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Supplementary Information
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Halomonas sulfidaeris-dominated Communities Inhabit 1.8km-deep Saline Sandstone
Reservoirs
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Yiran Dong1,2, Charu Gupta Kumar1, Nicholas Chia1,3,4, Pan-Jun Kim1,5, Philip A. Miller1,2,6,
Nathan D. Price1,3,7, Isaac K.O. Cann1,8,9, Theodore M. Flynn1,2,10, Robert A. Sanford1,2, Ivan
G. Krapac11, Randall A. Locke II11, Pei-Ying Hong12, Hideyuki Tamaki12,13, Wen-Tso Liu1,12,
Alvaro G. Hernandez14, Chris L. Wright14, Mark A. Mikel14, Jared L. Walker15, Mayandi
Sivaguru1, Glenn Fried1, Anthony C. Yannarell16, Roderick I. Mackie1, Bruce W.
Fouke1,2,3,9,11*
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Energy Biosciences Institute, Institute for Genomic Biology, University of Illinois UrbanaChampaign, 1206 W. Gregory Drive, Urbana, Illinois 61801 USA
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Department of Geology, University of Illinois Urbana-Champaign, 1301 W. Green Street,
Urbana, Illinois 61801 USA
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Institute for Systems Biology, 401 Terry Avenue North, Seattle, Washington 98109 USA
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Department of Surgery, Mayo Clinic, 200 First Street S.W., Rochester, Minnesota 55905,
USA
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Asia Pacific Center for Theoretical Physics, San 31 Hyoja-dong, Nam-gu, Pohang,
Gyeongbuk, 790-784 Korea
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Shell Oil Company, 150 N. Dairy Ashford Drive, Houston, Texas 77079 USA
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Chemical and Biomolecular Engineering, University of Illinois Urbana-Champaign, 600 S.
Mathews Avenue, Urbana, Illinois 61801 USA
Department of Animal Sciences, University of Illinois Urbana-Champaign, 1207 W.
Gregory Drive, Urbana, Illinois 61801 USA
Department of Microbiology, University of Illinois Urbana-Champaign, 601 S. Goodwin
Avenue, Urbana, Illinois 61801 USA
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Biosciences Division, Argonne National Laboratory, 9700 South Cass Avenue, Argonne,
Illinois 60439 USA
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Illinois State Geological Survey, Prairie Research Institute, University of Illinois UrbanaChampaign, 615 E. Peabody Drive, Urbana, Illinois 61801 USA
Civil and Environmental Engineering, University of Illinois Urbana-Champaign, 205 N.
Mathews Avenue, Urbana, Illinois 61801 USA
Bioproduction Research Institute, National Institute of Advanced Industrial Science and
Technology (AIST), Central 6, Higashi 1-1-1, Tsukuba, Ibaraki, 305-8566 Japan
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Roy J. Carver Biotechnology Center, University of Illinois Urbana-Champaign, 2613
Institute for Genomic Biology, 1206 W. Gregory Drive, Urbana, Illinois 61801 USA
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Schlumberger Carbon Services, 14090 SW Freeway, Suite 300, Sugar Land, Texas 77478
USA
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Department of Natural Resources and Environmental Sciences, University of Illinois at
Urbana-Champaign, 1102 South Goodwin Avenue, Urbana, Illinois 61801 USA
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Corresponding Author:
Bruce W. Fouke, Institute for Genomic Biology, University of Illinois Urbana-Champaign,
1206 W. Gregory Drive, Urbana, Illinois 61801 USA (E-mail: fouke@illinois.edu)
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Index
Geological and Hydrological Setting ...................................................................................... 1
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Drilling and Subsurface Reservoir Analyses
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Well Drilling ............................................................................................................................ 5
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Rock Core Collection .............................................................................................................. 6
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Schlumberger Quicksilver Probe ............................................................................................. 6
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Wire-Line Log Electrical Data Collection............................................................................... 8
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Drillstem Test (DST) ............................................................................................................... 8
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Collection of Quicksilver Probe MDT Water Samples ...................................................... 10
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Gas collection ........................................................................................................................ 11
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Centrifugation of water samples ............................................................................................ 11
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Geochemical Analyses .......................................................................................................... 12
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Strontium Isotopic Analysis .................................................................................................. 13
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Microscopic Analyses
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Lithology analysis of thin sections ........................................................................................ 14
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Fluorescence In-Situ Hybridization (FISH) for cell enumeration ......................................... 15
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Cultivation of Organisms ...................................................................................................... 17
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Microbial Community Analyses
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DNA extraction ..................................................................................................................... 18
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Development of 16S rRNA Clone library ............................................................................. 18
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Terminal Restriction Fragment Length Polymorphism (T-RFLP) ........................................ 20
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Metagenomic Analyses
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V1-V3 16S rRNA Hypervariable Region Pyrosequencing ................................................... 20
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Halomonas sulfidaeris strain Esulfide1 as a Reference Genome for D5872 Metagenome... 21
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454 pyrosequencing of D5872 Metagenome and H. sulfidaeris Esulfide1 genome ............. 22
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Metagenomic and Genomic Analyses ................................................................................... 22
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Figures and Tables
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Figure S1. Fluorescence hybridization of the D5872 microbial consortium ......................... 25
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Figure S2. T-RFLP analysis of D5872 microbial community and drilling mud ................... 26
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Figure S3. Phylogenetic relationship of Halomonas species in D5872 microbial community
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versus drilling mud ............................................................................................................... 27
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Figure S4. Rarefaction of D5872 and DDM microbial community richness ........................ 28
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Figure S5. Distribution of GC-content with length for D5872 Metagenome contigs ........... 29
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Figure S6. Taxonomic assignment of D5872 Metagenome in ORFs predicted in contigs and
singletons with ORFs longer than 100 aa ............................................................................ 30
Figure S7. Significant enrichment of SEED subsystems in D5872 Metagenome proteins
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over H. sulfidaeris Esulfide1 proteins .................................................................................. 31
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Figure S8. KEGG pathway depicting degradation of dicholorobenzoate as an example of the
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biodegradative capability of halogenated hydrocarbons amongst D5872 syntrophs ........... 32
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Figure S9. KEGG pathway depicting reductive fixation of ammonia into amino acids ....... 33
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Table S1. Geochemistry of formation fluid collected from the horizon 1.8 km (5872 feet) in
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depth of the Decatur Well, IL............................................................................................... 34
Table S2. Summary of D5872 microbial community identified by clone library and
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pyrosequencing..................................................................................................................... 35
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References ............................................................................................................................... 36
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Other Tables Attached as Excel Documents
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Table S3. Predicted proteins involved in translation, ribosome structure, and biogenesis in
D5872 Metagenome
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Table S4. Predicted proteins involved in transcription in D5872 Metagenome
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Table S5. Predicted proteins involved in replication, recombination, and DNA repair in
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D5872 Metagenome
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Table S6. tRNA genes identified in D5872 Metagenome
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Table S7. Comparison of SEED class2 category assignments in D5872 Metagenome contig
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and singleton proteins (D5872) and SEED microbial proteins (SEED)
Table S8. Predicted proteins involved in inorganic ion transport and metabolism in D5872
Metagenome
Table S9. Predicted proteins involved in carbohydrate transport and metabolisms in D5872
Metagenome
Table S10. Predicted proteins involved in osmolyte synthesis and degradation in D5872
Metagenome
Table S11. Predicted proteins involved in nucleotide transport and metabolism in D5872
Metagenome
Table S12. Predicted proteins involved in amino acid transport and metabolisms in D5872
Metagenome
Table S13. Predicted proteins involved in lipid transport and metabolisms in D5872
Metagenome
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Table S14. Predicted proteins involved in nitrate, nitrite and nitric oxide transport and
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metabolisms in D5872 Metagenome
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Table S15. Predicted proteins involved in defense mechanisms in D5872 Metagenome
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Table S16. Predicted proteins involved in cell motility in D5872 Metagenome
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Geologic and Hydrologic Setting
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A thorough understanding of the geologic and hydrologic history of the Paleozoic Mt. Simon
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Sandstone in the Illinois Basin is required to accurately decipher the ecology and evolution of
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its indigenous deep subsurface microbial community. Sedimentary rock reservoirs like the Mt.
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Simon Sandstone, often associated with hydrocarbon migration and entrapment, are also
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important targets for large-scale carbon capture, transport and storage (CCS) projects (Morse
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and Leetaru, 2003; DOE et al., 2007; Leetaru et al., 2009). The Mt. Simon Sandstone is a 494
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m-thick (1620 ft.) deposit of Cambrian-age sandstones at the base of the Sauk Sequence
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(Sloss, 1963; Buschbach, 1964) that unconformably overlies Precambrian weathered granites,
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granodiorites and rhyolites that comprise the basement of the Illinois Basin (Bradbury and
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Atherton, 1965; Atekwana, 1996). The uppermost basement surface was heavily eroded prior
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to deposition of the Mt. Simon Sandstone, creating a paleotopographic surface with as much
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as 600 m (2000 ft.) of vertical relief (Leetaru and McBride, 2009). The Mt. Simon Sandstone
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was therefore deposited in an arid high-relief depositional environment containing alluvial
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fans and braided river systems (Leetaru et al., 2009).
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The porosity and permeability of lithologies like the Mt. Simon Sandstone are
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strongly influenced by both depositional (primary) and post-depositional (diagenetic)
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alteration processes. The early Paleozoic section of the Illinois Basin in the Midcontinent of
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North America, which includes the Mt. Simon Sandstone, has experienced multiple events of
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regional subsurface fluid flow and resulting diagenesis that were driven by tectonic events
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(Bethke et al., 1984; Bethke, 1986; Bethke and Marshak, 1990; Bethke et al., 1991).
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Specifically, the Mt. Simon Sandstone experienced increasing subsidence and burial
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throughout the Paleozoic, having reached a maximum burial depth of 2.4 km and
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temperatures of 100°C in the early Mesozoic (Rowan et al., 2002; Makowitz et al., 2006).
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Several studies have been conducted on the Mt. Simon Sandstone formation waters
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(Visocky et al., 1985; Siegel, 1989). However, the wells from which these waters were
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sampled penetrated extremely shallow sections of the Mt. Simon Sandstone in the northern
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portions of the Illinois Basin. Furthermore, they do not provide a complete suite of elemental
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and isotopic water analyses. Therefore, interpretation of the origin and chemistry of the burial
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brines that precipitated the quartz cement overgrowths observed in the Mt. Simon Sandstone
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in the present study has been completed using analogous formation water collected from
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Silurian and Devonian strata in the Illinois Basin (Stueber and Walter, 1991). This approach
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was necessary because the geochemical analyses of deep subsurface Mt. Simon Sandstone
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formation water collected from the Illinois Basin–Decatur Project (IBDP) is currently being
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completed by the ISGS and the data are not yet available.
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Formation water from Silurian to Devonian strata in the Illinois Basin has a chloride
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(Cl)/bromide (Br) ratio that plots near a modeled seawater evaporation trajectory (Carpenter,
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1978) and stops just short of halite precipitation (Stueber and Walter, 1991). The Cl/Br ratio
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during seawater evaporation remains linear because both Cl and Br are inert in most
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biological and diagenetic processes until the brine reaches halite saturation (Stueber and
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Walter, 1991). At the point of halite precipitation Br (kD= 0.032) will preferentially remain
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within the brine as the Cl is incorporated into the halite (Carpenter, 1978; McCaffrey et al.,
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1987). This suggests that the Mt. Simon Sandstone formation water originated at the surface
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from evaporated seawater. In addition, cation/Br ratios within the Silurian to Devonian strata
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formation water suggest that water and rock interactions further modified the evaporated
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seawater (Stueber and Walter, 1991). For example, sodium (Na)/Br and potassium (K)/Br
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ratios are both depleted in the Silurian-Devonian formation water when compared to normal
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modern-day seawater, and fall below the seawater evaporation trajectory (Riley and Chester,
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1971; Carpenter, 1978; Stueber and Walter, 1991). This loss of Na is generally explained by
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the albitization of feldspar (Carpenter, 1978; Land and Prezbindowski, 1981; Stueber and
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Walter, 1991) and simultaneous loss of K during the precipitation of potassium feldspar
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cements (Egeberg and Aagaard, 1989), as represented by:
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2KAlSi3O8 + 2.5Al2Si2O5(OH)4 + Na+ =NaAlSi3O8 + 2KAl3Si3O10(OH)2 + 2SiO2 + 2.5H2O +
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H+
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This reaction is an additional potential source for the precipitation of quartz
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overgrowth cements within the Mt. Simon Sandstone, as well as clay cementation (Worden
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and Morad, 2000). Furthermore, the evaporation of seawater and the resulting increase in Na
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would tend to drive this reaction forward and further enhance quartz cementation.
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The distribution of Silurian and Devonian formation water Cl/Br along modeled
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seawater evaporation trajectories suggests that this modified subsurface evaporated seawater
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also experienced mixing with recharging meteoric water (Stueber and Walter, 1991). The
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possibility that evaporated seawater mixed with meteoric water and/or seawater has been
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evaluated by Land and Prezbindowski (1981). They suggest that the formation water
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chemistry, which is depleted in deuterium, is inconsistent with isotopic fractionation during
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seawater evaporation (Holser, 1979; Knauth and Beeunas, 1986; Stueber and Walter, 1991)
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Stueber and Walter (1991) suggest that, based on δD and δ18O covariations, the formation
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water was diluted by mixtures of as much as 50 % meteoric water. This addition of meteoric
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water into the Mt. Simon Sandstone may have been the source of oxygen to precipitate the
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multiple iron oxide cements in the Lower and Upper Stratigraphic Intervals.
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The Mt. Simon Sandstone formation water 87Sr/86Sr ratios are greater than the Sr-
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isotope ratios of Cambrian seawater (Veizer et al., 1999). This implies that dissolved Sr was
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derived from a source other than connate burial sweater within the Illinois Basin. Potential
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sources include: (1) overlying Paleozoic shales; (2) granites and rhyolites comprising the
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Precambrian igneous crystalline basement; and (3) shales and sands within the Mt. Simon
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Sandstone. Several previous studies have concluded that Silurian and Devonian formation
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water in the Illinois Basin derived dissolved radiogenic Sr from the New Albany shale group
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(Stueber et al., 1987; Bethke et al., 1991; Stueber and Walter, 1991). However, it is uncertain
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whether this would apply to the Mt. Simon Sandstone, which has no obvious vertical
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hydrologic connectivity with the overlying Eau Claire shale (Leetaru et al., 2009; Bowen et
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al., 2010). It is also possible that dissolved radiogenic Sr was derived from water-rock
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interaction with granites and rhyolites comprising the Precambrian basement (Clark and Fritz,
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1997). This hypothesis is viable due to the contact between the Mt. Simon Sandstone and the
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Precambrian basement. Further, the Precambrian basement would not need to contribute
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much radiogenic Sr because the 87Sr/86Sr ratio is likely very high (Clark and Fritz, 1997). The
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third hypothesis of deriving radiogenic Sr from shales and sands within the Mt. Simon
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Sandstone appears to be feasible, as well. Since the Mt. Simon Sandstone is primarily derived
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from Precambrian igneous basement, a significant concentration of radiogenic rubidium
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(87Rb) is expected (Clark and Fritz, 1997). 87Rb is directly linked to potassium (K)
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geochemistry, for which it will substitute readily for K+ in potassium feldspar, and decay into
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the daughter product, 87Sr, thus increasing the 87Sr/86Sr ratios (Clark and Fritz, 1997).
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Furthermore, the radiogenic Sr within the thin shale beds that occur in the Mt. Simon
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Sandstone appears to be highly accessible. Stueber et al. (1987) conducted leachate
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experiments on later Paleozoic shale formations within the Illinois Basin, which liberated
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high Sr ratios of 0.72793 to 0.75472. It is therefore reasonable to extrapolate these values to
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the shale within the Mt. Simon Sandstone.
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4
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Drilling and Subsurface Reservoir Analyses
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Well Drilling
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Drilling, geological analyses and formation water sampling were conducted at the IBDP well
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site in Decatur, Illinois (39°52’36.58” N, 88°53’36.00” W) by Schlumberger Carbon Services
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using multiple techniques and down hole tools. Detailed descriptions of these are available at
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http://www.slb.com and briefly summarized here. Drilling of the IBDP well began in
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February 2009. The maximum drilling depth of 2205 m was reached in April 26, 2009, and
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the well was completed in May 2009. Rotary drilling was conducted, which uses sections of
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hollow pipe (drillstem) that are screwed together and added as the depth of the well increases.
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Attached to the leading end of the drillstem is a tri-cone bit that cuts through the bedrock to
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produce a slurry mixture of drilling mud, formation water and broken up fragments of
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bedrock (cuttings). To prevent the borehole from becoming clogged by this drilling mud
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slurry and to cool and lubricate the drill bit, drilling fluids are continuously pumped down
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through the drillstem and out of the spinning drill bit. This mud and rock slurry then returns
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to the surface, where the cuttings are separated and the drilling muds are replenished and re-
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circulated back down into the drillstem. The drilling fluids consisted of proprietary mixture
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of bentonite clays, polymers (Guar Gum) and local groundwater, with the density, viscosity
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and other characteristics of the drilling mud continually adjusted based on the changing
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physical properties of the well with increasing depth of penetration (Phelps and Fredrickson,
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2002). Completion of the IBDP well required 16 tri-cone drill bits (ranging from 66 cm-, 45
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cm- and 31 cm-diameter) composed of a combination of steel, tungsten-carbide,
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polycrystalline diamond compact and diamond cutting and grinding edges. In addition, three
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well casings (surface, intermediate and production) were inserted into the borehole. During
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this drilling process, water and rock sampling was conducted with a drillstem test (DST),
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multiple downhole wireline logging tools, and borehole and sidewall rock coring.
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Rock Core Collection
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Three whole sections of borehole cores were taken during the drilling process; one in the Eau
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Claire Formation and two within the Mt. Simon Sandstone. Whole coring is the process of
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extracting cylindrical, intact rock samples from specific subsurface horizons using a
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specialized drill coring bit in conjunction with a core barrel and core catcher. The coring bit
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is open in the center, allowing the bit to travel around a cylindrical rock sample, which is then
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held in the core barrel. The core is held inside the core barrel with the core catcher until the
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core is extracted. These three whole cores were 9.14 m (30 ft) long and about 10.16 cm (4 in)
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in diameter. The depths at the top of the cores of the Eau Claire Formation and the two within
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the Mt. Simon Sandstone were 1668 m, 1950 m and 2057 m, respectively. The whole cores
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were used to establish the facies and depositional environments of the Mount Simon
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Sandstone (Bowen et al., 2010).
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Additional cores were acquired through the sidewall coring via percussion and
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mechanical sampling. Percussion sampling involves firing a hollow core bit into the
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formation and then extracting the bit with the rock core material inside (2.3 cm x 5 cm cores).
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Mechanical sampling utilized a Mechanical Sidewall Coring Tool (MSCT) hollow rotary drill
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core that cut into the formation and extracted similar sized core plugs. A total of 51 sidewall
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cores were collected from the Mount Simon Sandstone.
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Schlumberger Quicksilver Probe
The Quicksilver probe is a newly developed wireline formation tester (WFT)
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that was lowered into the IBDP wellbore on a wireline logging cable.
The
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WFT pumping system is designed such that the sterilized sampling bottles
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can be isolated from the formation water pumping system. Therefore,
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extensive pumping and associated pump tests can be completed without
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actually collecting the fluids and contaminating the sterilized and sealed
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sample collect bottles. A series preliminary pump tests were completed by
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positioning the probe at a variety of stratigraphic horizons within the Mt.
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Simon Sandstone that were identified from wire-line log data. This was
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completed in order to determine which stratigraphic horizons would produce
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a high enough flow rate for eventual pumping and sampling. Once the target
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stratigraphic sample horizon at 5872’ was determined with this technique,
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the probe was stabilized with lateral support arms against the wall of the
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well bore and dual straddle packers were positioned and inflated above and
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below the sample depth (Ayan et al., 2013).
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probes, the ®Quicksilver Probe is unique in that it is comprised of two pumps, which
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include: (1) an outer 20 cm-diameter concentric ring pump (called the guard probe); and (2)
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an inner 3 cm-diameter central pump (called the sample probe). When activated, both of
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these pumps extend horizontally from the probe to seat directly against the face of the
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borehole wall (described in detail at
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http://www.slb.com/services/characterization/wireline_open_hole/insitu_fluid/quicksilver_pr
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obe_wireline_sampling.aspx). The ®Quicksilver Probe simultaneously pumps fluid from the
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guard and sample probes with separated flowlines. The guard probe draws contaminated
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fluid from the outer portions of the target rock formation and these fluids are discarded into
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the wellbore below the packers. Conversely, the sample probe collects fluid from the cleaner
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central portion of the rock formation (Akkurt et al., 2006; Schlumberger, 2007). The water
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collected through the center sample pump is subjected to real-time geochemical monitoring
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with WFT sensors of K, Br and other drilling mud contaminants, until a minimum threshold
Compared to conventional WFT
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level of ≤ 4% mud infiltrate contamination is reached. In the case of the 1.8 km-deep sample
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collected from the Mt. Simon Sandstone, approximately 4L of formation water were
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collected after more than 2.5 hours of pumping and monitoring at depth. In addition, prior to
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the application in the present study, the ®Quicksilver Probe had been used exclusively to
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collect uncontaminated oil samples (Akkurt et al., 2006; Ayan et al., 2013).
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Wire-Line Log Electrical Data Collection
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The core data were used in conjunction with a suite of wireline collected using a
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Schlumberger Modular Formation Dynamics Tester (MDT), a wireline tool. A wireline tool
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is a cylindrical device that consists of one to multiple tools that each collects specific data,
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such as porosity and permeability. The wireline tool was lowered to the base of the well and
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was slowly pulled to the surface to provide wireline data. Wireline data, or logs, provide a
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better comprehension of the entire reservoir characteristics and well integrity by obtaining
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different parameters, such as porosity, density, lithologies, seismic profiling and casing
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cement, and can provide physical water and rock samples. The measurements used to create
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these logs were the gamma ray, neutron porosity, standard deviation permeability (SDR Perm)
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and Elemental Analysis (ELAN). The gamma ray log measurements recorded the amount of
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naturally occurring radioactivity within a deposit from elements, such as uranium and
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thorium. The neutron porosity log is based on the effect of a formation on neutrons emitted
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by a source. Hydrogen has the largest effect on slowing or capturing neutrons and because
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hydrogen is mainly in pore fluids, the porosity can be calculated (slb.com). The ELAN log is
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Schlumberger’s analysis of all the wireline logs that have been collected, which are combined
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together and adjusted to core data or fluid samples to create an overall log of the well.
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8
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Drillstem Test (DST)
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A drillstem test (DST) was conducted on drilling mud and formation water within the
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Ordovician St. Peter Formation on March 7, 2009. The DST tested well pressure and
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permeability, as well as permitted water collection by placing temporary packers, which are
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flexible elastomeric devices that inflate or expand to seal a wellbore, directly above and
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below the stratigraphic interval being sampled. The opposing pressure within drillstem was
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released through pressure step-down valves that allowed fluids within the targeted subsurface
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formation to flow up the drillstem as a result of subsurface formation pressure. The valves
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were then closed, the packers were removed and the drillstem was removed from the
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borehole. The mixture of cutting fluid slurry and formation water traveled up the drillstem
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and was collected at the surface. These drillstem samples, collected from each length of pipe
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as it is brought to the surface and disconnected, ranged in composition from nearly 100 %
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drilling mud to being formation water that was highly contaminated with drilling mud. The
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drilling mud collected was used as an “internal tracer” to evaluate the quality of formation
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water collected using a Quicksilver probe (as described below).
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The DST yielded 2 L of “less contaminated” formation water and approximately 212
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L of drilling mud and highly contaminated formation water from the St. Peter Formation.
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Water samples were collected in 19 liter buckets by catching the water that flowed out of the
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bottom of the each section of drillstem pipe as it was brought out of the ground and
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unscrewed. This broke the seal between the pipes and allowed the formation water to flow
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out from between the pipes at the surface. The 2 L sample of less contaminated formation
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water collected from the end of the drillstem was emptied straight into two autoclaved 1 L
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Nalgene bottles. Additionally, a small proportion of collected water was filtered with 0.45µm
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filters and analyzed by the ISGS to obtain the aqueous geochemistry. These 0.45µm filters
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were immediately collected, with sterile gloved hands and tweezers, and stored at -80˚C
9
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(Pinkart et al., 2002). After collection of the water samples and filters, the samples were
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returned to University of Illinois at Urbana-Champaign where the drilling mud and water
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samples were placed in a 4˚C dark cold room, with sample splits of drilling mud and
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formation water and the filters placed in a -80˚C freezer.
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Collection of Quicksilver Probe MDT Water Samples
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The clean formation water samples obtained from the Quicksilver probe modular dynamic
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formation tester (MDT) were used for gas collection, aqueous geochemical analysis and
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microbial DNA extractions. Prior to deployment and collection, the Quicksilver MDT
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probe was thoroughly cleaned and sanitized with multiple rinses of MilliQ water, bleach and
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ethanol in Schlumberger facilities in Brownstown, Texas, where it was immediately sterile
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wrapped, packed and shipped on Schlumberger trucks to Decatur, Illinois. Once the
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Quicksilver Probe MDT reached the IBDP well site, it was again cleaned and pressurized
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with 100 % ethanol before it was set down the borehole. When the probe reached the 1.8 km
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sample depth horizon within the borehole, collection of the formation water began by
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monitoring the resistivity in the flowlines in real-time. The factors that affect the actual
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contamination (e.g., vertical permeability, horizontal permeability, pump rate, formation fluid
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mobility and sanding potential) were taken into consideration and the tool pump was adjusted
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to the optimum speed and drawdown. Due to the contrast between the formation water and
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the liquid phase of drilling fluid, changes in resistivity could be observed during the transition
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from initial drilling mud filtrate to the virgin formation water. The pumping lasted until
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resistivity became asymptotical to be as low as 0.02 ohms and very constant. The resistivity
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acts a definitive tracer, which indicates that the drilling fluid has been evacuated and that
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contamination has been removed (Phelps and Fredrickson, 2002). Additional tracers and
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procedures that are presented by Phelps and Fredrickson (2002) were thoroughly considered,
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but ultimately abandoned due to high expenses.
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The 4 L sample bottle was sterilized, sealed and inserted into the ®Quicksilver Probe.
378
The sample bottle remained closed at depth until 2.5 hours of test pumping was completed
379
and the < 4% geochemical contamination level had been reached. At that time, the sample
380
bottle was filled in situ within the ®Quicksilver Probe and automatically sealed when full for
381
transport back to the surface. Immediately upon return to the surface, the 4 L sample bottle
382
was removed from the ®Quicksilver Probe within a mobile Schlumberger laboratory unit.
383
The formation water was then immediately offloaded under sterile conditions into sterilized 1
384
L Nalgene bottles, leaving no gas head in the bottles. There were then placed on ice within a
385
dark cooler and immediately transported 87 km back to the laboratories at the University of
386
Illinois at Urbana-Champaign (UIUC). Ensuing sample handling and treatment is described
387
below and under a series of sample preparation as introduced below.
388
389
Gas collection
390
The headspace gases of the MDT sample (D5872) were collected for isotopic analyses. Gas
391
collection was accomplished by using serum bottles that have been evacuated with nitrogen
392
gas, a 10 mL syringe, an 18-gauge needle, a 22-gauge needle and a T-valve. With gloved
393
hands and sterilized lab bench, the T-valve was placed onto the 10 mL syringe with the 18-
394
gauge needle. Using a Bunsen burner to heat up the 18-gauge needle, the needle was used to
395
penetrate through the cap of the 1L Nalgene bottle into the headspace. Quickly, the T-value
396
was opened and the syringe plunger was pulled back to 6 mL, which was followed
397
immediately by closing the T-valve. After replacing the 18-gauge needle with a 22-gauge
398
needle when the T-valve was closed, the 6 mL of gas was injected into the evacuated serum
399
bottles. The bottles were then sent to Isotech Laboratories Inc., IL, for isotopic analyses of
400
gas components.
401
11
402
Centrifugation of water samples
403
The formation water collected from the MDT was centrifuged on a Thermo Scientific Sorvall
404
Lengend RT Plus fixed angle centrifuge (Thermo Scientific, MA) immediately after it was
405
transported back to the laboratory at UIUC. The centrifugation process was conducted with
406
gloved hands on a clean lab bench and all the samples were kept at 4˚C. The 1L Nalgene
407
samples were opened in a high air flow environment to eliminate airborne contamination.
408
Negative controls were created and treated identically to the samples. The centrifugation was
409
performed at 9,000 rpm for 20 minutes at 4˚C. The supernatant from this operation was
410
decanted into sterilized Nalgene bottles and stored in the 4˚C room. The pellets were pooled
411
and washed with pre-cooled sterile 1×PBS buffer solution (pH 7.4) twice. The final pellets
412
were then re-suspended in RNAlater® (Ambion Inc., TX), incubated at 4 ˚C for 24 hours
413
before stored in the -80˚C freezer for long-term storage.
414
415
Geochemical Analyses
416
Geochemical parameters, such as pH, conductivity and electoral potential (Eh), were
417
measured on the freshly collected formation water in the field with the ORION® 5-Star
418
pH/RDO*/Conductivity portable meters (Thermo Scientific, MA) connected with appropriate
419
probes. Density was measured with a portable density meter (Mettler-Toledo Inc., OH).
420
Supernatant of centrifuged samples was promptly aliquoted and sent on ice to the
421
geochemistry laboratory of the ISGS, the Nuclear Engineering Teaching Laboratory of the
422
University of Texas, Austin and the Prairie Analytical Systems, Inc (Prairie Lab)
423
(Springfield, IL) to quantify ion and trace metal concentrations. The samples were kept in at
424
4 °C in dark before analyses. In order to assess the systematic errors at different laboratories,
425
some of the parameters were repeated in different labs. Ion composition and concentrations
426
were determined at ISGS by using a Dionex 211i ion chromatograph (IC) (Dionex Company,
12
427
CA) installed with an Ionpac AG14 Guard Column, an Ionpac AS14 Analytical Column, and
428
a CDM-3 conductivity detector cell with a DS4 detection stabilizer following U.S. EPA
429
Method 300.0 (Pfaff, 1993). Some of the ion and metal quantification were performed with
430
an IC and an inductively coupled plasma mass spectroscopy (ICP-MS) at the Prairie Lab.
431
TOC was measured with a Shimadzu TOC-V CSN (Shimadzu Cooperation, Japan).
432
433
Strontium Isotopic Analysis
434
Sr analyses were completed on the formation water from the MDT to obtain the modern
435
87
436
conjunction with solid rock samples. The preparation for these samples started with the Sr-
437
spec resin and Sr columns under laminar flow hoods with gloves in a clean lab. The Sr-spec
438
resin by Eichrom was thoroughly cleaned by adding ultrapure deionized (DI) water to the
439
resin and leaving under a heat lamp overnight, then decanted and refilled with more ultrapure
440
DI water. This process was repeated three times. The Teflon Sr columns were then loaded
441
with the Sr-spec resin slurry, using the upmost care not to create any bubbles within the resin
442
by letting the Sr-spec resin slowly settle. The columns were then washed with one full
443
reservoir >2mL) of 3N HNO3, followed by one full reservoir of ultrapure DI water and 1mL
444
of 3N HNO3 to precondition the column for samples.
445
Sr/86Sr ratios for further diagenetic episodes and basinal brine fluid migrations in
Once the resin and columns were created, Sr samples were prepared in laminar flow
446
hoods in a clean lab with gloved hands by drying down from 1mL from the centrifuged water
447
samples in acid (HCl) washed beakers. A negative control and a duplicate were treated
448
identically to the samples. When a small solid formed from the dry down process, the
449
samples were brought up into 4 mL of 3N nitric acid (HNO3). When the Sr columns were
450
preconditioned with the 1mL of 3 N HNO3 (see above), 100µL of the sample was added to
451
the columns and allowed to flow through the column completely. Next, three additions of
13
452
2mL 3N HNO3 was added to the columns and allowed to completely drip through the column.
453
This was followed by two additions of 2mL of warm 0.05N HNO3 to elute the Sr. The eluted
454
Sr was collected in new, HCl cleaned beakers. The collections from the 0.05N HNO3 were
455
evaporated to dryness and 50 µL of concentrated (10.5 N) HNO3 was added to the small solid
456
to dry down again, which will break up organic residues.
457
The samples were brought up in 2% HNO3 with a 1:100 dilution in a laminar flow
458
hood for mass spectrometer analyses. The Sr analyses were conducted on a Nu Plasma HR
459
(MC-ICP-MS) (multicollector inductively-coupled-plasma mass spectrometer) with
460
consistent plasma parameters of: (1) coolant gas: 13 L/min; (2) auxiliary gas: 0.9 L/min; (3)
461
RF power: 1300W. The mass spectrometer has an electrostatic analyzer (ESA) that is used to
462
filter the ions by their kinetic energies and is followed by a magnetic sector, used to separate
463
ions to their mass over charge. By combining this magnetic sector with an electrostatic
464
analyzer, creates double-focusing and allows for exceptional resolution with “flat-topped”
465
peak signals.
466
467
Microscopic Analyses
468
Lithology analysis of thin sections
469
Polished uncovered thin sections (30 µm in thickness) were prepared from the chosen
470
sidewall cores by Wagner Petrographic, UT (http://www.wagnerpetrophic.com). Each thin
471
section was analyzed under plane-light (PL) and polarized light using an Axiovert 200M
472
Fluorescent Microscope with Apotome (Carl Zeiss Company, Oberkochen, Germany)
473
operated by Axiovision software, allowing for ultrahigh resolution imaging and microscopy
474
(125-200 nanometer working resolution). Images were collected with an AxioCam MRc5
475
color camera and an AxioCam MRm high-resolution camera and analyzed for grain size and
14
476
shape using the Axiovision software. All of the sidewall core lithologies were described
477
using the siliciclastic sedimentary rock classification scheme (Williams et al., 1982).
478
479
Fluorescence In-Situ Hybridization (FISH) for cell enumeration
480
Microbial samples were stained with fluorescence dyes for enumeration of microorganisms
481
with a method modified from Pernthaler et al. (2000). Right after arriving at the laboratory
482
on ice, cells in 1 liter of MDT formation water and about 200 mL of the DST slurry were
483
concentrated by centrifuging at 8,500 rpm for 20 min at 4 °C to collect biomass. The pellets
484
were pooled, washed twice with 15 mL PBS buffer and fixed with 4 % paraformaldehyde
485
solution at 4 °C for 4 hours. The fixed sample was washed twice with PBS and the stored in
486
PBS and ethanol mixture (1:1, v:v) at -20 °C before staining and microscopy observation.
487
The fixed samples were diluted in 1 mL PBS buffer solution at the proper dilution ratio.
488
In order to minimize unspecific stain with soil matrix (e.g. clay), a series of dilution from
489
1:10 to 1:1000 were performed in PBS buffer. The samples were filtered through black
490
polycarbonate filters with 0.22 m pores (GE Water and Process Technologies, PA) and air-
491
dried. Then, they were sequentially hybridized with cy3 labeled EUB 338 probe (EUB338-
492
cy3) (5 µg/mL) (Integrated DNA Technologies Inc., IA), 5×Sybr Green I (Invitrogen Corp.,
493
CA) and TOPRO3 (5 µg/mL) (Sigma-Aldrich Corp., MO). A series of preliminary
494
experiments were performed on the samples from D5872 to optimize the experimental
495
conditions. Wavelength channels were carefully selected to achieve the minimal overlap of
496
different fluorescence dyes. As shown in Fig. S1, the spots visualized in the cy3 channel are
497
not all co-localized in the Sybr Green I, suggesting no significant overlap occurred between
498
these two channels. In our preliminary experiment, weak signal intensity and smear signals
499
were observed in the DAPI channel. Thus, the DAPI dye employed in the original protocol
500
(Pernthaler et al., 2000) was replaced with TOPRO3. Compared to DAPI, TOPRO3 exhibits
15
501
several advantages, including: (1) TOPRO3 is very specific to DNA; (2) the Far Red channel
502
(emission wavelengths above 650 nm) compared to DAPI (400-470 nm), does not interfere
503
with the natural autofluorescence inherent to organic matter (which is possible using the
504
DAPI channel). Therefore, emission the wavelengths above 650 nm observed for most
505
biological and organic compound autofluorescence is effectively minimized. EUB338-cy3,
506
the universal probe for bacteria (Amann et al., 1995), was used in this study for the
507
enumeration of the cell density in the D5872 and drilling mud samples. Staining was
508
performed with a hybridization buffer containing 30 % formamide (Sigma-Aldrich Corp.,
509
MO). It was followed with hybridization by 5×Sybr Green I solution for 30 min in dark and
510
staining by TOPRO3 for 3 min. For every hybridization step, excessive dye was washed by
511
washing solution (for EUB338-cy3) (Pernthaler et al., 2000) or PBS buffer (for Sybr Green I
512
and TOPRO3). The membranes were mounted with 1-2 drops of ProLong® Gold Antifade
513
Reagent (Invitrogen Corp., CA) and covered with cover slips. After keeping the samples at
514
room temperature in dark for 24 hours, the cover slips were sealed with nail polish and the
515
samples were stored at 4°C before they were viewed under the confocal microscope.
516
With 40×-oil emersion lens and the Zeiss Axiovert 200M with the Apotome Structured
517
Illumination Optical Sectioning System with appropriate filters, about 12 to 25 Z-stack
518
images were taken (at 5 m interval and 60-125 m total distance depending on the thickness
519
of the area) for each spot. An extended depth of focus image using all focal planes was
520
created in the Axiovision program on each channel. The pictures taken from the three
521
channels were overlapped and only the overlapping regions of EUB338, SYBR Green I and
522
TOPRO3 signals were identified and defined as stained cells. Such cells turned white after
523
this process and all the pixels in other colors than white were discarded with the aid of Adobe
524
Photoshop (Adobe Systems, CA). For each sample, such operations were repeated for at least
525
10 spots randomly distributed on the membrane. The number of cells per unit volume of
16
526
formation water was calculated as average number of signals per unit of area normalized by
527
the area of filtered sample on the membrane, dilution ratio during sample preparation and
528
initial volume of formation water used for FISH.
529
Cultivation of Organisms
530
Significant efforts were made to grow culturable organisms from the D5872 formation water
531
sample. The groundwater used for cultivation was flushed with sterile cotton filtered N2:CO2
532
(80:20, v:v) and stored at 4 °C in dark right after arriving at the UIUC laboratory before
533
development of microcosms within a week. A variety of experimental conditions, including
534
media, salt concentrations and nutrients were created. The pH of the media was buffered
535
with 20 mM bicarbonate and N2:CO2 (80:20, v:v) headspace for the anaerobic media. The
536
sterilized anaerobic media were prepared under N2:CO2 (80:20, v:v) headspace and in the
537
serum tubes sealed with butyl rubber stoppers and aluminum seals. All the amendments were
538
added from sterilized anaerobic stock solutions.
539
Two media, including the synthetic groundwater (Roh et al., 2002) and R2A medium
540
were selected. A series of geochemical conditions were created, including fermentation, iron-,
541
nitrate-, and sulfate-reducing conditions with glucose, Fe(III)-citrate, NaNO3 and Na2SO4 (5
542
mM each), respectively. A fatty acid mixture (acetate, lactate, butyrate (5 mM) each) as
543
electron donors and carbon source was amended in the non-fermentation microcosms. Under
544
each condition, groundwater was injected with the final volume 10 and 50 % of the total
545
volume, respectively, both to adjust the final salt concentrations and as the source of
546
indigenous organisms.
547
The cultures were stored still at 37 °C in dark and manually shaken every 3-5 days.
548
In order to determine whether organisms were successfully cultured, the cultures were read
549
for OD600 using a SPECTRONIC 20D+ spectrophotometer (Thermos Scientific, MA) and
550
observed under a Nikon optical microscopy (Nikon Instruments Inc., Japan) every 3-5 days
17
551
for the first month and at the weekly frequency for the rest of three months. The microcosms
552
filtered through 0.22 mm membrane was measured at monthly frequency for the changes of
553
fatty acid concentrations using an HPLC.
554
Microbial Community Analyses
555
DNA extraction
556
High molecular weight genomic DNA was extracted directly from the frozen centrifuged
557
pellets (as described in Centrifugation of water samples above) using a procedure similar to
558
that of Tsai and Olson (1991). All reagents used were of molecular-biology grade or higher
559
purity. Cells were lysed by incubating one filter at 37°C in the lysis solution (0.15 M NaCl,
560
0.1 M Na2EDTA, pH=8.0) with 15 mg lysozyme per mL, followed by another incubation in
561
STS solution (0.1 M NaCl, 0.5 M Tris-Cl, pH=8.0, 10% sodium dodecyl sulfate) (1991) .
562
After freeze-thaw cycling between a 55°C water bath and liquid nitrogen to further disrupt
563
cell membranes for three times, proteins were removed through sequential extractions with
564
equal volumes of phenol (pH=7.8), phenol:chloroform:isoamyl alcohol (25:24:1, pH=7.8)
565
and chloroform:isoamyl alcohol (24:1). DNA, which remained in the aqueous phase, was
566
precipitated in isopropanol containing 2.5 M ammonium acetate and 100 µg/mL glycogen
567
(Ambion Inc., TX). The centrifuged DNA pellet was then air dried and resuspended in
568
nuclease-free TE buffer. We confirmed both the quality and quantity of DNA recovered by
569
gel electrophoresis. DNA samples were loaded onto a 0.8 % agarose gel and stained with
570
ethidium bromide. The size and intensity of the genomic DNA bands were compared visually
571
with a quantitative DNA ladder (Hyperladder I, Bioline USA, Boston, MA). Due to the low
572
biomass in the environmental samples, DNA may be present but not visible via gel
573
electrophoresis.
574
18
575
Development of 16S rRNA Clone library
576
In order to develop clone libraries for full-length 16S rRNA genes, PCR was performed with
577
primers for Bacteria 8F (5′-AGA GTT TGA TCC TGG CTC AG-3′) or Archaea (21F 5’-
578
TCC GGT TGA TCC YGC CGG-3’) with a universally conserved reverse primer (1492R, 5′-
579
GGT TAC CTT GTT ACG ACT T-3′). PCR amplification was performed in 50 μL reaction
580
volumes using the TaKaRa Ex TaqTM kit (Takara Bio Inc., CA) following the manufacturer’s
581
instructions. Amplification began with a denaturing hot start at 95°C and was followed by 25
582
cycles of the following: 30 s of denaturing at 94°C, 30 s for primer annealing at 55°C, and 90
583
s for extension at 72°C. The PCR was completed with a final extension step at 72°C for 5
584
minutes. Size and intensity of the PCR product was confirmed with agarose electrophoresis
585
as described above. Considering the precaution of contamination, appropriate negative
586
controls were used to ensure no sample was contaminated with exogenous DNA. Equivalent
587
volume of sterile ultrapure water was filtered in the same manner as the formation water
588
samples and then subjected the filter to the same DNA extraction procedure. A negative
589
procedural control containing no sample during DNA extraction was prepared as well. No
590
DNA was detected in any control extraction by gel electrophoresis, and no amplification was
591
observed in any PCR reactions using these samples as a template. The amplified PCR
592
products were purified with Qiaquick PCR purification kit (Qiagen Inc., CA). Purified PCR
593
products were cloned into p-GEMT Easy Vector® and were transformed into JM109 High
594
Efficiency Competent Cells as instructed by the manufacturer (Promega Corporation, WI).
595
For each 16S rRNA clone library, a total of 196 recombinant plasmids were extracted from
596
randomly picked clones and sequenced at the Illinois Biotechnology Center of University of
597
Illinois, Urbana.
598
599
The full-length 16S rRNA gene sequences from clone libraries were assembled with
Sequencher 4.9 (Gene Codes Corporation, Ann Arbor, Michigan). Sequences with low
19
600
qualities were eliminated. Alignments and distant matrices of assembled sequences were
601
generated with the aid of NAST (DeSantis et al., 2006b) and PHYLIP (Retief, 2000)
602
programs from Greengenes (DeSantis et al., 2006a). All sequences were checked for chimeric
603
artifacts by Bellerophon (version 3) (Huber et al., 2004) and the chimeras were removed
604
from the datasets before the following sequence analyses. Taxonomic classification was
605
determined by using the sequence match tool of the Ribosomal Database Project (RDP) II
606
database (Maidak et al., 2001) and double checked with the sequence classification program
607
from the Greengenes (DeSantis et al., 2006a). Operational taxonomic units (OTU) were
608
determined at sequence similarity levels of 99 %. One representative sequence per OTU was
609
used to construct phylogenetic trees based on neighbor-joining algorithm (Jukes-Cantor
610
correction) in ARB (Ludwig et al., 2004). The robustness of the inferred tree topologies was
611
evaluated after 1000 bootstrap replicates of the neighbor-joining data.
612
613
Terminal Restriction Fragment Length Polymorphism (T-RFLP)
614
Preparation of T-RFLP samples was similar to that in development of clone library. Full-
615
length 16S rRNA genes were amplified from genomic DNA using FAM-labeled 8F and
616
1492R at the same thermal condition. The PCR products were purified using QIAquick® PCR
617
purification Kit (Qiagen Inc., CA). The purified PCR products were digested with RsaI (New
618
England Biolabs Inc., MA) and analyzed with an AB 3730xl DNA Analyzer and using
619
ROX1000 as the standard. The results were visualized using the GeneMapper® software and
620
analyzed with a Microsoft Excel script designed by our group.
621
20
622
Metagenomic Analyses
623
V1-V3 16S rRNA Hypervariable Region Pyrosequencing
624
PCR products for bar-coded pyrosequencing were amplified with the primer pair targeting
625
V1-V3 hypervariable region of 16S rRNA genes (position 27-338 (V1-V3), Escherichia coli
626
numbering). Each primer was attached with a 12 bp Roche 454 pyrosequencing bar-code
627
sequence, a TC liner and a Roche 454 sequencing adapter. Amplification, pooling and
628
pyrosequencing were performed as described (Jindou et al., 2008). The single band of PCR
629
products was purified from the cut agarose gel (0.8 %) with QIAGENE® Gel Extraction Kit
630
and the concentrations were quantified with a Qubit® Fluorometer (Invitrogen Co., CA) after
631
staining the DNA using a Quant-iT™ dsDNA BR Assay Kit (Invitrogen Co., CA). Equal
632
amount of PCR products for each sample were pooled and be run on a Roche Titanium 454
633
pyrosequencing machine at the W. M. Keck Center, University of Illinois, Urbana-
634
Champaign.
635
Pyrosequencing reads were first assessed for quality by using several criteria: reads that
636
had degenerate bases, overall low-quality score, without a complete barcode and primer at
637
one end or shorter than 300 bp were removed. The remaining reads were aligned against a
638
SILVA based bacterial reference (Pruesse et al., 2007), and chimeras were determined with
639
the ChimeraSlayer package in Mothur (Haas et al., 2011) and excluded from further analyses.
640
The remaining sequences were trimmed with the aid of Jalview (Clamp et al., 2004) to cover
641
the same aligned region. The reads were then preclustered (Huse et al., 2010) and assigned
642
into OTUs to generate rarefaction curves. Sequencing recovery was estimated using non-
643
parametric abundance-based coverage estimator (Chao and Lee, 1992) at different
644
dissimilarity levels. The pyrosequencing reads were classified into different taxonomic
645
groups by comparing them with those entries in the SILVA database bacterial reference
646
(Pruesse et al., 2007). The number of representative reads for each OUT determined at 97 %
21
647
as the cutoff value at each taxonomic level was counted and the proportions of each group in
648
a sample were calculated.
649
650
Halomonas sulfidaeris strain Esulfide1 as a Reference Genome for D5872 Metagenome
651
Halomonas sulfidaeris strain Esulfide 1 (H. sulfidaeris Esulfide1) was selected as the
652
reference genome for the D5872 Metagenome. The culture was purchased from the America
653
Type Culture Collection (ATCC) and grown on the agar plates containing the Halomonas
654
medium as recommended by ATCC and 15 % agar. Single colonies were picked and
655
inoculated in liquid Halomonas medium for 36-48 hours at 30 C. After the cells were
656
collected by centrifugation, genomic DNA was extracted with the method as described
657
above.
658
659
Pyrosequencing of D5872 Metagenome and H. sulfidaeris Esulfide1 genome
660
Genomic DNA of D5872 H. sulfidaeris-dominated microbial community and H. sulfidaeris
661
Esulfide1 was sequenced at the W. M. Keck Center, University of Illinois Urbana-
662
Champaign. A combination of 454 shotgun, 454 paired-ends (3-5kb inserts), and Illumina
663
sequences (500 bp inserts) were performed (Kumar et al., In preparation) using a Roche GS-
664
FLX sequencer and a Illumina HiSeq2000, respectively. The libraries for 454 Roche shotgun
665
and mate-pair sequencing of 3kb libraries were prepared following the standard protocols
666
from Roche (Roche). For Illumina sequencing, the genomic DNA was fragmented into
667
roughly 500 bp reads using standard Illumina protocols and paired-end sequenced.
668
669
Metagenomic and Genomic Analyses
670
The genomic reads sequenced using 454 array-based pyrosequencing (Ronaghi, 2001) were
671
assembled using the Newbler assembler v2.3 (www.454.com). Reads were quality-trimmed
22
672
using default trimming provided by Newbler. For the metagenome, the unassembled raw
673
reads (singletons) were trimmed using Newbler’s quality trimming, and singletons <100 bp
674
in length were removed. Sequences that were identical to each other were also removed in
675
order to retain a unique sequence set. Stringent parameters for minimum overlap between two
676
reads (60 bp in length and 95% in identity) were used to prevent assembly into chimeric
677
contigs. Assembly was compared with different parameters (default) and versions (v2.0) of
678
the Newbler assembler, and GC-content distribution of the contigs were compared between
679
these assemblies to ensure a high quality assembly.
680
tRNA genes were predicted in the contigs and singletons using tRNAScan-SE (Lowe
681
and Eddy, 1997; Schattner et al., 2005), and rRNA were predicted using a Python based
682
script, rna_hmm3 (Huang et al., 2011), that runs on HMMER3.0 (Eddy, 2009) to search for
683
partial 5S, 16S, and 23S genes in the metagenome contigs. From the predicted coordinates,
684
the rRNA sequences were parsed and the corresponding sequences were assembled using the
685
CAP3 sequence assembly program (Huang and Madan, 1999) to identify complete operons.
686
Metagenomic reads often contain partial open-reading frames (ORFs). MetaGeneMark (Zhu
687
et al., 2010) was used to predict protein encoded genes in the contigs and singletons allowing
688
for partial ORFs. The program uses hidden Markov models (HMM) trained on short bacterial
689
and archaeal sequences based on their GC-content and codon usage. MetaGeneMark
690
predictions were shown to compare favorably with other prediction tools, such as MetaGene
691
(Noguchi et al., 2006) and MetaGeneAnnotator (Noguchi et al., 2008). Duplicates amongst
692
the predicted proteins were removed from the corresponding ORFs. In order to determine the
693
percentage of proteins representative of the reference genome, the predicted proteins were
694
searched against the predicted proteins of the reference genome H. sulfidaeris Esulfide1
695
(Kumar et al., In preparation). Conserved protein families, domains, and motifs were
696
predicted in the metagenome translated ORFs by searching Pfam (Finn et al., 2010) and
23
697
TIGRfam (Haft et al., 2001) databases using HMMER3.0 (Eddy, 2009). Known enzymes
698
were predicted using PRIAM (Claudel-Renard et al., 2003) at an e-value cutoff of 10-10. This
699
was followed by searching the proteins against the non-redundant (nr) database at an e-value
700
of 10-5 using Blastp (Altschul et al., 1997). The Blastp results were used as input for
701
taxonomic assignment of the ORFs using MEGAN v4 (Huson et al., 2007), and also for
702
assigning ORFs to known SEED categories.
703
Taxonomic classification of the assembled contigs and singletons was performed
704
using NCBI Blastp (Altschul et al., 1990; Altschul et al., 1997) search against the NCBI non-
705
redundant protein database (nr) and the gi numbers (NCBI sequence identifiers) for the top
706
10 hits for each protein were picked. Of the 42,408 (91%) proteins that had hits at an E-value
707
of <10-5, only 109 proteins were unassigned to any taxon or assigned as root. Using a SEED
708
server script, the corresponding SEED protein-encoding gene (PEG) IDs for the gi
709
numbers were identified in theSEED database (Aziz et al., 2008; Meyer et al., 2008).
710
Sorted by e-values for each gene, the PEGs were mapped to the subsystems within
711
theSEED database and the first subsystem that exists for a PEG was picked. For
712
theSEED classification, Level 2 within the hierarchy was used where possible.
713
Otherwise, Level 1 classification of theSEED was used. In order to identify enriched
714
SEED categories in the metagenome, counts of proteins assigned to each SEED
715
category for the metagenome proteins were compared to those for proteins in the
716
entire SEED database using the Fischer’s exact test. To correct for multiple
717
comparisons, a false discovery rate error adjustment to the P-values was made using
718
the Bejamini-Hochberg method (Benjamini and Hochberg, 1995). All categories that
719
were below an adjusted P-value of <0.05 were considered significantly enriched in the
720
metagenome. The predicted proteins were also classified on the basis of Cluster of
24
721
Orthologous (COG) domains using reverse-position-specific Blast (rpsBlast)
722
(Marchler-Bauer et al., 2002) search against a database of position-specific scoring
723
matrices (PSSMs) of automatically aligned sequences classified in the COG resource
724
for unicellular species (Tatusov et al., 2003). The rpsBlast search results were parsed,
725
followed by generation of counts of metagenome proteins that were assigned to each
726
COG category.
727
Figures and Tables
(a) Sybr Green I
5 mm
(c) TOPRO3
5 mm
(b) EUB338-cy3
5 mm
(d) Overlapped
5 mm
728
729
Figure S1. Fluorescence hybridization of the D5872 microbial consortium by Sybr Green (a),
730
cy3 labeled probe EUB338 (b) and TOPRO3 (c). Subfigure (d) shows overlapped pictures (a-
731
c). The spots appearing at the same location in (a)-(c) will overlap in (d) and are thus
25
732
determined as an effectively stained cell. The arrows point to one of the identified stained
733
cells.
26
1
(a) D5872
872
Fraction
0.8
0.6
0.4
0.2
421
165
645 700
859
888
0
0
300
600
900
Length of Fragments
1
(b) DDM
Fraction
0.8
0.6
646
0.4
0.2
119
422-423
467
307
470 562
823
881
869 896
0
0
734
300
600
900
Length of Fragments
735
Figure S2. T-RFLP analysis of D5872 microbial community (a) and drilling mud (b). Full-
736
length 16S rRNA genes were multiplied with bacterial primers 8F and 1492R and digested
737
with restriction enzyme RsaI. Individual fragment abundance was normalized to the total
738
abundance and shown as the fraction. The values above the peaks indicate the length of the
739
corresponding fragments in the unit of base pairs.
740
27
741
(a)
(b)
742
743
744
745
746
747
748
749
750
751
752
753
754
Figure S3. Phylogenetic relationship of Halomonas species in D5872 microbial community
755
versus the DST drilling mud (DDM) by 454 pyrosequencing reads targeting V1-V3
756
hypervariable region of 16S rRNA genes (a) and comparison of abundance of each OTU in
757
the two samples (b). Each sample code indicates one OTU identified at 97 % cutoff value.
758
Escherichia coli (X80725) was used as the out-group and the scale bar indicates 0.1 changes
759
per nucleotide position. The accession number of each type strain is in parentheses. The
760
values listed in the parentheses after each OTU indicate numbers of sequences in the D5872
761
versus those in the drilling mud. The abundance of each OUT is proportional to the size of
762
bars in (b).
763
764
765
28
766
(a)
(b)
200
250
D5872
DDM
OTUs
150
OTUs
D5872
200
100
50
DDM
150
100
50
0
0
0
50
100
150
Number of Reads
200
0
2000
4000 6000 8000
Number of Reads
10000
767
768
Figure S4. Rarefaction of D5872 and the DST drilling mud (DDM) microbial community
769
richness using full-length 16S rRNA genes (a) and pyrosequencing reads of the V1-V3
770
hypervariable region of 16S rRNA genes (b). The cutoff values for (a) and (b) are 99 % and
771
97 %, respectively.
772
29
773
774
775
776
777
778
779
780
781
782
783
Figure S5. Distribution of GC-content with length for D5872 Metagenome contigs. The solid
784
black line represents the average GC-content for H. sulfidaeris Esulfide1 (0.54).
785
786
30
(a)
(b)
787
788
789
Figure S6. Taxonomic assignment of D5872 Metagenome ORFs predicted in contigs (a) and
790
singletons with ORFs longer than 100 aa (b). The ‘Others’ category comprises a mix of
791
diverse taxa and includes proteins, majority of which were classified within root, bacteria,
792
and cellular organisms.
793
31
794
795
Figure S7. Significant enrichment of SEED subsystems in D5872 Metagenome proteins over
796
H. sulfidaeris Esulfide1 proteins. X-axis indicates the negative log of P-value. Y-axis denotes
797
the difference in percentage of proteins in D5872 Metagenome and H. sulfidaeris Esulfide1
798
that are assigned to SEED subsystems. The enriched subsystems are highlighted in red.
799
800
801
32
802
803
Figure S8. KEGG pathway depicting degradation of dicholorobenzoate as an example of the
804
biodegradative capability of halogenated hydrocarbons amongst D5872 Metagenome.
805
Enzymes highlighted in green are the ones present in D5872 Metagenome.
806
33
807
808
Figure S9. KEGG pathway depicting reductive fixation of ammonia into amino acids.
809
Enzymes highlighted in green are the ones present in D5872 Metagenome.
34
810
Table S1. Geochemistry of formation fluid collected from the horizon 1.8 km (5872 feet) in
811
depth of the Decatur Well, ILa.
Geochemical Parameters
Depth (m)
T (°C)
Formation Pressure (bar)
Redox Potential (Eh) (mV/pH unit)
D5872
1789
49.9
181
166.5
Drilling Mud
Wellhead
26.5
NAb
NA
7.98
NA
c
BDL
NA
240011*
14412
15.70±0.28
213.4±16.2
52767±5552
2.025±0.035
6.45±0.27
BDL
2.30
503±15
7260
BDL
21200
642
12.20
12732±987
BDL
292.300.71
4.98±0.62
3.03
BDL
2392±11
NA
BDL
BDL
BDL
350±77
131.0±4.2
4.65±0.35
48±11
52.5±4.3
1.14±0.05
1581±128
0.0634±0.0032
-21.9
-35.16
-63
0.71123±0.00001
(1.5±0.4)106
NA
NA
NA
NA
(8.4±3.5)1010
pH (0.3)
Dissolved oxygen (DO)
TOC (mg/L)d
TDS (×103 ppm)e, f
F- (ppm)
Br- (ppm)
Cl- (ppm)
I- (ppm)
NO3--N (ppm)
PO43- (ppm)
Ba2+(ppm)
SO42-(ppm)
Ca2+(ppm)
Fe2+(ppm)
K+(ppm)
Mg2+(ppm)
Mn2+(ppm)
Na+(ppm)
Zn2+(ppm)
Isotope (‰)
13CO2
13CH4
13DC1
87Sr/86Sr
Cell density (cells/L)
812
a
813
on filtered formation fluid or gaseous phase by the laboratories at UIUC, ISGS, UT-Austin
814
and Isotech Inc. Average was calculated if a parameter was measured at different
815
laboratories and uncertainties indicate standard deviation of replicate measurements. b NA:
816
not measured. c BDL: below detection limit. d TOC: total organic carbon. e TDS: total
817
dissolved salts. The samples were measured after passing through 0.45 m filter membrane;
818
ppm indicates parts per million. f The detection limits for the ions are (ppm): F- (0.05); Br-
819
(0.25); Cl- (3.5); I- (0.1); NO3- (0.25); Ba2+ (0.5); PO43- (0.8); SO42- (0.25); Ca2+ (10); Fe2+ (14);
820
K+ (19); Mn2+ (0.1); Na+ (12); Zn2+ (0.001).
Temperature, Eh, pH and DO were measured in-situ, while other parameters were measured
35
821
822
Table S2. Summary of D5872 microbial community identified by clone library and
pyrosequencing
16S rRNA clone library
Number of sequences/reads
Number of OTUa
Number of sequences/reads
affiliated to genus Halomonas
Closest type strain related to the
dominant sequences/reads
Coverage
823
824
825
V1-V3 pyrosequencing
D5872
166
9
Drilling Mud
156
39
D5872
8856
120
Drilling Mud
9918
201
163
4
8637
237
H. sulfidaeris
H. neptunia
H. sulfidaeris
0.98
0.84
0.995
H. neptunia and
H. sulfidaeris
0.992
a
99 % and 97 % were used as the cutoff values for OTUs in analyses of 16S rRNA clone library and
pyrosequencing, respectively.
36
826
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