 3.2 7.4 7.2 7.4Magnesium Dissolved µg/L 50 45500 31500 27000 11000 2520 81000 31500 20200 18500 14300 23800 7630 57500 31800 123000Manganese Dissolved µg/L 1 1290 6080 6770 2350 777 15100 4080 2150 3390 2720 5250 1130 2700 6960 18600Mercury Dissolved µg/L 1 0.003Molybdenum Dissolved µg/L 10000 0.01 1.25 0.24 0.30 0.27 1.23 0.33 0.80 1.41 0.44 0.27 0.15 1.10 0.95 0.85 0.43Nickel Dissolved µg/L 0.05 0.24 0.20 0.71 1.00 0.73 0.40 0.57 0.46 0.53 1.01 0.76 0.49 1.20 0.37 0.48Potassium Dissolved µg/L 50 9400 8900 11200 8000 2000 19500 9200 13400 9000 12600 10700 4010 9960 10600 19500Selenium Dissolved µg/L 10 0.1 <0.1 <0.1 0.1 <0.1 <0.1 0.1 <0.1 <0.1 <0.1 <0.1 0.1 0.4 0.4 <0.1 1.0Silicon Dissolved µg/L 50 16100 11000 12100 20100 6800 8800 8400 11100 12400 14400 20500 11700 14500 10300 9790Silver Dissolved µg/L 0.005 <0.005 <0.005 <0.005 <0.005 <0.005 0.010 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005Sodium Dissolved µg/L 50 104000 201000 187000 57700 14400 768000 267000 462000 120000 353000 122000 65000 200000 344000 920000Strontium Dissolved µg/L 0.05 346 429 378 347 35.9 1790 463 422 280 251 433 157 605 661 3040Thallium Dissolved µg/L 3 0.002 0.004 <0.002 <0.002 0.002 <0.002 <0.002 <0.002 0.002 <0.002 <0.002 <0.002 0.005 0.003 0.003 <0.002Tin Dissolved µg/L 0.01 0.02 <0.01 0.02 0.01 0.03 0.02 0.02 0.01 0.01 0.01 0.01 <0.01 <0.01 <0.01 <0.01Titanium Dissolved µg/L 1000 0.1 5.4 3.3 5.6 5.6 3.1 4.4 2.9 3.3 3.9 5.3 10.1 4.0 4.8 3.2 6.6Uranium Dissolved µg/L 3000 0.005 <0.005 0.005 0.020 0.014 0.006 0.008 0.005 <0.005 0.006 0.009 0.007 <0.005 0.010 3.2 0.038Vanadium Dissolved µg/L 0.1 1.1 0.5 2.8 1.1 0.7 0.4 0.5 0.5 0.8 1.0 1.0 0.8 0.7 0.3 0.4Zinc Dissolved µg/L 0.5 2.1 1.2 2.2 4.4 4.2 2.0 4.1 3.1 1.4 1.3 1.2 0.6 0.7 <0.5 <0.5Hardness (calc) ug CaCO3/L 100 281000 266000 237000 162000 26300 813000 264000 182000 166000 134000 302000 72100 386000 289000 1160000Comments: RDL - Reported Detection Limit;     G / S - Guideline / StandardAlkalinityC01-FU C02-FU C03-FU C04-FU C05-FU C06-FU C08-FU C09-FU C10-FU C11-FU C12-FU D01-FU D02-FU D03-FU D04-FUmeq/L 0.01998 7.61 3.63 5.19 3.89 1.12 5.1 1.9 0.95 3.98 4.58 8.65 1.75 7.31 2.2 10.83mg/L CaCO3 1 380.9 181.7 259.8 194.7 56.1 255.3 95.1 47.5 199.2 229.2 432.9 87.6 365.9 110.1 542.0mg/L HCO3- 1.2192148 464.4 221.5 316.7 237.4 68.3 311.2 115.9 58.0 242.9 279.5 527.8 106.8 446.1 134.2 660.9Appendix A135Date Sampled 12/18/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 01/26/2015 01/26/2015 01/26/2015 01/26/2015 01/26/2015 01/26/2015 01/27/2015Time Sampled (time in + 30 min) 14:25 10:20 11:05 11:30 12:10 12:50 13:25 13:30 14:10 11:30 11:55 12:40 13:35 14:45 15:10 10:35Water Level (m, New West Encoder 1) 3.035 2.436 2.508 2.587 2.721 2.856 2.956 3.153 3.065 2.865 2.507 1.945 1.763 3.014Tide (rising/falling/H/L) ↗ ↘ ↗ ↗ ↗ ↗ ↗ ↗ ↗ ↘ ↘ ↘ ↘ ↘ ↘ ↗Electrical Conductivity (uS/cm) River WedgeD05 D06 D07 D08 D09 D10 D11 DR2 DR3 E01 E02 E03 E04 E05 E06 E075540 2040 4020 5340 5940 6290 5280 829 2760 2020 4140 5890 5270 1283pH River WedgeD05 D06 D07 D08 D09 D10 D11 DR2 DR3 E01 E02 E03 E04 E05 E06 E077.16 7.14 6.92 7.21 7.21 6.92 6.77 7.33 7.29 7.46 7.74 7.26 6.96 6.7620°‐Distance (m) to HWM River Wedgeomitted (silt): B01, B02, B07, B10, B11, C07, C11, C12 108 85 95 107 120 122 102 78 89 95 99 122 107 81Anion Scan River WedgeSample Description D05-FU D06-FU D07-FU D08-FU D09-FU D10-FU D11-FU DR2-FU DR3-FU E01-FU E02-FU E03-FU E04-FU E05-FU E06-FU E07-FUDate Sampled 12/18/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 01/26/2015 01/26/2015 01/26/2015 01/26/2015 01/26/2015 01/26/2015 01/27/2015Parameter Unit G / S RDL 6211761 6212282 6212283 6212284 6212285 6212286 6212287 6212300 6212301 6281980 6281981 6281982 6281983 6281984 6281985 6281986Chloride mg/L 1500 0.05 1530 483 1140 1520 1700 1800 1460 1.60 13100 183 687 503 1140 1710 1480 217Nitrate-N mg/L 400 0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 0.134 8.53 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005Nitrite-N mg/L 0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005Sulphate mg/L 1000 0.5 <0.5 56.6 134 43.5 8.5 6.2 <0.5 6.8 1710 5.4 32.3 69.8 68.4 10.9 1.0 <0.5Fluoride mg/L 0.02 <0.02 <0.02 0.03 0.12 0.16 0.07 <0.02 0.02 <0.02 0.06 0.03 <0.02 0.25 0.25 0.09 0.05Bromide mg/L 0.05 1.41 1.65 0.87 1.37 1.70 1.71 1.28 <0.05 43 0.60 2.41 1.72 0.95 1.78 1.49 0.82Dissolved Metals Low Level River WedgeSample Description D05-FA D06-FA D07-FA D08-FA D09-FA D10-FA D11-FA DR2-FU DR3-FU E01-FA E02-FA E03-FA E04-FA E05-FA E06-FA E07-FADate Sampled 12/18/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 12/19/2014 01/26/2015 01/26/2015 01/26/2015 01/26/2015 01/26/2015 01/26/2015 01/27/2015Parameter Unit G / S RDL 6211747 6211749 6211750 6211751 6211752 6211753 6211754 6212305 6212306 6282002 6282023 6282024 6282025 6282026 6282027 6282028Aluminum Dissolved µg/L 0.5 3.2 7.7 2.7 1.5 1.9 5.1 5.9 71.7 17 22 <5 9 <5 6 <5 13Antimony Dissolved µg/L 200 0.01 0.30 0.14 0.12 0.14 0.12 0.12 0.15 1.21 0.3 0.5 0.2 0.2 0.2 0.1 <0.1 0.1Arsenic Dissolved µg/L 50 0.05 0.70 7.92 4.45 0.46 0.12 3.80 4.14 0.32 1.6 13.6 4.5 2.5 0.7 5.1 1.4 49.1Barium Dissolved µg/L 10000 0.05 403 157 332 335 370 769 674 13.6 15.3 64.7 217 158 220 686 536 218Beryllium Dissolved µg/L 53 0.005 0.007 <0.005 <0.005 <0.005 0.006 0.013 0.006 <0.005 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05Bismuth Dissolved µg/L 0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.01 <0.1 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05Boron Dissolved µg/L 50000 1 128 232 198 147 123 107 100 7 2470 47 121 71 147 93 113 295Cadmium Dissolved µg/L 0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 0.010 0.08 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05Calcium Dissolved µg/L 50 241000 34800 99500 193000 238000 298000 235000 13100 237000 28800 90400 64900 126000 251000 197000 26200Chromium Dissolved µg/L 0.1 0.1 0.1 <0.1 <0.1 0.1 0.3 0.3 0.3 <1 <1 <1 <1 <1 <1 <1 <1Cobalt Dissolved µg/L 40 0.005 0.081 0.050 0.078 0.057 0.037 0.068 0.130 0.062 0.07 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05Copper Dissolved µg/L 0.05 0.23 0.17 0.06 <0.05 0.06 0.08 <0.05 1.16 1.6 1.2 <0.5 <0.5 <0.5 <0.5 <0.5 <0.5Iron Dissolved µg/L 10 32400 16200 66100 16400 20900 28700 64200 123 <10 13400 28900 12400 3810 16200 51300 40900Lead Dissolved µg/L 0.01 0.01 0.02 <0.01 <0.01 0.01 0.01 <0.01 0.07 <0.1 0.4 0.7 0.7 0.8 0.5 0.6 0.3Lithium Dissolved µg/L 0.1 6.8 5.1 9.9 8.7 5.0 5.5 5.4 0.6 68 3 6 4 4 4 6 6Magnesium Dissolved µg/L 50 91400 16300 53100 89700 94000 102000 98700 2720 858000 10300 45400 38400 64800 92500 83700 5050Manganese Dissolved µg/L 1 22300 3680 8370 18600 17900 34700 19800 16 7 2490 6790 4990 5130 29900 17700 1750Mercury Dissolved µg/L 1 0.003Molybdenum Dissolved µg/L 10000 0.01 0.15 1.61 0.54 0.32 0.10 0.28 0.10 0.69 8.2 0.9 9.9 0.9 0.6 0.6 1.6 4.4Nickel Dissolved µg/L 0.05 0.55 0.33 0.45 0.30 0.41 0.64 0.70 0.68 6.5 <0.5 <0.5 <0.5 0.7 1.0 0.5 0.8Potassium Dissolved µg/L 50 13800 10500 14000 15300 12200 15300 13300 783 254000 4800 10100 6700 9900 12200 10100 7200Selenium Dissolved µg/L 10 0.1 1.2 0.4 0.4 0.4 0.9 0.8 0.7 <0.1 1 <1 <1 <1 <1 <1 <1 <1Silicon Dissolved µg/L 50 9510 13100 11600 10000 7170 8630 9640 2810 1380 7100 9400 8000 7100 6100 9200 14400Silver Dissolved µg/L 0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 0.009 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05Sodium Dissolved µg/L 50 729000 352000 585000 728000 790000 748000 652000 3430 7050000 106000 279000 198000 537000 755000 666000 184000Strontium Dissolved µg/L 0.05 2040 382 1110 1550 2040 2720 1870 63.0 6860 222 864 580 1060 1560 1440 224Thallium Dissolved µg/L 3 0.002 0.002 <0.002 0.002 <0.002 <0.002 <0.002 <0.002 0.002 0.04 0.12 0.06 0.05 0.02 <0.02 <0.02 0.07Tin Dissolved µg/L 0.01 <0.01 <0.01 0.20 <0.01 <0.01 <0.01 <0.01 <0.01 <0.1 0.17 0.20 <0.05 0.09 0.09 <0.05 <0.05Titanium Dissolved µg/L 1000 0.1 5.6 4.6 3.9 4.4 4.2 5.5 6.0 4.5 2 4 3 3 2 2 3 9Uranium Dissolved µg/L 3000 0.005 0.011 <0.005 <0.005 0.017 0.024 0.295 0.012 0.189 2.31 <0.05 <0.05 <0.05 <0.05 0.70 <0.05 <0.05Vanadium Dissolved µg/L 0.1 0.6 0.8 0.3 0.3 0.5 1.1 0.7 0.4 <1 <1 <1 <1 <1 1 <1 6Zinc Dissolved µg/L 0.5 1.2 0.7 0.8 0.5 0.5 0.8 0.8 1.1 <5 <5 <5 <5 <5 <5 <5 <5Hardness (calc) ug CaCO3/L 100 978000 154000 467000 851000 981000 1160000 993000 43900 4130000 114000 413000 320000 581000 1010000 837000 86200Comments: RDL - Reported Detection Limit;     G / S - Guideline / StandardAlkalinity River WedgeD05-FU D06-FU D07-FU D08-FU D09-FU D10-FU D11-FU DR2-FU DR3-FU E01-FU E02-FU E03-FU E04-FU E05-FU E06-FU E07-FUmeq/L 0.01998 9.02 1.43 1.95 6.62 9.29 11.26 10.55 0.98 0.99 1.76 2.91 1.34 3.20 6.41 6.23 4.45mg/L CaCO3 1 451.5 71.6 97.6 331.3 465.0 563.6 528.0 49.0 49.5 88.1 145.6 67.1 160.2 320.8 311.8 222.7mg/L HCO3- 1.2192148 550.4 87.3 119.0 404.0 566.9 687.1 643.8 59.8 60.4 107.4 177.6 81.8 195.3 391.1 380.2 271.5Appendix A136Date Sampled 01/27/2015 01/27/2015 01/27/2015 01/27/2015 01/27/2015 01/27/2015 01/28/2015 01/28/2015 01/28/2015 01/28/2015 01/28/2015 01/28/2015Time Sampled (time in + 30 min) 11:15 11:45 12:50 13:30 14:10 14:55 10:25 11:25 11:55 12:30 14:55 15:30Water Level (m, New West Encoder 1) 3.101 3.105 2.882 2.675 2.428 2.088 2.552 2.748 2.794 2.798 2.202 1.96Tide (rising/falling/H/L) ↗ ↘ ↗ H ↘ ↘ ↘ ↘ ↗ ↗ ↗ H ↘ ↘Electrical Conductivity (uS/cm)E08 E09 E10 E11 E12 E13 E14 E15 E16 E17 E18 E192370 2470 4990 5770 5110 5760 3930 4970 5100 5090 2410 368pHE08 E09 E10 E11 E12 E13 E14 E15 E16 E17 E18 E196.6 6.880 6.57 6.86 6.65 6.5 6.94 7.05 7.1 7.24 6.91 7.0420°‐Distance (m) to HWMomitted (silt): B01, B02, B07, B10, B11, C07, C11, C12 86 97 104 95 94 99 89 100 106 109 104 84Anion ScanSample Description E08-FU E09-FU E10-FU E11-FU E12-FU E13-FU E14-FU E15-FU E16-FU E17-FU E18-FU E19-FUDate Sampled 01/27/2015 01/27/2015 01/27/2015 01/27/2015 01/27/2015 01/27/2015 01/28/2015 01/28/2015 01/28/2015 01/28/2015 01/28/2015 01/28/2015Parameter Unit G / S RDL 6281987 6281988 6281989 6281990 6281991 6281992 6282054 6281993 6281994 6281995 6281996 6281997Chloride mg/L 1500 0.05 509 587 1370 1710 1420 1630 1100 1440 1510 1480 541 36.1Nitrate-N mg/L 400 0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005Nitrite-N mg/L 0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005 <0.005Sulphate mg/L 1000 0.5 <0.5 16.9 <0.5 2.9 <0.5 <0.5 123 168 178 174 <0.5 0.8Fluoride mg/L 0.02 <0.02 <0.02 <0.02 0.08 0.06 0.07 0.15 0.06 0.08 0.06 <0.02 0.07Bromide mg/L 0.05 0.25 0.69 1.30 2.06 1.51 1.70 0.54 11.6 11.3 11.5 0.75 0.13Dissolved Metals Low LevelSample Description E08-FA E09-FA E10-FA E11-FA E12-FA E13-FA E14-FA E15-FA E16-FA E17-FA E18-FA E19-FADate Sampled 01/27/2015 01/27/2015 01/27/2015 01/27/2015 01/27/2015 01/27/2015 01/28/2015 01/28/2015 01/28/2015 01/28/2015 01/28/2015 01/28/2015Parameter Unit G / S RDL 6282029 6282030 6282031 6282032 6282033 6282034 6282035 6282036 6282037 6282038 6282040 6282042Aluminum Dissolved µg/L 0.5 <5 <5 6 <5 <5 <5 <5 <5 <5 <5 8 22Antimony Dissolved µg/L 200 0.01 <0.1 <0.1 0.3 <0.1 0.1 <0.1 0.1 0.1 <0.1 0.2 0.1 0.3Arsenic Dissolved µg/L 50 0.05 31.3 2.2 2.0 0.6 1.0 4.5 1.4 3.3 0.9 2.5 11.1 14.5Barium Dissolved µg/L 10000 0.05 345 290 662 461 410 463 419 429 505 425 284 84.0Beryllium Dissolved µg/L 53 0.005 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05Bismuth Dissolved µg/L 0.01 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05Boron Dissolved µg/L 50000 1 219 123 78 201 291 183 454 337 231 198 75 53Cadmium Dissolved µg/L 0.005 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05Calcium Dissolved µg/L 50 50300 114000 237000 139000 120000 179000 85900 90900 115000 150000 81000 22300Chromium Dissolved µg/L 0.1 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1Cobalt Dissolved µg/L 40 0.005 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05Copper Dissolved µg/L 0.05 <0.5 <0.5 <0.5 <0.5 <0.5 <0.5 <0.5 <0.5 <0.5 <0.5 <0.5 <0.5Iron Dissolved µg/L 10 95700 53900 40200 50400 73900 138000 25100 36900 42900 22300 43300 16100Lead Dissolved µg/L 0.01 0.5 0.4 0.4 0.4 0.4 0.3 0.4 0.4 0.3 0.3 0.3 <0.1Lithium Dissolved µg/L 0.1 10 6 6 9 8 10 6 8 8 6 3 3Magnesium Dissolved µg/L 50 13600 37800 94500 87400 86900 88100 94100 78000 96000 85100 34200 5000Manganese Dissolved µg/L 1 3280 11600 26200 5760 5750 11300 3910 6170 5890 9810 6770 1670Mercury Dissolved µg/L 1 0.003Molybdenum Dissolved µg/L 10000 0.01 1.2 0.1 0.1 0.2 <0.1 0.3 0.7 1.1 0.7 1.2 0.2 0.6Nickel Dissolved µg/L 0.05 0.6 0.7 0.5 <0.5 <0.5 <0.5 <0.5 0.6 0.5 0.6 0.6 0.7Potassium Dissolved µg/L 50 13400 7600 13200 14700 15700 14100 17800 16400 14800 12100 4800 4100Selenium Dissolved µg/L 10 0.1 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1 <1Silicon Dissolved µg/L 50 18300 7500 7700 12600 14900 13500 8700 10300 10000 7600 7900 12700Silver Dissolved µg/L 0.005 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05Sodium Dissolved µg/L 50 266000 219000 578000 807000 709000 734000 534000 725000 698000 717000 272000 35400Strontium Dissolved µg/L 0.05 569 906 1640 1420 1220 1500 962 1200 1480 1330 553 142Thallium Dissolved µg/L 3 0.002 0.04 0.02 <0.02 <0.02 <0.02 <0.02 <0.02 <0.02 <0.02 <0.02 <0.02 0.08Tin Dissolved µg/L 0.01 <0.05 <0.05 0.08 0.12 <0.05 0.13 0.12 <0.05 <0.05 0.06 0.10 0.05Titanium Dissolved µg/L 1000 0.1 5 2 3 4 5 5 4 3 3 3 3 4Uranium Dissolved µg/L 3000 0.005 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05 <0.05Vanadium Dissolved µg/L 0.1 2 <1 <1 <1 1 <1 <1 <1 <1 <1 <1 1Zinc Dissolved µg/L 0.5 <5 <5 <5 <5 <5 <5 <5 <5 <5 <5 <5 <5Hardness (calc) ug CaCO3/L 100 182000 440000 981000 707000 658000 810000 602000 548000 683000 725000 343000 76300Comments: RDL - Reported Detection Limit;     G / S - Guideline / StandardAlkalinityE08-FU E09-FU E10-FU E11-FU E12-FU E13-FU E14-FU E15-FU E16-FU E17-FU E18-FU E19-FUmeq/L 0.01998 5.41 3.82 6.87 4.99 6.69 7.58 1.89 1.61 1.31 1.83 5.54 1.93mg/L CaCO3 1 270.8 191.2 343.8 249.7 334.8 379.4 94.6 80.6 65.6 91.6 277.3 96.6mg/L HCO3- 1.2192148 330.1 233.1 419.2 304.5 408.2 462.5 115.3 98.2 79.9 111.7 338.1 117.8Appendix A137Date Sampled 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015Time Sampled (time in + 30 min)Water Level (m, New West Encoder 1)Tide (rising/falling/H/L) ↗ ↘Electrical Conductivity (uS/cm)MW14-02‐070MW14-02‐082MW14-02‐100MW14-02‐119MW14-02‐137MW14-02‐155MW14-02‐173MW14-02‐195MW14-06‐059MW14-06‐071MW14-06‐092MW14-06‐114MW14-06‐135MW14-06‐156MW14-06‐178MW14-06‐199482 472 1342 5810 10200 11490 11840 11640 437 517 754 7500 10320 10910 11670 10070pH02-070 02-082 02-100 02-119 02-137 02-155 02-173 02-195 06-059 06-071 06-092 06-114 06-135 06-156 06-178 06-1996.67 6.47 7.29 7.13 7.34 7.51 7.71 7.9 7.16 7.14 6.97 7.29 7.7 7.84 8.01 7.91Depth (m)omitted (silt): B01, B02, B07, B10, B11, C07, C11, C12 7.00 8.2 10 11.9 13.7 15.5 17.3 19.5 5.90 7.10 9.2 11.4 13.5 15.6 17.8 19.9Anion ScanSample Description 02-070-FU 02-082-FU 02-100-FU 02-119-FU 02-137-FU 02-155-FU 02-173-FU 02-195-FU 06-059-FU 06-071-FU 06-092-FU 06-114-FU 06-135-FU 06-156-FU 06-178-FU 06-199-FUDate Sampled 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015Parameter Unit G / S RDL 6802219 6802220 6802221 6802222 6802223 6802224 6802225 6802226 6802248 6802249 6802250 6802251 6802252 6802253 6802254 6802255Chloride mg/L 1500 0.05 21.0 33.7 90.9 1680 3230 3790 4540 4500 12.7 10.2 33.6 2320 3850 3450 3570 3070Nitrate-N mg/L 400 0.005 <0.005 0.031 0.065 <0.05 <0.05 <0.05 <0.05 <0.05 <0.005 <0.005 0.015 <0.05 <0.05 <0.05 <0.05 <0.05Nitrite-N mg/L 0.005 <0.005 <0.005 <0.005 <0.05 <0.05 <0.05 <0.05 <0.05 <0.005 <0.005 <0.005 <0.05 <0.05 <0.05 <0.05 <0.05Sulphate mg/L 1000 0.5 0.7 <0.5 <0.5 <5 6 <5 <5 <5 <0.5 <0.5 <0.5 42 119 <5 <5 5Fluoride mg/L 0.02 0.04 0.05 0.14 <0.2 <0.2 0.4 0.7 1.4 0.03 <0.02 0.07 <0.2 0.6 0.5 0.7 0.3Bromide mg/L 0.05 0.06 0.09 0.57 5.0 8.8 10.4 10.3 9.8 <0.05 <0.05 0.11 6.7 9.0 9.9 10.3 8.8Dissolved Metals Low LevelSample Description 02-070-FA 02-082-FA 02-100-FA 02-119-FA 02-137-FA 02-155-FA 02-173-FA 02-195-FA 06-059-FA 06-071-FA 06-092-FA 06-114-FA 06-135-FA 06-156-FA 06-178-FA 06-199-FADate Sampled 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/28/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015 7/29/2015Parameter Unit G / S RDL UBC-2 UBC-3 UBC-6 UBC-7 UBC-9 UBC-13 UBC-16 UBC-14 UBC-1 UBC-4 UBC-5 UBC-8 UBC-11 UBC-12 UBC-15 UBC-10Aluminum Dissolved µg/L 0.5Antimony Dissolved µg/L 200 0.01Arsenic Dissolved µg/L 50 0.05 0 0 50.1 0 528.2 0 0 442.4 30.9 41.6 64.3 0 375.9 163 0 0Barium Dissolved µg/L 10000 0.05Beryllium Dissolved µg/L 53 0.005Bismuth Dissolved µg/L 0.01Boron Dissolved µg/L 50000 1Cadmium Dissolved µg/L 0.005Calcium Dissolved µg/L 50 27910 27100 107300 432600 328800 215300 124900 75420 37820 52720 69670 303700 221500 146500 68810 60280Chromium Dissolved µg/L 0.1Cobalt Dissolved µg/L 40 0.005Copper Dissolved µg/L 0.05Iron Dissolved µg/L 10 36.37 36.44 15.32 45.05 12.25 7.34 3.437 0.2653 17.58 19.68 47.92 22.26 4.988 3.342 1.181 0Lead Dissolved µg/L 0.01Lithium Dissolved µg/L 0.1 43.6 47.2 39.7 206.8 407.9 384.7 394.9 383.6 40.2 42.3 43.6 415.9 384.7 409.7 408.7 385.4Magnesium Dissolved µg/L 50 10530 9970 30880 131200 128800 157600 135000 150100 4566 6212 18190 81870 140500 158700 156500 131200Manganese Dissolved µg/L 1 556.5 698.3 719.6 2622 3100 2585 1513 585 958.8 1362 1078 3954 2077 1451 519.1 511.2Mercury Dissolved µg/L 1 0.003Molybdenum Dissolved µg/L 10000 0.01Nickel Dissolved µg/L 0.05Potassium Dissolved µg/L 50 5452 1460 2316 5739 23100 34500 37910 38650 40790 5426 2024 26460 35400 39010 35160 32000Selenium Dissolved µg/L 10 0.1Silicon Dissolved µg/L 50Silver Dissolved µg/L 0.005Sodium Dissolved µg/L 50 33070 22190 123000 515200 1455000 2109000 2351000 2496000 39940 25810 30390 1091000 1906000 2201000 2338000 2024000Strontium Dissolved µg/L 0.05Thallium Dissolved µg/L 3 0.002Tin Dissolved µg/L 0.01Titanium Dissolved µg/L 1000 0.1Uranium Dissolved µg/L 3000 0.005Vanadium Dissolved µg/L 0.1 5.4 5.6 1.1 12.4 19 16.9 34.3 29.8 5.3 3.6 1.9 29.6 12.3 13.3 41.7 18.1Zinc Dissolved µg/L 0.5Hardness (calc) ug CaCO3/L 100Comments: RDL - Reported Detection Limit;     G / S - Guideline / StandardAlkalinity02-070-FU 02-082-FU 02-100-FU 02-119-FU 02-137-FU 02-155-FU 02-173-FU 02-195-FU 06-059-FU 06-071-FU 06-092-FU 06-114-FU 06-135-FU 06-156-FU 06-178-FU 06-199-FUmeq/L 0.01998 3.88 3.53 11.02 11.61 9.22 13.14 20.82 21.82 3.83 4.58 6.87 7.43 8.09 15.56 20.50 16.24mg/L CaCO3 1 194.2 176.7 551.6 581.1 461.5 657.7 1042.0 1092.1 191.7 229.2 343.8 371.9 404.9 778.8 1026.0 812.8mg/L HCO3- 1.2192148 236.8 215.4 672.5 708.5 562.6 801.8 1270.5 1331.5 233.7 279.5 419.2 453.4 493.7 949.5 1250.9 991.0Appendix A138ID River 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16Depth 0 4‐10cm 10‐15cm 15‐20cm 20‐25cm 25‐30cm 30‐35cm 35‐40cm 40‐45cm 45‐50cm 50‐55cm 55‐60cm 60‐65cm 65‐70cm 70‐75cm 75‐80cm 80‐85cmMidpoint (cm) 0 7 12.5 17.5 22.5 27.5 32.5 37.5 42.5 47.5 52.5 57.5 62.5 67.5 72.5 77.5 82.5Date 01/12/2014 13/12/2015 13/12/2015 14/12/2015 14/12/2015 14/12/2015 13/01/2016 14/01/2016 14/01/2016 14/01/2016 14/01/2016 15/01/2016 15/01/2016 15/01/2016 18/01/2016 18/01/2016 20/01/2016Time 20:31 23:09 1:03 2:28 10:55 22:32 0:15 1:34 11:33 15:45 0:16 1:35 2:45 20:05 21:38 13:47pH 7.07 7.22 7.33 7.4 7.64 7.76 7.42 7.36 7.64 7.6 7.82 7.82 7.8 7.83 7.87 7.82SC (µS/cm) 133.8 137.7 154.7 223 313 312 436 538 627 752 1164 1483 1620 1608 1908 2240DO (µg/L) 149 138 72 69 31 110 71 80 30 1 55 50 45 30 39 21DO (µM) 4.66 4.31 2.25 2.16 0.97 3.44 2.22 2.50 0.94 0.03 1.72 1.56 1.41 0.94 1.22 0.66Alk. (meq/L) 0.98 0.58 0.9 1.03 1.39 1.88 0.99 1.3 1.34 1.1 1.5 1.96 2.26 2.64 1.86 2.14 2.96CaCO₃ (mg/L) 49.05 29.03 45.05 51.55 69.57 94.09 49.55 65.07 67.07 55.06 75.08 98.10 113.11 132.13 93.09 107.11 148.15HCO₃‐ (mg/L) 59.80 35.39 54.92 62.85 84.82 114.72 60.41 79.33 81.77 67.12 91.53 119.60 137.91 161.10 113.50 130.59 180.62AnionsCl (mg/L) 1.6 6.94 3.88 4.43 7.49 21.3 43.1 77.4 106 147 173 247 401 446 447 545 661NO₃ (mg/L) 0.134 3.54 1.41 1.35 2.35 1.31 0.687 0.804 0.815 1.03 0.764 1.98 2.46 1.95 1.96 2.08 2.1SO₄ (mg/L) 6.8 8 3.9 6.1 6.3 4.8 3.3 2.1 1.3 1.5 6.6 1.6 1.3 1.5 0.6 0.8 0.6Cl (mM) 0.045 0.196 0.109 0.125 0.211 0.601 1.216 2.183 2.990 4.146 4.880 6.967 11.311 12.580 12.608 15.372 18.644NO₃ (mM) 0.002 0.057 0.023 0.022 0.038 0.021 0.011 0.013 0.013 0.017 0.012 0.032 0.040 0.031 0.032 0.034 0.034SO₄ (mM) 0.071 0.083 0.041 0.063 0.066 0.050 0.034 0.022 0.014 0.016 0.069 0.017 0.014 0.016 0.006 0.008 0.006CationsNa (mg/L) 3.43 2.129 1.472 1.628 3.625 8.657 10.58 26.16 35.3 53.72 83.65 161.2 235.6 238.6 253.8 304.9 353.9Ca (mg/L) 13.1 15.29 17.9433 22.0067 30.5 42.8333 32.24 46.3467 48.58 42 40.9767 40.56 56.05 52.9033 48.6733 63.5767 76.7Mg (mg/L) 2.72 3.098 3.5705 4.2545 6.0795 8.731 7.497 10.0005 10.85 12.21 16.99 17.92 23.13 20.985 20.39 26.24 33.3K (mg/L) 0.783 0.7694 0.7408 0.8681 1.187 1.59 1.702 2.562 2.984 4.654 4.701 5.706 6.649 6.022 5.161 7.243 7.08Fe (mg/L) 0.123 0.1636 0.119 0.0874 0.3177 0.119 0.25455 0.3303 1.7425 2.947 5.611 4.8625 8.0195 12.31 11.565 13.415 24.7Fe(II) (mg/L) 0 0 0 0 0 0.08 0.11 1.6 2.58 2.83 3.9 5.85 10 10 11.75 21.92Mn (mg/L) 0.016 0.10705 0.67385 0.9558 2.4775 3.39 3.144 3.4765 3.4485 2.424 3.5865 3.0445 3.3485 2.865 2.5835 3.3025 3.695Na (mM) 0.149 0.093 0.064 0.071 0.158 0.377 0.460 1.138 1.535 2.337 3.639 7.012 10.248 10.379 11.040 13.262 15.394Ca (mM) 0.327 0.382 0.448 0.549 0.761 1.069 0.804 1.156 1.212 1.048 1.022 1.012 1.399 1.320 1.214 1.586 1.914Mg (mM) 0.112 0.127 0.147 0.175 0.250 0.359 0.308 0.411 0.446 0.502 0.699 0.737 0.952 0.863 0.839 1.080 1.370K (mM) 0.020 0.020 0.019 0.022 0.030 0.041 0.044 0.066 0.076 0.119 0.120 0.146 0.170 0.154 0.132 0.185 0.181Fe (mM) 0.002 0.003 0.002 0.002 0.006 0.002 0.005 0.006 0.031 0.053 0.100 0.087 0.144 0.220 0.207 0.240 0.442Fe(II) (mM) 0.000 0.000 0.000 0.000 0.000 0.000 0.001 0.002 0.029 0.046 0.051 0.070 0.105 0.179 0.179 0.210 0.393Mn (mM) 0.000 0.002 0.012 0.017 0.045 0.062 0.057 0.063 0.063 0.044 0.065 0.055 0.061 0.052 0.047 0.060 0.067Appendix B139ID AV BW CX DY EZRadius 48‐52mm 44‐48mm 39‐44mm 32‐39mm 24‐32mmMidpoint (mm) 50 46 41.5 35.5 28Date 21/01/2016 21/01/2016 21/01/2016 20/01/2016 20/01/2016Time 14:41 13:50 13:01 19:00 17:40pH 7.62 7.24 7.56 7.81 7.73SC (µS/cm) 222 221 223 299 291DO (µg/L) 85 57 55 60 56DO (µM) 2.66 1.78 1.72 1.88 1.75Alk. (meq/L) 1.15 1.31 1.09 1.7 1.6CaCO₃ (mg/L) 57.56 65.57 54.55 85.09 80.08HCO₃‐ (mg/L) 70.18 79.94 66.51 103.74 97.63Cl (mg/L) 18.7 20.1 15.1 15.2 12.5NO₃ (mg/L) 1.93 3.5 1.58 1.21 0.627SO₄ (mg/L) 8.4 8.5 7.3 7.8 7.6Cl (mM) 0.527 0.567 0.426 0.429 0.353NO₃ (mM) 0.031 0.056 0.025 0.020 0.010SO₄ (mM) 0.087 0.088 0.076 0.081 0.079Na (mg/L) 8.397 9.198 5.977 7.485 6.754Ca (mg/L) 39.36 38.18 25.42 33.14 32.14Mg (mg/L) 8.196 7.826 5.478 6.828 6.56K (mg/L) 2.722 1.6966 0.3188 4.238 2.982‐ ‐ ‐ ‐ ‐ ‐‐ ‐ ‐ ‐ ‐ ‐Mn (mg/L) 3.114 2.97 1.9492 2.194 2.898Na (mM) 0.365 0.400 0.260 0.326 0.294Ca (mM) 0.982 0.953 0.634 0.827 0.802Mg (mM) 0.337 0.322 0.225 0.281 0.270K (mM) 0.070 0.043 0.008 0.108 0.076‐ ‐ ‐ ‐ ‐ ‐‐ ‐ ‐ ‐ ‐ ‐Mn (mM) 0.057 0.054 0.035 0.040 0.053Appendix B140       01020304050607080900.01 0.1 1 10Depth (cm)Concentration (mM)NaCaMgK01020304050607080900.001 0.01 0.1 1Depth (cm)Concentration (mM)FeFe(II)MnAppendix C14101020304050607080900.0Depth (cm)01 001020304050607080900Depth (cm).01ConceDi  0.1ntration2DO (µM)ssolved O1 (mM)4₂ 10ClNO₃SO₄6 Appendix C142Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function0 0 7.01 20 9.77237E‐08 1.95447E‐06 0 0 7.22 20 6.0256E‐08 1.20512E‐06100 0.1 4.11 20.1 7.76247E‐05 0.001560257 60 0.06 6.35 20.06 4.46684E‐07 8.96047E‐06112 0.112 3.87 20.112 0.000134896 0.002713034 100 0.1 5.57 20.1 2.69153E‐06 5.40998E‐05120 0.12 3.68 20.12 0.00020893 0.004203664 120 0.12 4.95 20.12 1.12202E‐05 0.00022575140 0.14 3.43 20.14 0.000371535 0.00748272 130 0.13 4.66 20.13 2.18776E‐05 0.000440396180 0.18 3.16 20.18 0.000691831 0.013961149 140 0.14 4.39 20.14 4.0738E‐05 0.000820464240 0.24 2.93 20.24 0.001174898 0.023779927 150 0.15 4.17 20.15 6.76083E‐05 0.001362307340 0.34 2.69 20.34 0.002041738 0.04152895 170 0.17 3.73 20.17 0.000186209 0.00375583440 0.44 2.55 20.44 0.002818383 0.057607747 210 0.21 3.34 20.21 0.000457088 0.009237752640 0.64 2.36 20.64 0.004365158 0.090096868 290 0.29 2.97 20.29 0.001071519 0.021741127390 0.39 2.74 20.39 0.001819701 0.037103701490 0.49 2.59 20.49 0.002570396 0.05266741A B A B0.1659 0.0157 0.1535 0.0227x y x y0.0946353 0 0.1478827 0POI at V = 0.09 mL POI at V = 0.15 mLID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)ID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)Acid #3 20 0.09 0.122 1.15455E‐05 0.000577275 0.58 Acid #3 20 0.15 0.122 1.80417E‐05 0.000902085 0.90Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function0 0 7.32 20 4.7863E‐08 9.5726E‐07 0 0 7.4 20 3.98107E‐08 7.96214E‐0760 0.06 6.42 20.06 3.80189E‐07 7.6266E‐06 80 0.08 6.71 20.08 1.94984E‐07 3.91529E‐06120 0.12 5.6 20.12 2.51189E‐06 5.05392E‐05 120 0.12 6.37 20.12 4.2658E‐07 8.58278E‐06160 0.16 4.59 20.16 2.5704E‐05 0.000518192 150 0.15 6.07 20.15 8.51138E‐07 1.71504E‐05170 0.17 4.33 20.17 4.67735E‐05 0.000943422 180 0.18 5.8 20.18 1.58489E‐06 3.19831E‐05180 0.18 4.1 20.18 7.94328E‐05 0.001602954 210 0.21 5.21 20.21 6.16595E‐06 0.000124614200 0.2 3.75 20.2 0.000177828 0.003592124 220 0.22 4.88 20.22 1.31826E‐05 0.000266552240 0.24 3.39 20.24 0.00040738 0.008245377 230 0.23 4.51 20.23 3.0903E‐05 0.000625167310 0.31 3.08 20.31 0.000831764 0.016893122 250 0.25 3.96 20.25 0.000109648 0.002220368410 0.41 2.87 20.41 0.001348963 0.027532332 290 0.29 3.48 20.29 0.000331131 0.00671865510 0.51 2.72 20.51 0.001905461 0.039080999 370 0.37 3.12 20.37 0.000758578 0.015452225610 0.61 2.6 20.61 0.002511886 0.051769979 470 0.47 2.91 20.47 0.001230269 0.025183602570 0.57 2.75 20.57 0.001778279 0.036579207A B A B0.1162 0.0196 0.1062 0.0242x y x y0.1686747 0 0.2278719 0POI at V = 0.17 mL POI at V = 0.23 mLID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)ID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)Acid #3 20 0.17 0.122 2.05783E‐05 0.001028916 1.03 Acid #3 20 0.23 0.122 2.78004E‐05 0.001390019 1.39Gran function alkalinity calculations Gran function alkalinity calculationsSample 1 Sample 2Gran function alkalinity calculations Gran function alkalinity calculationsSample 3 Sample 4y = 0.1659x ‐ 0.015700.010.020.030.040.050.060.070.080.090.10 0.1 0.2 0.3 0.4 0.5 0.6 0.7Gran functionVolume of acid added (mL)y = 0.1535x ‐ 0.022700.010.020.030.040.050.060 0.1 0.2 0.3 0.4 0.5 0.6Gran functionVolume of acid added (mL)y = 0.1162x ‐ 0.019600.010.020.030.040.050.060 0.1 0.2 0.3 0.4 0.5 0.6 0.7Gran functionVolume of acid added (mL)y = 0.1062x ‐ 0.024200.0050.010.0150.020.0250.030.0350.040 0.1 0.2 0.3 0.4 0.5 0.6Gran functionVolume of acid added (mL)Appendix D143Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function0 0 7.64 20 2.29087E‐08 4.58174E‐07 0 0 7.76 20 1.7378E‐08 3.4756E‐07100 0.1 6.79 20.1 1.62181E‐07 3.25984E‐06 100 0.1 6.09 20.1 8.12831E‐07 1.63379E‐05160 0.16 6.39 20.16 4.0738E‐07 8.21279E‐06 140 0.14 5.52 20.14 3.01995E‐06 6.08218E‐05200 0.2 6.1 20.2 7.94328E‐07 1.60454E‐05 170 0.17 4.27 20.17 5.37032E‐05 0.001083193220 0.22 5.98 20.22 1.04713E‐06 2.11729E‐05 180 0.18 3.9 20.18 0.000125893 0.002540511240 0.24 5.9 20.24 1.25893E‐06 2.54807E‐05 190 0.19 3.68 20.19 0.00020893 0.004218289260 0.26 5.68 20.26 2.0893E‐06 4.23291E‐05 220 0.22 3.35 20.22 0.000446684 0.009031942280 0.28 5.34 20.28 4.57088E‐06 9.26975E‐05 280 0.28 3.04 20.28 0.000912011 0.01849558300 0.3 4.82 20.3 1.51356E‐05 0.000307253 380 0.38 2.78 20.38 0.001659587 0.033822381310 0.31 4.45 20.31 3.54813E‐05 0.000720626 480 0.48 2.62 20.48 0.002398833 0.049128098320 0.32 4.17 20.32 6.76083E‐05 0.001373801340 0.34 3.77 20.34 0.000169824 0.003454228380 0.38 3.37 20.38 0.00042658 0.008693691450 0.45 3.06 20.45 0.000870964 0.017811205530 0.53 2.84 20.53 0.00144544 0.029674878650 0.65 2.68 20.65 0.002089296 0.043143965750 0.75 2.58 20.75 0.002630268 0.054578061A B A B0.1254 0.0386 0.1551 0.0252x y x y0.307815 0 0.1624758 0POI at V = 0.31 mL POI at V = 0.16 mLID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)ID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)Acid #3 20 0.31 0.122 3.75534E‐05 0.001877671 1.88 Acid #3 20 0.16 0.122 1.98221E‐05 0.000991103 0.99Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function0 0 7.42 20 3.80189E‐08 7.60379E‐07 0 0 7.42 20 3.80189E‐08 7.60379E‐07100 0.1 6.47 20.1 3.38844E‐07 6.81077E‐06 100 0.1 6.55 20.1 2.81838E‐07 5.66495E‐06160 0.16 5.9 20.16 1.25893E‐06 2.53799E‐05 160 0.16 5.94 20.16 1.14815E‐06 2.31468E‐05200 0.2 5.09 20.2 8.12831E‐06 0.000164192 190 0.19 5.5 20.19 3.16228E‐06 6.38464E‐05220 0.22 4.34 20.22 4.57088E‐05 0.000924232 210 0.21 4.99 20.21 1.02329E‐05 0.000206808230 0.23 3.99 20.23 0.000102329 0.002070122 220 0.22 4.54 20.22 2.88403E‐05 0.000583151240 0.24 3.76 20.24 0.00017378 0.003517309 240 0.24 3.85 20.24 0.000141254 0.002858976260 0.26 3.49 20.26 0.000323594 0.006556007 280 0.28 3.36 20.28 0.000436516 0.008852541320 0.32 3.12 20.32 0.000758578 0.015414296 360 0.36 3.01 20.36 0.000977237 0.01989655420 0.42 2.84 20.42 0.00144544 0.02951588 460 0.46 2.8 20.46 0.001584893 0.032426915520 0.52 2.68 20.52 0.002089296 0.042872357 560 0.56 2.63 20.56 0.002344229 0.048197344A B A B0.1412 0.0302 0.1393 0.0305x y x y0.213881 0 0.2189519 0POI at V = 0.21 mL POI at V = 0.22 mLID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)ID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)Acid #3 20 0.21 0.122 2.60935E‐05 0.001304674 1.30 Acid #3 20 0.22 0.122 2.67121E‐05 0.001335607 1.34Sample 5 Sample 6Gran function alkalinity calculations Gran function alkalinity calculationsSample 7Gran function alkalinity calculationsSample 8Gran function alkalinity calculationsy = 0.1659x ‐ 0.015700.010.020.030.040.050.060.070.080.090.10 0.1 0.2 0.3 0.4 0.5 0.6 0.7Gran functionVolume of acid added (mL)y = 0.1659x ‐ 0.015700.010.020.030.040.050.060.070.080.090.10 0.1 0.2 0.3 0.4 0.5 0.6 0.7Gran functionVolume of acid added (mL)y = 0.1254x ‐ 0.03860.010.0230.040.050.060.1 0.2 0.3 0.4 0.5 0.6 0.7 8Gran functionGran function y = 0.1062x ‐ 0.02420.0050.010.01520.0250.030.0350.040.1 0.2 0.3 0.4 0.5 6Gran functionGran functiony = 0.1551x ‐ 0.02520.010.0230.040.056Gran functiony = 0.1659x ‐ 0.015700.010.020.030.040.050.060.070.080.090.10 0.1 0.2 0.3 0.4 0.5 0.6 0.7Gran functionVolume of acid added (mL)Gran functionGran function y = 0.1062x ‐ 0.02420.0050.010.01520.0250.030.0350.040.1 0.2 0.3 0.4 0.5 6Gran functionGran functionGran functionGran functionGran function y = 0.1412x ‐ 0.03020.0 510.01520.0230.03540.0455Gran functiony = 0.1659x ‐ 0.015700.010.020.030.040.050.060.070.080.090.10 0.1 0.2 0.3 0.4 0.5 0.6 0.7Gran functionVolume of acid added (mL)y = 0.1162x ‐ 0.01960.010.0230.040.050.06Gran functionGran functiony = 0.1393x ‐ 0.03050.1 0.2 0.3 0.4 0.5 6Gran functionAppendix D144Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function0 0 7.64 20 2.29087E‐08 4.58174E‐07 0 0 7.62 20 2.39883E‐08 4.79767E‐07100 0.1 6.35 20.1 4.46684E‐07 8.97834E‐06 100 0.1 6.62 20.1 2.39883E‐07 4.82165E‐06170 0.17 4.91 20.17 1.23027E‐05 0.000248145 160 0.16 6.12 20.16 7.58578E‐07 1.52929E‐05180 0.18 4.46 20.18 3.46737E‐05 0.000699715 200 0.2 5.68 20.2 2.0893E‐06 4.22038E‐05190 0.19 4.05 20.19 8.91251E‐05 0.001799436 220 0.22 5.36 20.22 4.36516E‐06 8.82635E‐05210 0.21 3.58 20.21 0.000263027 0.005315772 230 0.23 5.11 20.23 7.76247E‐06 0.000157035250 0.25 3.19 20.25 0.000645654 0.013074498 240 0.24 4.75 20.24 1.77828E‐05 0.000359924350 0.35 2.81 20.35 0.001548817 0.031518418 250 0.25 4.24 20.25 5.7544E‐05 0.001165266450 0.45 2.62 20.45 0.002398833 0.049056133 270 0.27 3.64 20.27 0.000229087 0.004643589310 0.31 3.2 20.31 0.000630957 0.012814744380 0.38 2.88 20.38 0.001318257 0.026866072480 0.48 2.64 20.48 0.002290868 0.04691697A B A B0.1825 0.0328 0.2011 0.0496x y x y0.179726 0 0.2466435 0POI at V = 0.18 mL POI at V = 0.25 mLID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)ID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)Acid #3 20 0.18 0.122 2.19266E‐05 0.001096329 1.10 Acid #3 20 0.25 0.122 3.00905E‐05 0.001504525 1.50Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function0 0 7.63 20 2.34423E‐08 4.68846E‐07 0 0 7.76 20 1.7378E‐08 3.4756E‐07100 0.1 6.83 20.1 1.47911E‐07 2.97301E‐06 100 0.1 6.9 20.1 1.25893E‐07 2.53044E‐06170 0.17 6.39 20.17 4.0738E‐07 8.21686E‐06 180 0.18 6.44 20.18 3.63078E‐07 7.32692E‐06280 0.28 5.97 20.28 1.07152E‐06 2.17304E‐05 240 0.24 6.11 20.24 7.76247E‐07 1.57112E‐05290 0.29 5.28 20.29 5.24807E‐06 0.000106483 290 0.29 5.75 20.29 1.77828E‐06 3.60813E‐05320 0.32 4.42 20.32 3.80189E‐05 0.000772545 330 0.33 5.32 20.33 4.7863E‐06 9.73055E‐05330 0.33 4.07 20.33 8.51138E‐05 0.001730364 350 0.35 4.92 20.35 1.20226E‐05 0.000244661340 0.34 3.8 20.34 0.000158489 0.003223673 370 0.37 4.37 20.37 4.2658E‐05 0.000868942360 0.36 3.47 20.36 0.000338844 0.006898867 380 0.38 4.05 20.38 8.91251E‐05 0.001816369400 0.4 3.15 20.4 0.000707946 0.014442094 400 0.4 3.62 20.4 0.000239883 0.004893619480 0.48 2.86 20.48 0.001380384 0.02827027 440 0.44 3.22 20.44 0.00060256 0.012316318580 0.58 2.65 20.58 0.002238721 0.046072881 520 0.52 2.9 20.52 0.001258925 0.025833149620 0.62 2.68 20.62 0.002089296 0.043081286A B A B0.178 0.0571 0.1724 0.0638x y x y0.3207865 0 0.3700696 0POI at V = 0.32 mL POI at V = 0.37 mLID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)ID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)Acid #3 20 0.32 0.122 3.9136E‐05 0.001956798 1.96 Acid #3 20 0.37 0.122 4.51485E‐05 0.002257425 2.26Gran function alkalinity calculations Gran function alkalinity calculationsSample 9 Sample 10Gran function alkalinity calculations Gran function alkalinity calculationsSample 11 Sample 12y = 0.1535x ‐ 0.022700.010.020.030.040.050.060 0.1 0.2 0.3 0.4 0.5 0.6Gran functionVolume of acid added (mL)y = 0.1162x ‐ 0.019600.010.020.030.040.050.060 0.1 0.2 0.3 0.4 0.5 0.6 0.7Gran functionVolume of acid added (mL)y = 0.1659x ‐ 0.01570.010.020.030.0450.060.070.080.090.1Gran functiony = 0.2011x ‐ 0.04960.0 510.01520.0230.03540.04550.1 0.2 0.3 0.4 0.5 6Gran functiony = 0.1062x ‐ 0.02420.0050.010.01520.0250.030.0354Gran functiony = 0.1659x ‐ 0.01570.010.020.030.0450.060.070.080.090.10.1 0.2 0.3 0.4 0.5 0.6 7Gran function y = 0.1825x ‐ 0.03280.1 0.2 0.3 0.4 5Gran functiony = 0.1062x ‐ 0.024200.0050.010.0150.020.0250.030.0350.040 0.1 0.2 0.3 0.4 0.5 0.6Gran functionVolume of acid added (mL)y = 0.1659x ‐ 0.015700.010.020.030.040.050.060.070.080.090.10 0.1 0.2 0.3 0.4 0.5 0.6 0.7Gran functionVolume of acid added (mL)y = 0.1724x ‐ 0.06380.0 510.01520.0230.03540.0450.05Gran functiony = 0.1659x ‐ 0.01570.010.020.030.0450.060.070.080.090.10.1 0.2 0.3 0.4 0.5 0.6 7Gran functiony = 0.178x ‐ 0.05710.0 510.01520.0230.03540.0455Gran functionAppendix D145Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function0 0 7.65 20 2.23872E‐08 4.47744E‐07 0 0 7.89 20 1.28825E‐08 2.5765E‐07100 0.1 6.98 20.1 1.04713E‐07 2.10473E‐06 100 0.1 6.85 20.1 1.41254E‐07 2.8392E‐06180 0.18 6.65 20.18 2.23872E‐07 4.51774E‐06 170 0.17 6.32 20.17 4.7863E‐07 9.65397E‐06260 0.26 6.26 20.26 5.49541E‐07 1.11337E‐05 210 0.21 6.05 20.21 8.91251E‐07 1.80122E‐05340 0.34 5.77 20.34 1.69824E‐06 3.45423E‐05 250 0.25 5.68 20.25 2.0893E‐06 4.23082E‐05400 0.4 5.08 20.4 8.31764E‐06 0.00016968 270 0.27 5.42 20.27 3.80189E‐06 7.70644E‐05420 0.42 4.63 20.42 2.34423E‐05 0.000478692 290 0.29 4.86 20.29 1.38038E‐05 0.00028008430 0.43 4.35 20.43 4.46684E‐05 0.000912575 300 0.3 4.38 20.3 4.16869E‐05 0.000846245450 0.45 3.83 20.45 0.000147911 0.003024777 310 0.31 4.06 20.31 8.70964E‐05 0.001768927490 0.49 3.32 20.49 0.00047863 0.009807131 320 0.32 3.96 20.32 0.000109648 0.002228044560 0.56 2.97 20.56 0.001071519 0.022030437 340 0.34 3.52 20.34 0.000301995 0.006142582600 0.6 2.74 20.6 0.001819701 0.037485838 360 0.36 3.3 20.36 0.000501187 0.010204172400 0.4 3.06 20.4 0.000870964 0.017767657470 0.47 2.83 20.47 0.001479108 0.030277349570 0.57 2.63 20.57 0.002344229 0.048220787A B A B0.173 0.0749 0.1833 0.056x y x y0.432948 0 0.3055101 0POI at V = 0.43 mL POI at V = 0.31 mLID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)ID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)Acid #3 20 0.43 0.122 5.28197E‐05 0.002640983 2.64 Acid #3 20 0.31 0.122 3.72722E‐05 0.001863612 1.86Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function Acid added (uL) Acid added (mL) pH Tot. Vol. 10^‐pH Gran Function0 0 7.82 20 1.51356E‐08 3.02712E‐07 0 0 7.73 20 1.86209E‐08 3.72417E‐07100 0.1 6.95 20.1 1.12202E‐07 2.25526E‐06 100 0.1 7.12 20.1 7.58578E‐08 1.52474E‐06180 0.18 6.58 20.18 2.63027E‐07 5.30788E‐06 200 0.2 6.73 20.2 1.86209E‐07 3.76142E‐06240 0.24 6.14 20.24 7.24436E‐07 1.46626E‐05 300 0.3 6.33 20.3 4.67735E‐07 9.49502E‐06280 0.28 5.85 20.28 1.41254E‐06 2.86463E‐05 400 0.4 5.78 20.4 1.65959E‐06 3.38556E‐05310 0.31 5.53 20.31 2.95121E‐06 5.99391E‐05 460 0.46 5.16 20.46 6.91831E‐06 0.000141549340 0.34 4.93 20.34 1.1749E‐05 0.000238974 480 0.48 4.71 20.48 1.94984E‐05 0.000399328350 0.35 4.5 20.35 3.16228E‐05 0.000643524 490 0.49 4.34 20.49 4.57088E‐05 0.000936574360 0.36 4.03 20.36 9.33254E‐05 0.001900106 500 0.5 4.02 20.5 9.54993E‐05 0.001957735370 0.37 3.73 20.37 0.000186209 0.003793071 520 0.52 3.65 20.52 0.000223872 0.004593856390 0.39 3.42 20.39 0.000380189 0.007752062 560 0.56 3.29 20.56 0.000512861 0.01054443450 0.45 3.02 20.45 0.000954993 0.019529598 640 0.64 2.97 20.64 0.001071519 0.022116158550 0.55 2.78 20.55 0.001659587 0.034104511 740 0.74 2.77 20.74 0.001698244 0.035221573A B A B0.1962 0.0688 0.1392 0.0675x y x y0.3506626 0 0.4849138 0POI at V = 0.35 mL POI at V = 0.48 mLID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)ID Sample Vol. (mL)Acid Req. (mL)Acid Conc. (N)mol. of protons added Alk. (N)Alk. (meq/L)Acid #3 20 0.35 0.122 4.27808E‐05 0.002139042 2.14 Acid #3 20 0.48 0.122 5.91595E‐05 0.002957974 2.96Sample 14Gran function alkalinity calculationsSample 16Gran function alkalinity calculationsSample 13Gran function alkalinity calculationsSample 15Gran function alkalinity calculationsy = 0.1659x ‐ 0.015700.010.020.030.040.050.060.070.080.090.10 0.1 0.2 0.3 0.4 0.5 0.6 0.7Gran functionVolume of acid added (mL)Gran function y = 0.1062x ‐ 0.02420.0050.010.01520.0250.030.0350.040.1 0.2 0.3 0.4 0.5 6Gran functionGran functiony = 0.1551x ‐ 0.02520.010.0230.040.056Gran functiony = 0.1162x ‐ 0.0196Gran functionGran functiony = 0.173x ‐ 0.0749Gran functiony = 0.1659x ‐ 0.015700.010.020.030.040.050.060.070.080.090.10 0.1 0.2 0.3 0.4 0.5 0.6 0.7Gran functionVolume of acid added (mL)y = 0.1162x ‐ 0.01960.010.0230.040.050.06Gran functionGran functiony = 0.1254x ‐ 0.03860.1 0.2 0.3 0.4 0.5 0.6 0.7 8Gran functionGran functionGran functionGran functiony = 0.1962x ‐ 0.06880.0050.010.01520.0250.030.03540.1 0.2 3 0.4 0.5 6Gran functiony = 0.1659x ‐ 0.015700.010.020.030.040.050.060.070.080.090.10 0.1 0.2 0.3 0.4 0.5 0.6 0.7Gran functionVolume of acid added (mL)Gran functionGran function y = 0.1062x ‐ 0.02420.0050.010.01520.0250.030.0350.040.1 0.2 0.3 0.4 0.5 6Gran functionGran functionGran functionGran functionGran function y = 0.1412x ‐ 0.03020.0 510.01520.0230.03540.0455Gran functionGran functionGran function y = 0.1833x ‐ 0.0560.010.0230.040.056Gran functiony = 0.1535x ‐ 0.022700.010.020.030.040.050.060 0.1 0.2 0.3 0.4 0.5 0.6Gran functionVolume of acid added (mL)y = 0.1062x ‐ 0.02420.0050.010.01520.0250.030.0354Gran functiony = 0.1659x ‐ 0.01570.010.020.030.0450.060.070.080.090.10.1 0.2 0.3 0.4 0.5 0.6 7Gran functiony = 0.1254x ‐ 0.03860.1 0.2 0.3 4 0.5 0.6 0.7 8Gran functionGran functionGran functionGran function y = 0.1392x ‐ 0.0675Gran functionAppendix D146Depth (cm)Porosity Tray weight (g)Tray + wet weight (g)Tray + dry weight (g)Water weight (g) Dry weight (g)Dry weight ratioCentrifuge tube (g)Tube + sediment (g)sediment mass (g)Extractant volume (mL)OES dilution factor[Fe] from OES (mg/L)Fe content (mg/g)1w 7 0.19034182 4.1 9.15 8.702 0.448 5.05 0.9191a 7 0.919 10.91 11.48 0.57 40 1 0.03375 0.002578531b 7 0.919 10.87 11.42 0.55 40 1 1.06852w 12.5 0.238226622 4.136 9.161 8.568 0.593 5.025 0.8942a 12.5 0.894 10.913 11.44 0.527 40 1 0.0702 0.0059570632b 12.5 0.894 10.91 11.468 0.558 40 1 0.03805 0.0030494823w 17.5 0.243146294 4.128 9.234 8.615 0.619 5.106 0.8923a 17.5 0.892 10.911 11.436 0.525 40 1 0.0681 0.0058175823b 17.5 0.892 10.912 11.489 0.577 40 1 0.06845 0.0053204984w 22.5 0.242267256 4.129 9.102 8.502 0.6 4.973 0.8924a 22.5 0.892 10.916 11.398 0.482 40 1 0.05465 0.0050824574b 22.5 0.892 10.914 11.406 0.492 40 1 0.0687 0.0062624794c 22.5 Total Fe check 10.914 11.412 0.4985w 27.5 0.266675863 4.131 9.181 8.488 0.693 5.05 0.8795a 27.5 . 0.879 10.94 11.436 0.496 40 1 0.06845 0.006277685b 27.5 0.879 10.927 11.428 0.501 40 1 0.09605 0.0087210166w 32.5 0.30728547 4.115 9.151 8.308 0.843 5.036 0.8576a 32.5 0.857 10.731 11.205 0.474 40 1 0.11885 0.0117084286b 32.5 0.857 10.908 11.391 0.483 40 1 0.0893 0.0086334047w 37.5 0.285401863 4.087 9.07 8.319 0.751 4.983 0.8697a 37.5 0.869 10.914 11.438 0.524 40 1 0.08515 0.0074796317b 37.5 0.869 10.737 11.244 0.507 40 1 0.15055 0.0136678318w 42.5 0.291441044 4.175 9.239 8.453 0.786 5.064 0.8668a 42.5 0.866 10.911 11.398 0.487 40 1 0.42235 0.0400742778b 42.5 0.869 10.873 11.415 0.542 40 1 0.3399 0.0288654728c 42.5 Total Fe check 10.917 11.407 0.499w 47.5 0.233998308 4.142 9.078 8.509 0.569 4.936 0.8979a 47.5 0.897 10.792 11.327 0.535 40 1 0.9475 0.0790073699b 47.5 0.897 10.734 11.258 0.524 40 1 1.0785 0.09181867610w 52.5 0.288653442 4.182 9.204 8.435 0.769 5.022 0.86710a 52.5 0.867 10.904 11.431 0.527 40 1 1.2805 0.11207424310b 52.5 0.867 10.793 11.335 0.542 40 1 1.1585 0.09859016611w 57.5 0.29666229 4.138 9.227 8.417 0.81 5.089 0.86311a 57.5 0.863 10.79 11.338 0.548 40 1 1.328 0.11236303311b 57.5 0.863 10.867 11.384 0.517 40 1 1.2345 0.11071500612w 62.5 0.257006009 4.089 9.084 8.432 0.652 4.995 0.88512a 62.5 0.885 10.871 11.372 0.501 40 1 0.95755 0.086430312b 62.5 0.863 10.915 11.418 0.503 40 1 0.96535 0.08898621912c 62.5 Total Fe check 10.91 11.454 0.54413w 67.5 0.280358522 4.107 9.093 8.36 0.733 4.986 0.87213a 67.5 0.872 10.909 11.408 0.499 40 1 1.94 0.17837295113b 67.5 0.872 10.914 11.417 0.503 40 1 1.8615 0.16979420714w 72.5 0.257331871 4.146 8.513 7.942 0.571 4.367 0.88414a 72.5 0.884 10.908 11.437 0.529 40 1 1.1225 0.09597509814b 72.5 0.884 10.908 11.442 0.534 40 1 1.0915 0.09245073515w 77.5 0.303227605 4.228 8.728 7.989 0.739 4.5 0.85915a 77.5 0.859 10.914 11.409 0.495 40 1 1.476 0.1388599615b 77.5 0.859 10.873 11.385 0.512 40 1 1.557 0.14161671916w 82.5 0.327140717 4.213 9.222 8.303 0.919 5.009 0.84516a 82.5 0.845 10.988 11.485 0.497 40 1 1.6845 0.16044706716b 82.5 0.859 10.908 11.418 0.51 40 1 1.7215 0.15719282816c 82.5 Total Fe check 10.88 11.417 0.537Step 11151M CaCl2 extractable Fe16621014893111213457Appendix E147Step 4Total digestable FeExtractant volume (mL)OES dilution factor[Fe] from OES (mg/L)Fe content (mg/g)Hach dil. factor[Fe(II)] from Hach (mg/L)Fe(II) cont. (mg/g)Fe(II)/Fe(tot) ratioExtractant volume (mL)OES dilution factor[Fe] from OES (mg/L)Fe content (mg/g)Fe content (mg/g) Sum of Steps 1‐3Sum of Steps 1‐4 for total Fe checkDifferencePost‐extraction mass (g)40 10 41.77 3.191265729 2.5 0.21 0.040110474 0.0126 40 5 23.395 1.787399131 16.7 4.981243391 21.68124339 11.41440 10 39.375 3.117677768 2.5 0.11 0.021774257 0.0070 40 5 21.21 1.679389091 16.6 4.797066859 21.39706686 11.34540 10 36.545 3.101151699 2.5 0.18 0.038186298 0.0123 40 5 23.195 1.968291522 16.8 5.075400283 21.87540028 11.3540 10 40.37 3.235416609 2.5 0.21 0.042075643 0.0130 40 5 22.55 1.807249059 16.6 5.04571515 21.64571515 11.37640 10 36.235 3.095448865 2.5 0.12 0.025628113 0.0083 40 5 21.115 1.803791991 15.5 4.905058437 20.40505844 11.37640 10 46.635 3.624856678 2.5 0.13 0.025261679 0.0070 40 5 25.965 2.018213866 17.3 5.648391042 22.94839104 11.41340 10 35.54 3.305224478 2.5 0.16 0.03720005 0.0113 40 5 20.09 1.868372531 15.2 5.178679466 20.37867947 4.3% 11.33940 10 33.565 3.0596815 2.5 0.16 0.036462762 0.0119 40 5 18.015 1.642191635 15.6 4.708135614 20.30813561 4.7% 11.34321.3 21.3 11.35140 10 36.16 3.316302779 2.5 0.12 0.027513574 0.0083 40 5 19.79 1.814978761 17.4 5.13755922 22.53755922 11.38540 10 36.1 3.277758147 2.5 0.2 0.045398312 0.0139 40 5 19.365 1.758276635 16.7 5.044755798 21.7447558 11.36140 10 32.635 3.215015014 2.5 0.16 0.03940573 0.0123 40 5 17.61 1.734837273 15.6 4.961560715 20.56156071 11.12840 10 37.12 3.588711669 2.5 0.18 0.043505395 0.0121 40 5 20.28 1.960643121 15.9 5.557988194 21.45798819 11.32240 10 39.225 3.445549218 2.5 0.17 0.037332273 0.0108 40 5 19.895 1.747589591 14.8 5.20061844 20.00061844 11.38540 10 40.505 3.677286522 2.5 0.21 0.047662645 0.0130 40 5 20.965 1.903328279 15.5 5.594282631 21.09428263 11.19940 10 33.1 3.14066195 2.5 0.11 0.026093113 0.0083 40 5 1.718850142 18 4.899586369 22.89958637 10.2% 11.33340 10 36.445 3.09503426 2.5 0.18 0.038215542 0.0123 40 5 20.24 1.718850142 17.1 4.842749875 21.94274988 14.0% 11.33325.5 25.5 11.34540 10 40.405 3.369174404 2.5 0.31 0.064623442 0.0192 40 5 20.85 1.738579045 19.4 5.186760819 24.58676082 11.23940 10 35.895 3.055940079 2.5 0.21 0.044696157 0.0146 40 5 20.145 1.715055381 19.9 4.862814135 24.76281414 11.1540 10 39.21 3.431808732 2.5 0.23 0.050326193 0.0147 40 5 21.605 1.890951993 17.5 5.434834969 22.93483497 11.33940 10 44.46 3.783615707 2.5 0.2 0.042550784 0.0112 40 5 25.15 2.140304432 18.6 6.022510305 24.62251031 11.2540 10 46.905 3.968665707 2.5 0.23 0.048651163 0.0123 40 5 23.87 2.019657828 16.9 6.100686567 23.00068657 11.24640 10 40.06 3.592744544 2.5 0.38 0.085199883 0.0237 40 5 19.795 1.775296511 15.7 5.478756061 21.17875606 11.29640 10 46.025 4.154304804 2.5 0.27 0.060926795 0.0147 40 5 22.99 2.075121509 16.8 6.315856613 23.11585661 4.1% 11.31340 10 45.865 4.227847847 2.5 0.2 0.046090132 0.0109 40 5 20.815 1.918732213 15.4 6.235566278 21.63556628 10.2% 11.36324.1 24.1 11.39740 10 46.07 4.235897869 2.5 0.27 0.062062754 0.0147 40 5 19.575 1.799819856 14.3 6.214090676 20.51409068 11.31840 10 39.695 3.620725786 2.5 0.32 0.072970919 0.0202 40 5 19.46 1.775017604 13.4 5.565537597 18.9655376 11.32240 10 41.16 3.519229416 2.5 0.22 0.04702566 0.0134 40 5 21.16 1.809205404 18 5.424409917 23.42440992 11.37640 10 45.27 3.834397409 2.5 0.32 0.067760502 0.0177 40 5 22.56 1.910846157 17.9 5.837694301 23.7376943 11.37640 10 38.43 3.615439192 2.5 0.33 0.077614815 0.0215 40 5 20.2 1.900386981 15.6 5.654686132 21.25468613 11.33340 10 45.96 4.180285417 2.5 0.29 0.065942274 0.0158 40 5 21.21 1.929152604 16.1 6.25105474 22.35105474 11.31440 10 44.525 4.240965056 2.5 0.36 0.085724167 0.0202 40 5 19.52 1.85926194 15.6 6.260674062 21.86067406 6.2% 11.440 10 44.31 4.046014641 2.5 0.23 0.052504139 0.0130 40 5 21.42 1.955893333 15.6 6.159100802 21.7591008 6.6% 11.32323.3 23.3 11.336Step 31M NH2OH∙HCl in 25% CH3COOH extractable Fe0.5M HCl extractable Fe(tot) 0.5M HCl extractable Fe(II)Step 2Appendix E148Depth (cm)Notes Tray weight (g)Tray + wet weight (g)Tray + dry weight (g)Water weight (g) Dry weight (g)Dry weight ratioCentrifuge tube (g)Tube + sediment (g)sediment mass (g)Extractant volume (mL)OES dilution factor[Mn] from OES (mg/L)Mn content (mg/g)1w 7 4.1 9.15 8.702 0.448 5.05 0.9191a 7 0.919 10.91 11.48 0.57 40 1 0.23495 0.0179503921b 7 0.919 10.87 11.42 0.55 40 1 0.2595 0.0205469812w 12.5 4.136 9.161 8.568 0.593 5.025 0.8942a 12.5 0.894 10.913 11.44 0.527 40 1 0.15295 0.0129790982b 12.5 0.894 10.91 11.468 0.558 40 1 0.25795 0.0206731663w 17.5 4.128 9.234 8.615 0.619 5.106 0.8923a 17.5 0.892 10.911 11.436 0.525 40 1 0.26055 0.0222580163b 17.5 0.892 10.912 11.489 0.577 40 1 0.34805 0.0270533154w 22.5 4.129 9.102 8.502 0.6 4.973 0.8924a 22.5 0.892 10.916 11.398 0.482 40 1 0.38695 0.0359863994b 22.5 0.892 10.914 11.406 0.492 40 1 0.3361 0.0306378364c 22.5 Total Fe check 10.914 11.412 0.4985w 27.5 4.131 9.181 8.488 0.693 5.05 0.8795a 27.5 . 0.879 10.94 11.436 0.496 40 1 0.44985 0.0412566045b 27.5 0.879 10.927 11.428 0.501 40 1 0.52055 0.0472641836w 32.5 4.115 9.151 8.308 0.843 5.036 0.8576a 32.5 0.857 10.731 11.205 0.474 40 1 0.4669 0.0459963396b 32.5 0.857 10.908 11.391 0.483 40 1 0.49785 0.0481314687w 37.5 4.087 9.07 8.319 0.751 4.983 0.8697a 37.5 0.869 10.914 11.438 0.524 40 1 0.4814 0.0422864867b 37.5 0.869 10.737 11.244 0.507 40 1 0.49855 0.0452613558w 42.5 4.175 9.239 8.453 0.786 5.064 0.8668a 42.5 0.866 10.911 11.398 0.487 40 1 0.4165 0.0395192068b 42.5 0.869 10.873 11.415 0.542 40 1 0.43725 0.0371327688c 42.5 Total Fe check 10.917 11.407 0.499w 47.5 4.142 9.078 8.509 0.569 4.936 0.8979a 47.5 0.897 10.792 11.327 0.535 40 1 0.343 0.0286010859b 47.5 0.897 10.734 11.258 0.524 40 1 0.32575 0.02773290110w 52.5 4.182 9.204 8.435 0.769 5.022 0.86710a 52.5 0.867 10.904 11.431 0.527 40 1 0.34075 0.02982373910b 52.5 0.867 10.793 11.335 0.542 40 1 0.36205 0.03081102311w 57.5 4.138 9.227 8.417 0.81 5.089 0.86311a 57.5 0.863 10.79 11.338 0.548 40 1 0.3832 0.03242282711b 57.5 0.863 10.867 11.384 0.517 40 1 0.3287 0.0294791612w 62.5 4.089 9.084 8.432 0.652 4.995 0.88512a 62.5 0.885 10.871 11.372 0.501 40 1 0.33845 0.03054914612b 62.5 0.863 10.915 11.418 0.503 40 1 0.3234 0.02981109812c 62.5 Total Fe check 10.91 11.454 0.54413w 67.5 4.107 9.093 8.36 0.733 4.986 0.87213a 67.5 0.872 10.909 11.408 0.499 40 1 0.30635 0.02816729613b 67.5 0.872 10.914 11.417 0.503 40 1 0.25095 0.02289006514w 72.5 4.146 8.513 7.942 0.571 4.367 0.88414a 72.5 0.884 10.908 11.437 0.529 40 1 0.19315 0.01651455714b 72.5 0.884 10.908 11.442 0.534 40 1 0.1873 0.01586442815w 77.5 4.228 8.728 7.989 0.739 4.5 0.85915a 77.5 0.859 10.914 11.409 0.495 40 1 0.1934 0.01819479415b 77.5 0.859 10.873 11.385 0.512 40 1 0.2121 0.01929152616w 82.5 4.213 9.222 8.303 0.919 5.009 0.84516a 82.5 0.845 10.988 11.485 0.497 40 1 0.1977 0.01883074216b 82.5 0.859 10.908 11.418 0.51 40 1 0.20865 0.01905215416c 82.5 Total Fe check 10.88 11.417 0.5371M CaCl2 extractable Mn3Step 11245678910111314151612Appendix E149Step 4Total digestable MnExtractant volume (mL)OES dilution factor[Mn] from OES (mg/L)Mn content (mg/g)Extractant volume (mL)OES dilution factor[Mn] from OES (mg/L)Mn content (mg/g)Mn content (mg/g)Sum of Steps 1‐3Sum of Steps 1‐4 for total Fe Difference Post‐extraction mass (g)40 10 1.7 0.129881536 40 5 0.4294 0.032806548 0.314 0.180638476 0.494638476 11.41440 10 1.627 0.128824425 40 5 0.39335 0.031145106 0.32 0.180516512 0.500516512 11.34540 10 1.5325 0.13004556 40 5 0.4163 0.035326569 0.322 0.178351227 0.500351227 11.3540 10 1.576 0.126307074 40 5 0.40855 0.032742865 0.316 0.179723105 0.495723105 11.37640 10 1.4745 0.125962173 40 5 0.38875 0.033209763 0.298 0.181429952 0.479429952 11.37640 10 1.843 0.143253154 40 5 0.45215 0.035144826 0.338 0.205451296 0.543451296 11.41340 10 1.233 0.114669155 40 5 0.3665 0.034084546 0.286 0.1847401 0.4707401 3.3% 11.33940 10 1.305 0.11895976 40 5 0.34245 0.031216682 0.337 0.180814278 0.517814278 ‐6.3% 11.3430.487 0.487 11.35140 10 1.209 0.110879703 40 5 0.37345 0.034249814 0.336 0.186386121 0.522386121 11.38540 10 1.1225 0.101919211 40 5 0.36075 0.032754882 0.308 0.181938276 0.489938276 11.36140 10 1.0195 0.100435355 40 5 0.34425 0.033913557 0.295 0.18034525 0.47534525 11.12840 10 1.1635 0.112485615 40 5 0.36915 0.035688925 0.299 0.196306009 0.495306009 11.32240 10 1.266 0.111206254 40 5 0.36735 0.03226826 0.279 0.185761 0.464761 11.38540 10 1.3285 0.120609188 40 5 0.38275 0.034748338 0.295 0.200618881 0.495618881 11.19940 10 1.1035 0.104704546 40 5 0.031570284 0.379 0.175794035 0.554794035 ‐6.1% 11.33340 10 1.054 0.089509291 40 5 0.37175 0.031570284 0.328 0.158212343 0.486212343 7.0% 11.3330.523 0.523 11.34540 10 1.024 0.085386328 40 5 0.37225 0.031040098 0.381 0.145027511 0.526027511 11.23940 10 0.9027 0.076851849 40 5 0.37175 0.031649136 0.394 0.136233885 0.530233885 11.1540 10 1.0087 0.088285271 40 5 0.3834 0.03355663 0.328 0.15166564 0.47966564 11.33940 10 1.119 0.095228654 40 5 0.43745 0.037227681 0.343 0.163267358 0.506267358 11.2540 10 1.254 0.10610184 40 5 0.42545 0.03599763 0.314 0.174522297 0.488522297 11.24640 10 1.041 0.093361135 40 5 0.3656 0.032788502 0.309 0.155628797 0.464628797 11.29640 10 1.0795 0.097437741 40 5 0.39575 0.035721154 0.301 0.163708042 0.464708042 2.0% 11.31340 10 1.0335 0.095268304 40 5 0.3535 0.032585724 0.276 0.157665125 0.433665125 8.5% 11.3630.474 0.474 11.39740 10 1.161 0.106747936 40 5 0.33675 0.030962418 0.262 0.16587765 0.42787765 11.31840 10 1.0105 0.092171392 40 5 0.3529 0.032189297 0.25 0.147250754 0.397250754 11.32240 10 0.96735 0.082709586 40 5 0.3835 0.03278971 0.34 0.132013854 0.472013854 11.37640 10 1.0825 0.091688429 40 5 0.40295 0.034130118 0.347 0.141682975 0.488682975 11.37640 10 0.8797 0.082760912 40 5 0.3627 0.034122295 0.331 0.135078001 0.466078001 11.33340 10 1.115 0.10141467 40 5 0.37605 0.034203576 0.306 0.154909772 0.460909772 11.31440 10 1.054 0.100392525 40 5 0.3518 0.033508625 0.313 0.152731891 0.465731891 ‐2.8% 11.440 10 1.199 0.109482545 40 5 0.38415 0.035077331 0.281 0.16361203 0.44461203 1.9% 11.3230.453 0.453 11.336Step 30.5M HCl extractable Mn 1M NH2OH∙HCl in 25% CH3COOH extractable MnStep 2Appendix E1507 12.5 17.5 22.5 27.5 32.5 37.5 42.5 47.5 52.5 57.5 62.5 67.5 72.5 77.5 82.5 BlankAcidobacteria 21.92% 23.23% 21.93% 22.38% 24.68% 24.31% 21.73% 20.51% 14.21% 12.58% 10.01% 8.39% 8.04% 8.00% 8.87% 10.43% 0.00%Acidobacteria 21.35% 22.78% 21.08% 21.64% 23.80% 23.54% 20.89% 19.81% 13.31% 11.67% 8.86% 7.45% 7.25% 7.01% 7.86% 9.55% 0.00%AT‐s3‐28 0.10% 0.08% 0.20% 0.08% 0.21% 0.12% 0.13% 0.14% 0.09% 0.05% 0.01% 0.02% 0.05% 0.01% 0.03% 0.06% 0.00%Subgroup_11 0.09% 0.05% 0.05% 0.06% 0.09% 0.10% 0.08% 0.07% 0.00% 0.01% 0.06% 0.05% 0.06% 0.03% 0.04% 0.05% 0.00%Subgroup_15 0.19% 0.12% 0.16% 0.23% 0.22% 0.24% 0.19% 0.12% 0.14% 0.04% 0.04% 0.08% 0.04% 0.03% 0.10% 0.14% 0.00%Subgroup_17 2.06% 2.37% 2.90% 2.52% 3.41% 3.07% 3.06% 2.74% 1.51% 1.28% 1.23% 1.41% 1.08% 1.04% 1.00% 1.25% 0.00%Subgroup_18 0.08% 0.04% 0.08% 0.16% 0.14% 0.12% 0.17% 0.24% 0.41% 0.36% 0.24% 0.45% 0.52% 0.40% 0.32% 0.30% 0.00%Subgroup_3 1.32% 1.26% 1.09% 1.15% 1.39% 1.07% 1.06% 0.86% 0.73% 0.45% 0.37% 0.36% 0.33% 0.44% 0.40% 0.46% 0.00%Subgroup_4 9.96% 11.31% 8.36% 8.59% 7.98% 8.44% 7.18% 6.77% 4.26% 3.61% 2.43% 1.65% 1.35% 1.45% 2.21% 2.57% 0.00%Subgroup_5 0.20% 0.32% 0.19% 0.17% 0.24% 0.19% 0.21% 0.15% 0.14% 0.08% 0.12% 0.05% 0.05% 0.09% 0.09% 0.09% 0.00%Subgroup_6 7.35% 7.22% 8.06% 8.67% 10.13% 10.19% 8.80% 8.70% 6.03% 5.78% 4.37% 3.36% 3.77% 3.52% 3.67% 4.65% 0.00%Holophagae 0.57% 0.46% 0.85% 0.74% 0.88% 0.77% 0.84% 0.70% 0.90% 0.91% 1.15% 0.94% 0.79% 0.99% 1.01% 0.88% 0.00%Holophagae_Incertae_Sedis 0.00% 0.00% 0.04% 0.01% 0.01% 0.03% 0.05% 0.06% 0.24% 0.33% 0.58% 0.60% 0.36% 0.69% 0.55% 0.44% 0.00%Unknown_FamilyThermoanaerobaculum 0.00% 0.00% 0.04% 0.01% 0.01% 0.03% 0.05% 0.06% 0.24% 0.33% 0.58% 0.60% 0.36% 0.69% 0.55% 0.44% 0.00%Subgroup_10 0.45% 0.32% 0.48% 0.56% 0.53% 0.55% 0.50% 0.42% 0.29% 0.35% 0.27% 0.11% 0.17% 0.14% 0.27% 0.22% 0.00%Subgroup_7 0.12% 0.13% 0.33% 0.17% 0.33% 0.19% 0.30% 0.22% 0.37% 0.23% 0.30% 0.23% 0.26% 0.16% 0.19% 0.22% 0.00%Actinobacteria 3.31% 3.33% 2.75% 3.85% 4.71% 4.73% 4.87% 4.81% 5.06% 6.18% 4.43% 4.16% 3.69% 3.02% 4.08% 4.01% 1.43%Acidimicrobiia 1.32% 1.35% 0.90% 1.56% 1.59% 1.97% 1.87% 1.54% 2.06% 2.07% 1.54% 1.33% 1.37% 0.92% 1.67% 1.50% 0.48%Acidimicrobiales 1.32% 1.35% 0.90% 1.56% 1.59% 1.97% 1.87% 1.54% 2.06% 2.07% 1.54% 1.33% 1.37% 0.92% 1.67% 1.50% 0.48%Acidimicrobiaceae 0.38% 0.39% 0.25% 0.42% 0.53% 0.47% 0.62% 0.46% 0.54% 0.41% 0.12% 0.15% 0.22% 0.13% 0.23% 0.24% 0.48%Acidimicrobiales_Incertae_Sedis 0.04% 0.01% 0.01% 0.04% 0.06% 0.02% 0.04% 0.04% 0.05% 0.03% 0.01% 0.02% 0.02% 0.04% 0.00% 0.01% 0.00%Iamiaceae 0.33% 0.28% 0.30% 0.47% 0.42% 0.41% 0.43% 0.34% 0.60% 0.55% 0.26% 0.30% 0.29% 0.14% 0.33% 0.30% 0.00%OM1_clade 0.21% 0.23% 0.19% 0.25% 0.33% 0.45% 0.38% 0.40% 0.46% 0.59% 0.80% 0.54% 0.59% 0.39% 0.59% 0.47% 0.00%Sva0996_marine_group 0.06% 0.08% 0.05% 0.07% 0.11% 0.23% 0.13% 0.12% 0.12% 0.19% 0.18% 0.18% 0.07% 0.08% 0.12% 0.15% 0.00%unclassified 0.29% 0.36% 0.11% 0.32% 0.14% 0.38% 0.27% 0.18% 0.29% 0.31% 0.18% 0.14% 0.18% 0.14% 0.40% 0.34% 0.00%Thermoleophilia 1.99% 1.98% 1.84% 2.29% 3.12% 2.76% 3.00% 3.28% 3.00% 4.11% 2.89% 2.83% 2.32% 2.10% 2.41% 2.51% 0.95%Gaiellales 1.89% 1.78% 1.69% 2.13% 2.91% 2.62% 2.78% 2.89% 2.64% 3.51% 2.50% 2.46% 2.04% 1.81% 2.09% 2.20% 0.48%GaiellaceaeGaiella 1.78% 1.69% 1.57% 2.00% 2.68% 2.45% 2.51% 2.67% 2.54% 3.23% 2.22% 2.22% 1.68% 1.51% 1.86% 1.99% 0.48%unclassified 0.11% 0.09% 0.12% 0.13% 0.23% 0.18% 0.28% 0.22% 0.11% 0.28% 0.28% 0.25% 0.36% 0.30% 0.23% 0.21% 0.00%Solirubrobacterales 0.10% 0.20% 0.15% 0.16% 0.21% 0.14% 0.21% 0.38% 0.35% 0.60% 0.39% 0.36% 0.28% 0.28% 0.32% 0.31% 0.48%Bacteroidetes 12.18% 11.46% 11.34% 11.19% 8.88% 10.50% 9.24% 6.91% 5.80% 4.19% 3.03% 2.87% 2.61% 3.05% 4.47% 5.18% 3.33%Flavobacteriia 0.70% 0.46% 0.41% 0.36% 0.15% 0.18% 0.26% 0.17% 0.15% 0.12% 0.07% 0.07% 0.06% 0.04% 0.06% 0.06% 2.38%Flavobacteriales 0.70% 0.46% 0.41% 0.36% 0.15% 0.18% 0.26% 0.17% 0.15% 0.12% 0.07% 0.07% 0.06% 0.04% 0.06% 0.06% 2.38%Cryomorphaceae 0.03% 0.01% 0.07% 0.04% 0.02% 0.01% 0.02% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.48%Flavobacteriaceae 0.47% 0.31% 0.25% 0.17% 0.04% 0.07% 0.14% 0.08% 0.06% 0.11% 0.06% 0.05% 0.05% 0.04% 0.03% 0.03% 1.90%NS9_marine_group 0.19% 0.13% 0.09% 0.16% 0.09% 0.10% 0.10% 0.08% 0.09% 0.00% 0.01% 0.02% 0.02% 0.00% 0.03% 0.03% 0.00%Sphingobacteriia 11.48% 11.01% 10.93% 10.83% 8.73% 10.32% 8.98% 6.74% 5.65% 4.07% 2.97% 2.80% 2.55% 3.01% 4.41% 5.11% 0.95%Sphingobacteriales 11.48% 11.01% 10.93% 10.83% 8.73% 10.32% 8.98% 6.74% 5.65% 4.07% 2.97% 2.80% 2.55% 3.01% 4.41% 5.11% 0.95%AKYH767 0.96% 1.08% 0.96% 1.09% 0.81% 0.75% 0.78% 0.40% 0.40% 0.20% 0.12% 0.07% 0.10% 0.19% 0.24% 0.23% 0.00%ChitinophagaceaeChitinophaga 0.01% 0.00% 0.00% 0.00% 0.00% 0.01% 0.01% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Ferruginibacter 0.30% 0.25% 0.24% 0.42% 0.39% 0.33% 0.28% 0.27% 0.22% 0.09% 0.02% 0.02% 0.06% 0.02% 0.06% 0.08% 0.48%Filimonas 0.01% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Flavihumibacter 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.03% 0.00% 0.01% 0.00% 0.00%Flavisolibacter 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Flavitalea 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00%Hydrotalea 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00%Lacibacter 0.27% 0.23% 0.25% 0.22% 0.12% 0.15% 0.14% 0.07% 0.20% 0.11% 0.04% 0.05% 0.02% 0.05% 0.06% 0.05% 0.00%Parafilimonas 0.00% 0.01% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00%Sediminibacterium 0.02% 0.01% 0.02% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Segetibacter 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Taibaiella 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Terrimonas 5.24% 5.12% 5.27% 5.43% 4.24% 5.04% 4.32% 3.22% 2.49% 1.74% 1.79% 1.72% 1.60% 2.01% 3.17% 3.40% 0.00%unclassified 1.58% 1.38% 1.66% 1.46% 1.41% 1.47% 1.30% 1.22% 0.81% 0.57% 0.25% 0.34% 0.35% 0.19% 0.41% 0.74% 0.00%env.OPS_17 0.44% 0.20% 0.22% 0.16% 0.05% 0.06% 0.06% 0.02% 0.00% 0.00% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00% 0.00%KD1‐131 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.01% 0.05% 0.00% 0.02% 0.05% 0.00% 0.03% 0.01% 0.00%KD3‐93 0.01% 0.00% 0.03% 0.02% 0.00% 0.01% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%LiUU‐11‐161 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%NS11‐12_marine_group 0.16% 0.28% 0.20% 0.17% 0.13% 0.24% 0.13% 0.14% 0.05% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%PHOS‐HE51 0.30% 0.25% 0.32% 0.20% 0.19% 0.29% 0.23% 0.13% 0.07% 0.03% 0.00% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00%S15A‐MN91 0.00% 0.01% 0.03% 0.01% 0.01% 0.04% 0.00% 0.01% 0.06% 0.03% 0.01% 0.01% 0.01% 0.03% 0.02% 0.03% 0.00%Saprospiraceae 1.44% 1.30% 1.14% 0.94% 0.84% 1.29% 0.98% 0.83% 0.54% 0.49% 0.17% 0.15% 0.07% 0.07% 0.06% 0.17% 0.00%Sphingobacteriaceae 0.02% 0.01% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.48%unclassified 0.71% 0.78% 0.59% 0.64% 0.52% 0.64% 0.51% 0.35% 0.38% 0.17% 0.09% 0.03% 0.05% 0.11% 0.12% 0.07% 0.00%WCHB1‐69 0.00% 0.07% 0.00% 0.04% 0.03% 0.02% 0.22% 0.05% 0.37% 0.57% 0.45% 0.38% 0.18% 0.34% 0.22% 0.34% 0.00%Chlorobi 4.46% 4.50% 4.38% 4.32% 3.89% 4.17% 4.48% 5.02% 6.90% 8.43% 7.78% 6.78% 5.71% 6.53% 6.55% 7.10% 0.48%Ignavibacteria 4.46% 4.50% 4.38% 4.32% 3.89% 4.17% 4.48% 5.02% 6.90% 8.43% 7.78% 6.78% 5.71% 6.53% 6.55% 7.10% 0.48%Ignavibacteriales 4.46% 4.50% 4.38% 4.32% 3.89% 4.17% 4.48% 5.02% 6.90% 8.43% 7.78% 6.78% 5.71% 6.53% 6.55% 7.10% 0.48%BSV26 3.66% 3.44% 3.36% 3.55% 3.10% 3.46% 3.70% 3.79% 5.09% 5.56% 5.34% 4.32% 3.65% 4.09% 4.50% 5.04% 0.48%IgnavibacteriaceaeIgnavibacterium 0.54% 0.75% 0.74% 0.48% 0.52% 0.41% 0.45% 0.85% 1.37% 2.27% 1.96% 1.85% 1.56% 1.73% 1.45% 1.32% 0.00%IheB3‐7 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00%LD‐RB‐34 0.12% 0.12% 0.17% 0.11% 0.15% 0.15% 0.15% 0.09% 0.17% 0.29% 0.18% 0.16% 0.17% 0.23% 0.29% 0.27% 0.00%PHOS‐HE36 0.06% 0.04% 0.06% 0.10% 0.04% 0.10% 0.08% 0.09% 0.15% 0.13% 0.11% 0.16% 0.16% 0.34% 0.24% 0.25% 0.00%SM1H02 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00%SR‐FBR‐L83 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.03% 0.01% 0.00% 0.01% 0.04% 0.06% 0.01% 0.01% 0.02% 0.03% 0.00%unclassified 0.08% 0.15% 0.05% 0.07% 0.07% 0.07% 0.07% 0.18% 0.11% 0.16% 0.14% 0.23% 0.15% 0.14% 0.06% 0.18% 0.00%Appendix F151Chloroflexi 2.54% 2.38% 2.14% 2.28% 2.11% 2.25% 2.92% 4.22% 7.15% 9.75% 14.56% 15.00% 16.03% 13.55% 12.31% 11.19% 0.48%Anaerolineae 2.54% 2.38% 2.14% 2.28% 2.11% 2.25% 2.92% 4.22% 7.15% 9.75% 14.56% 15.00% 16.03% 13.55% 12.31% 11.19% 0.48%Anaerolineales 2.54% 2.38% 2.14% 2.28% 2.11% 2.25% 2.92% 4.22% 7.15% 9.75% 14.56% 15.00% 16.03% 13.55% 12.31% 11.19% 0.48%AnaerolineaceaeAnaerolinea 0.08% 0.04% 0.15% 0.11% 0.08% 0.08% 0.20% 0.19% 0.24% 0.13% 0.30% 0.26% 0.39% 0.32% 0.30% 0.22% 0.00%Anaerolineaceae_UCG‐001 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00%Leptolinea 0.00% 0.00% 0.00% 0.01% 0.00% 0.01% 0.07% 0.04% 0.06% 0.01% 0.07% 0.15% 0.15% 0.11% 0.09% 0.08% 0.00%Levilinea 0.00% 0.00% 0.01% 0.00% 0.00% 0.01% 0.00% 0.01% 0.02% 0.00% 0.00% 0.03% 0.00% 0.01% 0.00% 0.02% 0.00%Longilinea 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.01% 0.01% 0.00% 0.00% 0.01% 0.00%Ornatilinea 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00%Pelolinea 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.01% 0.06% 0.06% 0.11% 0.09% 0.15% 0.13% 0.11% 0.15% 0.14% 0.00%Thermomarinilinea 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00% 0.00%unclassified 2.46% 2.34% 1.99% 2.16% 2.03% 2.15% 2.63% 3.92% 6.76% 9.50% 14.10% 14.38% 15.35% 13.00% 11.76% 10.73% 0.48%Euryarchaeota 2.75% 2.65% 3.43% 2.31% 2.83% 2.63% 3.11% 3.33% 2.69% 2.98% 3.23% 5.25% 6.78% 5.19% 4.49% 3.23% 0.00%Methanomicrobia 2.75% 2.64% 3.39% 2.29% 2.80% 2.61% 3.10% 3.16% 2.08% 1.95% 1.45% 2.40% 3.12% 3.41% 2.88% 1.97% 0.00%Methanomicrobiales 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.01% 0.52% 0.05% 0.03% 0.05% 0.00%Methanosarcinales 2.75% 2.64% 3.39% 2.29% 2.80% 2.61% 3.10% 3.15% 2.08% 1.95% 1.45% 2.40% 2.60% 3.36% 2.86% 1.92% 0.00%ANME‐2a‐2b 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.03% 0.04% 0.39% 0.42% 0.50% 0.41% 0.07% 0.00%GOM_Arc_ICandidatus_Methanoperedens 2.72% 2.62% 3.39% 2.27% 2.80% 2.59% 3.04% 3.10% 2.06% 1.87% 1.38% 1.84% 1.63% 2.74% 2.31% 1.77% 0.00%Methanosaetaceae 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.02% 0.01% 0.03% 0.01% 0.14% 0.46% 0.10% 0.12% 0.08% 0.00%Methanosarcinaceae 0.01% 0.01% 0.00% 0.00% 0.00% 0.01% 0.03% 0.02% 0.00% 0.03% 0.00% 0.02% 0.07% 0.01% 0.01% 0.00% 0.00%Methermicoccaceae 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00%unclassified 0.02% 0.00% 0.00% 0.02% 0.01% 0.01% 0.03% 0.00% 0.00% 0.00% 0.02% 0.01% 0.00% 0.01% 0.01% 0.00% 0.00%Thermoplasmata 0.00% 0.01% 0.05% 0.02% 0.03% 0.01% 0.01% 0.17% 0.61% 1.03% 1.78% 2.85% 3.66% 1.77% 1.60% 1.26% 0.00%Thermoplasmatales 0.00% 0.01% 0.05% 0.02% 0.03% 0.01% 0.01% 0.17% 0.61% 1.03% 1.78% 2.85% 3.66% 1.77% 1.60% 1.26% 0.00%AMOS1A‐4113‐D04 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.08% 0.08% 0.03% 0.01% 0.00% 0.00%ANT06‐05 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 0.08% 0.11% 0.03% 0.04% 0.02% 0.00%ASC21 0.00% 0.00% 0.02% 0.01% 0.02% 0.00% 0.00% 0.01% 0.12% 0.11% 0.18% 0.51% 0.52% 0.17% 0.20% 0.14% 0.00%CCA47 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 0.09% 0.07% 0.09% 0.03% 0.02% 0.04% 0.04% 0.02% 0.00%Marine_Benthic_Group_D_and_DHVEG‐1 0.00% 0.00% 0.02% 0.01% 0.00% 0.01% 0.01% 0.07% 0.21% 0.77% 1.36% 1.76% 2.63% 1.40% 1.21% 0.97% 0.00%Terrestrial_Miscellaneous_Gp(TMEG) 0.00% 0.01% 0.00% 0.00% 0.01% 0.00% 0.00% 0.07% 0.07% 0.05% 0.06% 0.18% 0.15% 0.06% 0.06% 0.07% 0.00%Thermoplasmatales_Incertae_Sedis 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.02% 0.00% 0.02% 0.00% 0.00%unclassified 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.01% 0.12% 0.03% 0.06% 0.22% 0.13% 0.03% 0.03% 0.03% 0.00%Gemmatimonadetes 6.33% 6.13% 5.24% 5.53% 6.01% 5.98% 5.51% 7.00% 6.20% 5.57% 5.67% 5.40% 4.10% 3.77% 4.93% 5.85% 0.00%Gemmatimonadetes 6.33% 6.13% 5.24% 5.53% 6.01% 5.98% 5.51% 7.00% 6.20% 5.57% 5.67% 5.40% 4.10% 3.77% 4.93% 5.85% 0.00%BD2‐11_terrestrial_group 0.17% 0.17% 0.22% 0.38% 0.43% 0.29% 0.28% 0.32% 0.08% 0.12% 0.08% 0.11% 0.08% 0.08% 0.06% 0.13% 0.00%Gemmatimonadales 6.16% 5.96% 5.02% 5.15% 5.58% 5.68% 5.23% 6.68% 6.12% 5.45% 5.59% 5.29% 4.01% 3.69% 4.87% 5.72% 0.00%GemmatimonadaceaeGemmatimonas 0.06% 0.00% 0.03% 0.01% 0.00% 0.03% 0.01% 0.05% 0.01% 0.00% 0.01% 0.01% 0.00% 0.00% 0.01% 0.01% 0.00%unclassified 6.09% 5.96% 4.99% 5.14% 5.58% 5.65% 5.22% 6.63% 6.11% 5.45% 5.58% 5.28% 4.01% 3.69% 4.86% 5.71% 0.00%Nitrospirae 3.33% 3.03% 4.83% 3.31% 3.40% 3.16% 3.43% 4.72% 6.17% 6.99% 8.06% 7.32% 5.92% 7.55% 6.72% 5.62% 0.48%Nitrospira 3.33% 3.03% 4.83% 3.31% 3.40% 3.16% 3.43% 4.72% 6.17% 6.99% 8.06% 7.32% 5.92% 7.55% 6.72% 5.62% 0.48%Nitrospirales 3.33% 3.03% 4.83% 3.31% 3.40% 3.16% 3.43% 4.72% 6.17% 6.99% 8.06% 7.32% 5.92% 7.55% 6.72% 5.62% 0.48%0319‐6A21 0.27% 0.35% 0.47% 0.33% 0.54% 0.56% 0.44% 0.78% 0.91% 0.80% 0.82% 0.85% 0.58% 0.80% 0.92% 0.61% 0.48%4‐29 0.24% 0.42% 0.86% 0.42% 0.38% 0.29% 0.53% 0.97% 1.47% 1.75% 1.33% 1.10% 0.86% 0.85% 0.64% 0.52% 0.00%FTL22 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00%MIZ17 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.04% 0.01% 0.01% 0.00% 0.00% 0.00%NitrospiraceaeNitrospira 2.38% 2.06% 2.30% 2.24% 2.21% 1.96% 2.08% 1.75% 1.54% 1.43% 1.00% 1.39% 1.39% 1.13% 1.58% 1.41% 0.00%unclassified 0.39% 0.16% 1.17% 0.25% 0.21% 0.25% 0.30% 1.11% 2.09% 2.89% 4.55% 3.59% 2.85% 4.54% 3.33% 2.96% 0.00%Nitrospirales_Incertae_Sedis 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00%Sh765B‐TzT‐35 0.06% 0.04% 0.02% 0.07% 0.07% 0.09% 0.08% 0.10% 0.15% 0.12% 0.35% 0.35% 0.24% 0.21% 0.22% 0.11% 0.00%unclassified 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.02% 0.00% 0.00%wb1‐A12 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00%Planctomycetes 4.83% 4.70% 5.21% 5.25% 5.70% 5.55% 4.57% 4.36% 3.33% 2.93% 2.45% 3.11% 3.18% 2.32% 2.26% 2.94% 0.00%Phycisphaerae 0.39% 0.59% 0.55% 0.41% 0.50% 0.53% 0.48% 0.57% 0.54% 0.44% 0.71% 1.21% 1.22% 0.73% 0.52% 0.62% 0.00%CCM11a 0.09% 0.11% 0.12% 0.04% 0.08% 0.15% 0.09% 0.06% 0.06% 0.01% 0.02% 0.05% 0.07% 0.02% 0.03% 0.05% 0.00%MSBL9 0.01% 0.00% 0.02% 0.00% 0.01% 0.01% 0.02% 0.07% 0.08% 0.13% 0.25% 0.50% 0.61% 0.30% 0.20% 0.25% 0.00%Phycisphaerales 0.17% 0.29% 0.20% 0.18% 0.25% 0.18% 0.22% 0.26% 0.28% 0.24% 0.29% 0.51% 0.32% 0.33% 0.19% 0.26% 0.00%WD2101_soil_group 0.12% 0.19% 0.21% 0.19% 0.17% 0.19% 0.15% 0.18% 0.12% 0.05% 0.15% 0.14% 0.22% 0.08% 0.09% 0.06% 0.00%Planctomycetacia 4.44% 4.11% 4.67% 4.85% 5.20% 5.03% 4.09% 3.79% 2.78% 2.48% 1.74% 1.91% 1.96% 1.59% 1.74% 2.33% 0.00%Planctomycetales 4.44% 4.11% 4.67% 4.85% 5.20% 5.03% 4.09% 3.79% 2.78% 2.48% 1.74% 1.91% 1.96% 1.59% 1.74% 2.33% 0.00%PlanctomycetaceaeBlastopirellula 0.28% 0.27% 0.47% 0.29% 0.36% 0.30% 0.33% 0.21% 0.20% 0.21% 0.25% 0.07% 0.09% 0.07% 0.18% 0.21% 0.00%Candidatus_Anammoximicrobium 0.00% 0.01% 0.02% 0.01% 0.02% 0.02% 0.03% 0.04% 0.02% 0.03% 0.02% 0.01% 0.02% 0.03% 0.04% 0.01% 0.00%Candidatus_Nostocoida 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Gemmata 0.21% 0.24% 0.17% 0.21% 0.16% 0.13% 0.13% 0.08% 0.06% 0.01% 0.01% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00%Isosphaera 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00%Pir1_lineage 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.01% 0.00% 0.01% 0.02% 0.00% 0.02% 0.01% 0.02% 0.00% 0.00%Pir2_lineage 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.02% 0.00% 0.00% 0.00% 0.02% 0.00% 0.02% 0.01% 0.00% 0.00%Pir3_lineage 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.01% 0.00% 0.01% 0.00%Pir4_lineage 0.15% 0.11% 0.15% 0.23% 0.09% 0.07% 0.15% 0.18% 0.14% 0.23% 0.19% 0.35% 0.29% 0.16% 0.21% 0.28% 0.00%Pirellula 0.89% 0.76% 1.02% 0.79% 0.90% 0.97% 0.85% 0.78% 0.58% 0.33% 0.12% 0.26% 0.19% 0.28% 0.16% 0.26% 0.00%Planctomyces 1.23% 0.92% 1.05% 1.23% 1.36% 1.31% 0.93% 1.02% 0.67% 0.43% 0.27% 0.40% 0.42% 0.23% 0.30% 0.37% 0.00%Rhodopirellula 0.10% 0.05% 0.08% 0.13% 0.13% 0.16% 0.13% 0.11% 0.08% 0.08% 0.08% 0.04% 0.12% 0.03% 0.08% 0.13% 0.00%Schlesneria 0.19% 0.21% 0.11% 0.14% 0.16% 0.11% 0.18% 0.12% 0.11% 0.01% 0.01% 0.05% 0.02% 0.01% 0.00% 0.02% 0.00%Singulisphaera 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%unclassified 1.30% 1.39% 1.45% 1.71% 1.90% 1.79% 1.26% 1.15% 0.90% 1.12% 0.78% 0.69% 0.76% 0.71% 0.75% 1.01% 0.00%Zavarzinella 0.09% 0.13% 0.16% 0.11% 0.11% 0.16% 0.08% 0.07% 0.00% 0.01% 0.00% 0.02% 0.00% 0.02% 0.00% 0.03% 0.00%Appendix F152Proteobacteria 32.70% 33.74% 33.89% 34.50% 33.64% 32.51% 36.29% 36.05% 39.96% 38.41% 39.05% 39.69% 41.68% 44.73% 43.94% 42.49% 93.33%Alphaproteobacteria 7.38% 7.02% 7.99% 8.66% 7.54% 8.01% 8.69% 8.21% 7.82% 7.72% 7.18% 6.48% 7.82% 6.41% 7.41% 8.03% 48.57%Caulobacterales 0.32% 0.25% 0.25% 0.30% 0.24% 0.24% 0.36% 0.22% 0.29% 0.31% 0.25% 0.18% 0.26% 0.18% 0.24% 0.26% 0.00%Caulobacteraceae 0.06% 0.04% 0.02% 0.05% 0.07% 0.05% 0.05% 0.03% 0.05% 0.13% 0.09% 0.05% 0.10% 0.07% 0.09% 0.09% 0.00%Hyphomonadaceae 0.26% 0.21% 0.23% 0.25% 0.17% 0.19% 0.31% 0.19% 0.25% 0.17% 0.16% 0.12% 0.16% 0.11% 0.15% 0.18% 0.00%Rhizobiales 5.62% 5.52% 6.44% 7.08% 6.06% 6.60% 6.84% 6.56% 5.86% 6.16% 5.86% 5.21% 6.08% 5.02% 5.74% 6.20% 41.90%1174‐901‐12 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.48%A0839 0.60% 0.33% 0.47% 0.57% 0.36% 0.46% 0.46% 0.41% 0.26% 0.19% 0.26% 0.15% 0.14% 0.14% 0.14% 0.26% 0.00%Beijerinckiaceae 0.01% 0.00% 0.00% 0.10% 0.05% 0.01% 0.05% 0.05% 0.01% 0.00% 0.04% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Bradyrhizobiaceae 0.08% 0.00% 0.04% 0.04% 0.03% 0.01% 0.05% 0.04% 0.01% 0.01% 0.02% 0.04% 0.08% 0.04% 0.02% 0.06% 1.43%D05‐2 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00%DUNssu044 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00%DUNssu371 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00%HyphomicrobiaceaeDevosia 0.02% 0.00% 0.02% 0.01% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.01% 0.03% 0.00% 0.00% 0.00% 0.00%Hyphomicrobium 0.80% 0.82% 0.84% 1.14% 0.96% 0.97% 1.05% 0.93% 0.80% 1.00% 0.96% 0.87% 1.15% 0.95% 1.05% 0.95% 0.00%Pedomicrobium 0.19% 0.19% 0.30% 0.25% 0.24% 0.30% 0.30% 0.26% 0.37% 0.35% 0.19% 0.36% 0.36% 0.12% 0.18% 0.19% 0.00%Prosthecomicrobium 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00%Rhodomicrobium 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00%Rhodoplanes 0.02% 0.01% 0.02% 0.01% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.01% 0.01% 0.00% 0.01% 0.00% 0.00% 0.00%unclassified 0.12% 0.23% 0.13% 0.16% 0.15% 0.16% 0.16% 0.22% 0.15% 0.23% 0.16% 0.12% 0.14% 0.13% 0.28% 0.18% 0.00%JG35‐K1‐AG5 0.00% 0.00% 0.00% 0.00% 0.01% 0.01% 0.02% 0.00% 0.00% 0.00% 0.01% 0.01% 0.04% 0.00% 0.00% 0.01% 0.00%KF‐JG30‐B3 0.26% 0.43% 0.41% 0.42% 0.36% 0.40% 0.53% 0.39% 0.46% 0.56% 0.35% 0.19% 0.29% 0.29% 0.33% 0.44% 0.48%Methylobacteriaceae 0.03% 0.01% 0.02% 0.00% 0.01% 0.03% 0.05% 0.02% 0.13% 0.11% 0.15% 0.12% 0.14% 0.14% 0.06% 0.09% 37.62%Methylocystaceae 0.00% 0.00% 0.00% 0.01% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 0.01% 0.00% 0.00% 0.00% 0.00%MNG7 2.18% 2.21% 2.75% 2.75% 2.33% 2.58% 2.65% 3.00% 2.11% 2.58% 2.57% 2.02% 2.17% 2.27% 2.60% 2.67% 0.00%Phyllobacteriaceae 0.08% 0.03% 0.07% 0.05% 0.03% 0.05% 0.09% 0.04% 0.08% 0.03% 0.09% 0.05% 0.08% 0.06% 0.10% 0.11% 0.00%Rhizobiaceae 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 1.90%Rhizobiales_Incertae_Sedis 0.06% 0.04% 0.03% 0.03% 0.04% 0.05% 0.03% 0.01% 0.01% 0.00% 0.04% 0.05% 0.04% 0.02% 0.02% 0.01% 0.00%Rhodobiaceae 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 0.01% 0.00% 0.00% 0.00% 0.00%unclassified 0.84% 0.75% 0.83% 0.94% 0.79% 0.82% 0.76% 0.61% 0.73% 0.52% 0.61% 0.55% 0.83% 0.43% 0.47% 0.82% 0.00%Xanthobacteraceae 0.33% 0.47% 0.50% 0.58% 0.72% 0.76% 0.61% 0.59% 0.71% 0.59% 0.42% 0.60% 0.59% 0.41% 0.51% 0.41% 0.00%Rhodobacterales 0.14% 0.05% 0.08% 0.07% 0.06% 0.07% 0.10% 0.14% 0.17% 0.20% 0.17% 0.23% 0.11% 0.17% 0.12% 0.10% 0.95%Rhodospirillales 0.70% 0.75% 0.87% 0.80% 0.84% 0.71% 0.88% 0.82% 1.01% 0.76% 0.76% 0.69% 0.96% 0.89% 1.10% 1.14% 1.43%Sphingomonadales 0.60% 0.44% 0.36% 0.40% 0.34% 0.40% 0.51% 0.48% 0.48% 0.29% 0.15% 0.18% 0.42% 0.14% 0.21% 0.32% 4.29%Betaproteobacteria 10.36% 11.03% 10.22% 10.95% 10.55% 10.24% 11.24% 10.35% 10.77% 9.94% 7.77% 5.84% 5.71% 6.42% 7.66% 7.80% 6.67%B1‐7BS 0.57% 0.42% 0.54% 0.55% 0.62% 0.48% 0.46% 0.61% 0.71% 0.60% 0.56% 0.47% 0.34% 0.45% 0.37% 0.40% 0.00%Burkholderiales 4.21% 4.02% 4.12% 3.68% 3.63% 3.74% 4.24% 4.14% 4.42% 3.98% 3.33% 2.40% 2.44% 2.98% 3.68% 3.98% 5.24%Alcaligenaceae 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.02% 0.00% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00% 0.00%Burkholderiaceae 0.03% 0.00% 0.00% 0.00% 0.01% 0.01% 0.02% 0.02% 0.04% 0.01% 0.03% 0.02% 0.02% 0.00% 0.02% 0.00% 0.00%CM1G08 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Comamonadaceae 4.18% 3.99% 3.99% 3.63% 3.60% 3.69% 4.21% 4.11% 4.34% 3.97% 3.28% 2.33% 2.38% 2.96% 3.66% 3.93% 5.24%Oxalobacteraceae 0.00% 0.01% 0.00% 0.00% 0.00% 0.01% 0.01% 0.01% 0.00% 0.00% 0.00% 0.00% 0.02% 0.01% 0.00% 0.00% 0.00%unclassified 0.00% 0.01% 0.14% 0.04% 0.01% 0.05% 0.01% 0.00% 0.02% 0.00% 0.02% 0.02% 0.02% 0.01% 0.01% 0.05% 0.00%Hydrogenophilales 0.11% 0.09% 0.10% 0.04% 0.12% 0.06% 0.07% 0.11% 0.25% 0.41% 0.20% 0.23% 0.15% 0.24% 0.27% 0.18% 0.00%Methylophilales 0.47% 0.55% 0.58% 0.40% 0.45% 0.34% 0.47% 0.43% 0.44% 0.39% 0.23% 0.28% 0.39% 0.26% 0.43% 0.53% 0.00%Nitrosomonadales 1.73% 1.78% 0.60% 1.87% 1.85% 2.04% 1.92% 1.94% 1.79% 1.60% 1.31% 0.91% 0.88% 1.00% 1.17% 1.10% 0.00%Gallionellaceae 0.08% 0.07% 0.30% 0.07% 0.08% 0.05% 0.06% 0.16% 0.25% 0.16% 0.08% 0.08% 0.04% 0.12% 0.02% 0.08% 0.00%Nitrosomonadaceae 1.66% 1.71% 0.30% 1.80% 1.76% 1.99% 1.86% 1.78% 1.54% 1.44% 1.23% 0.82% 0.82% 0.87% 1.10% 1.01% 0.00%unclassified 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.02% 0.00% 0.06% 0.02% 0.00%Rhodocyclales 0.18% 0.24% 0.14% 0.15% 0.08% 0.09% 0.35% 0.11% 0.13% 0.09% 0.06% 0.08% 0.08% 0.06% 0.15% 0.04% 1.43%SC‐I‐84 0.23% 0.42% 0.37% 0.33% 0.33% 0.39% 0.41% 0.24% 0.31% 0.31% 0.19% 0.35% 0.26% 0.27% 0.37% 0.30% 0.00%TRA3‐20 2.86% 3.52% 3.77% 3.94% 3.48% 3.09% 3.32% 2.76% 2.72% 2.55% 1.90% 1.13% 1.17% 1.16% 1.22% 1.28% 0.00%Deltaproteobacteria 8.63% 9.13% 9.31% 8.98% 8.97% 8.26% 9.47% 10.70% 11.92% 12.77% 15.88% 19.45% 19.86% 19.84% 17.84% 16.87% 0.48%43F‐1404R 1.11% 1.02% 0.90% 1.03% 1.21% 1.24% 1.26% 2.05% 1.46% 1.87% 2.32% 1.88% 1.22% 2.14% 1.85% 2.03% 0.00%Bdellovibrionales 0.26% 0.17% 0.24% 0.28% 0.25% 0.19% 0.24% 0.14% 0.15% 0.11% 0.10% 0.25% 0.12% 0.13% 0.28% 0.10% 0.00%Deltaproteobacteria_Incertae_Sedis 0.02% 0.00% 0.02% 0.04% 0.03% 0.03% 0.04% 0.04% 0.06% 0.05% 0.15% 0.17% 0.49% 0.22% 0.26% 0.20% 0.00%Desulfarculales 0.05% 0.05% 0.06% 0.03% 0.04% 0.03% 0.03% 0.13% 0.21% 0.27% 0.47% 0.76% 0.83% 0.73% 0.92% 0.70% 0.00%DesulfarculaceaeDesulfarculus 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00%Desulfatiglans 0.05% 0.05% 0.06% 0.03% 0.04% 0.03% 0.03% 0.12% 0.21% 0.27% 0.47% 0.76% 0.83% 0.73% 0.91% 0.70% 0.00%Desulfobacterales 0.38% 0.39% 0.38% 0.35% 0.31% 0.40% 0.60% 0.63% 0.81% 1.02% 1.92% 4.65% 4.88% 4.91% 3.50% 3.19% 0.00%Desulfobacteraceae 0.04% 0.12% 0.07% 0.04% 0.06% 0.07% 0.19% 0.16% 0.21% 0.32% 0.81% 0.73% 0.62% 0.30% 0.31% 0.42% 0.00%DesulfobulbaceaeDesulfobulbus 0.02% 0.00% 0.00% 0.01% 0.03% 0.01% 0.04% 0.00% 0.02% 0.03% 0.05% 0.03% 0.06% 0.05% 0.02% 0.00% 0.00%Desulfocapsa 0.01% 0.01% 0.03% 0.01% 0.00% 0.01% 0.04% 0.00% 0.01% 0.00% 0.01% 0.02% 0.02% 0.02% 0.00% 0.00% 0.00%Desulfurivibrio 0.00% 0.00% 0.00% 0.00% 0.01% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.01% 0.00% 0.00% 0.00%MSBL7 0.15% 0.07% 0.04% 0.04% 0.03% 0.07% 0.06% 0.14% 0.25% 0.31% 0.59% 2.86% 3.23% 2.93% 2.43% 2.29% 0.00%unclassified 0.00% 0.00% 0.03% 0.00% 0.00% 0.00% 0.04% 0.03% 0.14% 0.05% 0.04% 0.04% 0.08% 0.01% 0.03% 0.02% 0.00%Nitrospinaceae 0.15% 0.19% 0.21% 0.25% 0.18% 0.23% 0.22% 0.25% 0.11% 0.16% 0.07% 0.08% 0.07% 0.05% 0.02% 0.05% 0.00%unclassified 0.00% 0.00% 0.00% 0.00% 0.01% 0.01% 0.00% 0.05% 0.07% 0.15% 0.37% 0.88% 0.80% 1.55% 0.69% 0.42% 0.00%Desulfurellales 0.43% 0.39% 0.39% 0.41% 0.27% 0.27% 0.31% 0.30% 0.24% 0.36% 0.22% 0.29% 0.17% 0.12% 0.28% 0.35% 0.00%Desulfuromonadales 0.23% 0.42% 0.68% 0.15% 0.64% 0.20% 0.42% 0.13% 0.19% 0.17% 0.23% 0.26% 0.26% 0.14% 0.11% 0.22% 0.48%GR‐WP33‐30 2.27% 3.16% 2.49% 2.90% 2.44% 2.23% 2.15% 2.18% 3.05% 2.84% 2.68% 2.36% 1.97% 1.97% 1.94% 2.17% 0.00%Myxococcales 2.46% 2.28% 2.75% 2.50% 2.33% 2.28% 2.74% 2.27% 2.11% 1.47% 1.75% 2.17% 2.30% 2.21% 2.30% 2.41% 0.00%27F‐1492R 0.09% 0.09% 0.02% 0.07% 0.03% 0.02% 0.07% 0.03% 0.04% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00%Amb‐16S‐1034 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%bacteriap25 0.00% 0.00% 0.01% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%BIrii41 0.04% 0.07% 0.08% 0.05% 0.05% 0.03% 0.13% 0.13% 0.05% 0.03% 0.03% 0.03% 0.03% 0.03% 0.03% 0.07% 0.00%Blfdi19 0.04% 0.03% 0.04% 0.04% 0.02% 0.02% 0.02% 0.01% 0.04% 0.00% 0.01% 0.00% 0.01% 0.04% 0.02% 0.01% 0.00%CystobacteraceaeAnaeromyxobacter 0.62% 0.64% 0.90% 0.75% 0.73% 0.71% 1.05% 0.84% 0.63% 0.64% 0.90% 1.12% 1.25% 1.22% 1.47% 1.63% 0.00%unclassified 0.17% 0.16% 0.11% 0.19% 0.13% 0.15% 0.06% 0.13% 0.05% 0.03% 0.03% 0.03% 0.03% 0.05% 0.02% 0.03% 0.00%Eel‐36e1D6 0.00% 0.00% 0.01% 0.00% 0.01% 0.01% 0.00% 0.00% 0.06% 0.01% 0.02% 0.04% 0.02% 0.02% 0.02% 0.04% 0.00%Haliangiaceae 0.45% 0.55% 0.56% 0.57% 0.57% 0.55% 0.57% 0.37% 0.32% 0.17% 0.14% 0.13% 0.20% 0.15% 0.09% 0.19% 0.00%KD3‐10 0.00% 0.01% 0.03% 0.01% 0.00% 0.01% 0.02% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%MidBa8 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.03% 0.05% 0.03% 0.09% 0.12% 0.15% 0.08% 0.10% 0.06% 0.00%mle1‐27 0.02% 0.01% 0.02% 0.01% 0.02% 0.00% 0.01% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.01% 0.00%MSB‐4B10 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.02% 0.00% 0.03% 0.01% 0.02% 0.00% 0.03% 0.00% 0.00% 0.00%Myxococcaceae 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.02% 0.00% 0.00% 0.00% 0.00% 0.00%Appendix F153Nannocystaceae 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%P3OB‐42 0.28% 0.15% 0.27% 0.08% 0.13% 0.07% 0.15% 0.09% 0.19% 0.08% 0.02% 0.02% 0.00% 0.05% 0.04% 0.02% 0.00%Phaselicystidaceae 0.02% 0.00% 0.04% 0.04% 0.01% 0.03% 0.01% 0.01% 0.04% 0.00% 0.00% 0.02% 0.02% 0.00% 0.01% 0.01% 0.00%Polyangiaceae 0.15% 0.13% 0.11% 0.13% 0.14% 0.10% 0.06% 0.08% 0.17% 0.04% 0.04% 0.04% 0.05% 0.03% 0.02% 0.02% 0.00%PS‐B29 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.01% 0.04% 0.01% 0.02% 0.03% 0.03% 0.02% 0.00%Sandaracinaceae 0.16% 0.17% 0.08% 0.14% 0.16% 0.09% 0.13% 0.12% 0.05% 0.07% 0.04% 0.02% 0.00% 0.03% 0.02% 0.01% 0.00%UASB‐TL25 0.01% 0.01% 0.01% 0.00% 0.03% 0.03% 0.01% 0.04% 0.04% 0.03% 0.06% 0.10% 0.01% 0.12% 0.05% 0.04% 0.00%unclassified 0.39% 0.24% 0.47% 0.40% 0.30% 0.46% 0.44% 0.35% 0.40% 0.31% 0.32% 0.42% 0.26% 0.32% 0.34% 0.22% 0.00%VHS‐B3‐70 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 0.00% 0.01% 0.04% 0.25% 0.03% 0.05% 0.02% 0.00%VHS‐B4‐70 0.01% 0.00% 0.01% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Sh765B‐TzT‐29 0.97% 0.90% 0.89% 0.97% 1.07% 1.04% 1.16% 1.26% 1.06% 1.00% 1.34% 0.83% 0.84% 0.98% 1.13% 1.04% 0.00%Sva0485 0.28% 0.24% 0.24% 0.17% 0.21% 0.20% 0.26% 0.66% 1.17% 1.87% 2.50% 2.59% 3.01% 3.09% 2.73% 2.09% 0.00%Syntrophobacterales 0.18% 0.12% 0.27% 0.15% 0.16% 0.15% 0.24% 0.92% 1.40% 1.74% 2.19% 3.24% 3.77% 3.19% 2.54% 2.37% 0.00%SyntrophaceaeDesulfobacca 0.08% 0.03% 0.07% 0.03% 0.03% 0.02% 0.06% 0.07% 0.12% 0.19% 0.29% 0.38% 0.56% 0.42% 0.37% 0.31% 0.00%Desulfomonile 0.00% 0.00% 0.01% 0.01% 0.01% 0.01% 0.00% 0.03% 0.05% 0.03% 0.04% 0.03% 0.08% 0.06% 0.02% 0.00% 0.00%Smithella 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.01% 0.00% 0.00% 0.05% 0.23% 0.61% 0.20% 0.21% 0.09% 0.00%Syntrophus 0.10% 0.07% 0.11% 0.10% 0.10% 0.11% 0.16% 0.75% 1.19% 1.42% 1.49% 1.85% 1.36% 1.67% 1.39% 1.39% 0.00%unclassified 0.00% 0.00% 0.02% 0.00% 0.00% 0.01% 0.01% 0.00% 0.01% 0.00% 0.09% 0.34% 0.52% 0.36% 0.22% 0.24% 0.00%Syntrophobacteraceae 0.01% 0.03% 0.05% 0.01% 0.02% 0.02% 0.02% 0.06% 0.04% 0.11% 0.24% 0.41% 0.66% 0.49% 0.33% 0.34% 0.00%unclassified 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Epsilonproteobacteria 0.02% 0.01% 0.00% 0.00% 0.01% 0.01% 0.02% 0.00% 0.01% 0.01% 0.05% 0.10% 0.58% 1.26% 0.49% 0.15% 0.95%Campylobacterales 0.02% 0.01% 0.00% 0.00% 0.01% 0.01% 0.02% 0.00% 0.01% 0.01% 0.05% 0.10% 0.58% 1.26% 0.49% 0.15% 0.95%CampylobacteraceaeArcobacter 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.01% 0.01% 0.01% 0.00% 0.00% 0.02% 0.00% 0.95%HelicobacteraceaeSulfuricurvum 0.01% 0.01% 0.00% 0.00% 0.01% 0.00% 0.01% 0.00% 0.01% 0.00% 0.04% 0.09% 0.57% 1.26% 0.44% 0.15% 0.00%Sulfurimonas 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.03% 0.00% 0.00%Sulfurovum 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Gammaproteobacteria 6.31% 6.55% 6.37% 5.92% 6.58% 6.00% 6.88% 6.79% 9.45% 7.97% 8.17% 7.81% 7.71% 10.81% 10.55% 9.64% 36.67%Chromatiales 1.05% 0.98% 1.06% 0.96% 0.66% 0.82% 0.98% 1.09% 1.72% 1.74% 1.21% 0.97% 0.93% 1.44% 1.28% 0.94% 0.00%Chromatiaceae 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%EctothiorhodospiraceaeAcidiferrobacter 1.05% 0.98% 1.06% 0.96% 0.66% 0.82% 0.98% 1.09% 1.71% 1.74% 1.21% 0.97% 0.93% 1.44% 1.28% 0.94% 0.00%Methylococcales 5.17% 5.37% 5.15% 4.81% 5.66% 5.01% 5.59% 5.53% 7.45% 5.93% 6.61% 6.53% 6.30% 9.02% 9.00% 8.55% 0.00%CrenotrichaceaeCrenothrix 1.39% 1.35% 1.68% 1.26% 1.63% 1.38% 1.78% 1.70% 2.49% 1.67% 1.81% 1.76% 1.84% 2.44% 2.87% 2.93% 0.00%MethylococcaceaeMethylobacter 0.46% 0.84% 0.76% 0.74% 0.75% 0.76% 0.83% 0.88% 1.27% 0.91% 1.58% 1.82% 1.46% 1.77% 1.66% 1.73% 0.00%Methylocaldum 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Methylococcus 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Methyloglobulus 0.49% 0.67% 0.95% 0.73% 1.33% 0.99% 1.15% 1.13% 1.27% 1.03% 0.89% 0.89% 1.18% 1.68% 1.79% 1.37% 0.00%Methylomonas 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Methylosarcina 0.00% 0.00% 0.00% 0.00% 0.01% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%unclassified 0.01% 0.01% 0.02% 0.10% 0.04% 0.02% 0.02% 0.04% 0.06% 0.03% 0.05% 0.04% 0.01% 0.05% 0.05% 0.04% 0.00%pLW‐20 0.04% 0.03% 0.02% 0.01% 0.01% 0.01% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%unclassified 2.77% 2.46% 1.72% 1.98% 1.88% 1.84% 1.81% 1.76% 2.35% 2.30% 2.28% 2.02% 1.82% 3.08% 2.63% 2.49% 0.00%Oceanospirillales 0.01% 0.05% 0.04% 0.01% 0.09% 0.03% 0.03% 0.07% 0.14% 0.24% 0.25% 0.16% 0.29% 0.27% 0.20% 0.03% 33.81%Alcanivoracaceae 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00%HalomonadaceaeHalomonas 0.01% 0.04% 0.04% 0.01% 0.08% 0.03% 0.03% 0.07% 0.13% 0.24% 0.25% 0.12% 0.29% 0.27% 0.19% 0.03% 31.90%unclassified 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00%Oleiphilaceae 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%OM182_clade 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%SUP05_cluster 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.04% 0.00% 0.00% 0.01% 0.00% 1.90%unclassified 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Xanthomonadales 0.08% 0.15% 0.13% 0.13% 0.18% 0.14% 0.27% 0.11% 0.13% 0.07% 0.09% 0.15% 0.19% 0.08% 0.06% 0.11% 2.86%unclassified 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.02% 0.03% 0.02% 0.00% 0.01% 0.02% 0.00%Xanthomonadaceae 0.01% 0.01% 0.05% 0.01% 0.05% 0.06% 0.07% 0.04% 0.06% 0.04% 0.05% 0.05% 0.07% 0.03% 0.00% 0.06% 2.86%Xanthomonadales_Incertae_Sedis 0.06% 0.13% 0.08% 0.13% 0.13% 0.08% 0.20% 0.07% 0.07% 0.01% 0.03% 0.07% 0.10% 0.05% 0.06% 0.03% 0.00%Spirochaetae 0.06% 0.11% 0.15% 0.10% 0.13% 0.10% 0.28% 0.36% 0.60% 0.73% 1.26% 1.60% 1.84% 1.82% 0.87% 1.16% 0.00%Spirochaetes 0.06% 0.11% 0.15% 0.10% 0.13% 0.10% 0.28% 0.36% 0.60% 0.73% 1.26% 1.60% 1.84% 1.82% 0.87% 1.16% 0.00%Spirochaetales 0.06% 0.11% 0.15% 0.10% 0.13% 0.10% 0.28% 0.36% 0.60% 0.73% 1.26% 1.60% 1.84% 1.82% 0.87% 1.16% 0.00%Brevinemataceae 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00%Leptospiraceae 0.00% 0.00% 0.06% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.02% 0.02% 0.07% 0.03% 0.03% 0.02% 0.00%PL‐11B10 0.00% 0.03% 0.02% 0.01% 0.01% 0.02% 0.04% 0.04% 0.04% 0.01% 0.00% 0.01% 0.02% 0.06% 0.00% 0.01% 0.00%Spirochaetaceae 0.06% 0.08% 0.08% 0.08% 0.11% 0.08% 0.21% 0.31% 0.57% 0.72% 1.24% 1.57% 1.73% 1.72% 0.84% 1.13% 0.00%unclassified 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00%V2072‐189E03 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00%Verrucomicrobia 5.59% 4.73% 4.70% 4.98% 4.03% 4.12% 3.58% 2.71% 1.93% 1.26% 0.46% 0.42% 0.44% 0.47% 0.53% 0.80% 0.48%Spartobacteria 4.56% 3.90% 3.57% 3.81% 3.19% 3.00% 2.77% 2.13% 1.57% 1.04% 0.38% 0.36% 0.37% 0.34% 0.46% 0.69% 0.00%Chthoniobacterales 4.56% 3.90% 3.57% 3.81% 3.19% 3.00% 2.77% 2.13% 1.57% 1.04% 0.38% 0.36% 0.37% 0.34% 0.46% 0.69% 0.00%01D2Z36 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%ChthoniobacteraceaeChthoniobacter 4.35% 3.62% 3.43% 3.68% 3.02% 2.91% 2.72% 2.04% 1.44% 0.92% 0.34% 0.32% 0.34% 0.30% 0.43% 0.64% 0.00%DA101_soil_group 0.18% 0.28% 0.14% 0.12% 0.16% 0.09% 0.05% 0.07% 0.11% 0.12% 0.01% 0.04% 0.02% 0.01% 0.02% 0.05% 0.00%FukuN18_freshwater_group 0.00% 0.00% 0.00% 0.01% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00%LD29 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%unclassified 0.02% 0.00% 0.00% 0.01% 0.00% 0.01% 0.00% 0.01% 0.01% 0.00% 0.02% 0.00% 0.02% 0.03% 0.01% 0.00% 0.00%Xiphinematobacteraceae 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.01% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00%Verrucomicrobiae 1.03% 0.83% 1.14% 1.17% 0.84% 1.11% 0.81% 0.58% 0.37% 0.21% 0.09% 0.06% 0.07% 0.13% 0.06% 0.11% 0.48%Verrucomicrobiales 1.03% 0.83% 1.14% 1.17% 0.84% 1.11% 0.81% 0.58% 0.37% 0.21% 0.09% 0.06% 0.07% 0.13% 0.06% 0.11% 0.48%DEV007 0.01% 0.01% 0.04% 0.01% 0.04% 0.05% 0.02% 0.03% 0.04% 0.00% 0.02% 0.00% 0.01% 0.02% 0.00% 0.01% 0.00%P._palm_C_85 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.00% 0.02% 0.00% 0.00% 0.00% 0.00% 0.00%unclassified 0.91% 0.72% 1.02% 1.09% 0.68% 1.03% 0.68% 0.49% 0.29% 0.17% 0.03% 0.01% 0.05% 0.07% 0.04% 0.09% 0.00%Verrucomicrobiaceae 0.11% 0.09% 0.08% 0.07% 0.12% 0.03% 0.11% 0.07% 0.04% 0.04% 0.04% 0.03% 0.02% 0.04% 0.03% 0.02% 0.48%Grand Total 100.00% 100.00% 100.00% 100.00% 100.00% 100.00% 100.00% 100.00% 100.00% 100.00% 100.00% 100.00% 100.00% 100.00% 100.00% 100.00% 100.00%Appendix F154 2Th Degrees75706560555045403530252015105Sqrt(Counts)200150100500MZ_7.raw_1 Quartz low 50.38 %Albite low 16.60 %Albite low, calcian 13.33 %Actinolite 2.38 %Orthoclase 4.38 %Ankerite 0.63 %Illite/Muscovite 2M1 3.74 %Illite/Muscovite 1M 3.20 %Chamosite 1MIIb 4.82 %Lizardite 1T ? 0.53 %Appendix G155 2Th Degrees75706560555045403530252015105Sqrt(Counts)200150100500MZ_9.raw_1 Quartz low 48.61 %Albite low 18.07 %Albite low, calcian 12.72 %Actinolite 2.66 %Orthoclase 4.41 %Ankerite 0.64 %Illite/Muscovite 2M1 3.70 %Illite/Muscovite 1M 3.65 %Chamosite 1MIIb 4.93 %Lizardite 1T ? 0.61 %Appendix G156 2Th Degrees75706560555045403530252015105Sqrt(Counts)200150100500MZ_13.raw_1 Quartz low 50.92 %Albite low 17.57 %Albite low, calcian 12.67 %Actinolite 1.96 %Orthoclase 4.64 %Ankerite 0.51 %Illite/Muscovite 2M1 3.44 %Illite/Muscovite 1M 3.12 %Chamosite 1MIIb 4.53 %Lizardite 1T ? 0.63 %Appendix G157  Appendix H158                             Appendix H159
https://iiif.library.ubc.ca/presentation/dsp.24.1-0314121/manifest