Meredith Nevers Muruleedhara Byappanahali 20200211 spreadsheet Reston, VA U.S. Geological Survey https://doi.org/10.5066/P9NFKBEB The data associated with the following data release were collected between 2016 and 2017 at three locations on Lake Michigan: Racine, WI; Chicago, IL; and East Chicago, IN. Individual water samples were collected one day a week for ten weeks between June and August. Samples were collected from eight specific sites made up of two river and six shoreline type environments. Sampling was completed at sites where various morphology (embayment, sand and sediment characteristics, size and shape) and hydrologic conditions (currents and waves) were present. Then samples were analyzed using microbial communities (metagenomic analysis), markers of contamination (microbial source tracking), and fecal indicator bacteria (E. coli). The data being released were part of a project funded by the Great Lakes Restoration Initiative (GLRI) explaining the dynamics of water quality contamination. The project sought to characterize shoreline health, focusing on water and sand interactions. More specifically the data were collected to examine and compare the interactions and contamination potential between nearshore water and the shoreline sand and underlying sediment. 20160609 20170810 publication date Complete None planned -87.7961 -87.4310 41.6497 42.7517 ISO 19115 Topic Category farming USGS Thesaurus freshwater ecosystems water sampling polymerase chain reaction ecology microbiology DNA sequencing Coastal and Marine Ecological Classification Standard Microbial Communities USGS Metadata Identifier USGS:5da87ed2e4b09fd3b0c9c598 Getty Thesaurus of Geographic Names Lake Michigan Grand Calumet River Root River Canal No data access constraints No data use constraints Meredith Nevers U.S. Geological Survey, Great Lakes Science Center, Lake Michigan Ecological Research Station mailing

1574 Kemil Road N 300 E

Chesterton IN 46304 219-926-8336 x425 219-929-5792 mnevers@usgs.gov Great Lakes Restoration Initiative Bio-Rad CFX Manager 3.1 was used to obtain qPCR data Sequences were generated using a MiSeq Illumina system Sequences were analyzed using the QIIME pipeline (version 1.9.1) Meredith B. Nevers Muruleedhara N. Byappanahalli Cindy H. Nakatsu Julie L. Kinzelman Mantha S. Phanikumar Dawn A. Shively Ashley M. Spoljaric 202007 publication Water Research vol. 178 n/a Elsevier BV ppg. 115671 https://doi.org/10.1016/j.watres.2020.115671 Meredith B. Nevers Paul M. Buszka Muruleedhara N. Byappanahalli Travis Cole Steven R. Corsi P. Ryan Jackson Julie L. Kinzelman Cindy H. Nakatsu Mantha S. Phanikumar 202204 publication Journal of Great Lakes Research vol. 48, issue 2 n/a Elsevier BV ppg. 441-454 https://doi.org/10.1016/j.jglr.2022.01.009 High standard quality assurance practices were employed in laboratory and field protocols to collect the data, ensuring the accuracy of values in datasets. All technicians collecting data undergo rigorous training to adhere to SOPs. Appropriate positive and negative controls were included in all qPCR assays and inhibition determination took place on 20% of the samples. All standard curve R-squared values and amplification efficiencies were inspected, and all fell within the expected ranges; water sample data did not have expected ranges. Any questionable result was reanalyzed, if sample result was still questionable, the sample was not used in analysis. Datasets were checked for errors, including duplication or omission. All manually entered data were checked for errors against hard copies of the data. Data set is considered complete for the information presented, as described in the abstract. Users are advised to read the rest of the metadata record carefully for additional details. As noted in the attribute accuracy, if a data point was questionable after being reanalyzed the datum was removed. Coordinates were created from Google Earth (WGS84) A formal accuracy assessment of the vertical positional information in the data set has either not been conducted, or is not applicable. Andrew D. Eaton Lenore S. Clesceri Eugene W. Rice Arnold E. Greenberg Mary Ann H. Franson 2005 publication Washington, DC American Public Health Association ISBN: 0875530478 9780875530475 n/a Digital and/or Hardcopy 20051015 publication date American Public Health Association, 2005 Methodology used in processing step 1 Richard L. Whitman Meredith B. Nevers 200408 publication Environmental Science & Technology vol. 38, issue 16 n/a American Chemical Society (ACS) ppg. 4241-4246 https://doi.org/10.1021/es034978i Digital and/or Hardcopy 20040710 publication date Whitman and Nevers, 2004 Methodology used in processing step 1 World Health Organization 19990301 publication Geneva, Switzerland World Health Organization Technical document: Government Document number WHO/SDE/WSH/99.1 https://www.who.int/water_sanitation_health/bathing/annapolis.pdf Digital and/or Hardcopy 19990301 publication date World Health Organization, 1999 Methodology used in processing step 1 S. C. Edburg M. J. Allen D. B. Smith 19910501 publication Association of Official Analytical Chemists Volume 74, issue 3 ISSN: 0004-5756 n/a Digital and/or Hardcopy 19910501 publication date Edberg, Allen et al, 1991 Methodology used in processing step 2 Hyatt C. Green Richard A. Haugland Manju Varma Hana T. Millen Mark A. Borchardt Katharine G. Field William A. Walters R. Knight Mano Sivaganesan Catherine A. Kelty Orin C. Shanks 20140307 publication Applied and Environmental Microbiology vol. 80, issue 10 n/a American Society for Microbiology ppg. 3086-3094 https://doi.org/10.1128/AEM.04137-13 Digital and/or Hardcopy 20140317 publication date Green, H. C., et al, 2014 Methodology used in processing step 4 J. Lu J. W. Santo Domingo R. Lamendella T. Edge S. Hill 20080509 publication Applied and Environmental Microbiology vol. 74, issue 13 n/a American Society for Microbiology ppg. 3969-3976 https://doi.org/10.1128/AEM.00019-08 Digital and/or Hardcopy 20080626 publication date Lu, J. et al, 2008 Methodology used in processing step 4 John F. Griffith Blythe A. Layton Alexandria B. Boehm Patricia A. Holden Jennifer A. Jay Charles Hagedorn Charles D. McGee Steven B. Weisberg 20131201 publication Technical report 804 https://www.waterboards.ca.gov/water_issues/programs/beaches/cbi_projects/docs/sipp_manual.pdf Digital and/or Hardcopy 20131201 publication date Griffith, J. F., et al, 2013 Methodology used in processing step 4 J Gregory Caporaso Justin Kuczynski Jesse Stombaugh Kyle Bittinger Frederic D Bushman Elizabeth K Costello Noah Fierer Antonio Gonzalez Peña Julia K Goodrich Jeffrey I Gordon Gavin A Huttley Scott T Kelley Dan Knights Jeremy E Koenig Ruth E Ley Catherine A Lozupone Daniel McDonald Brian D Muegge Meg Pirrung Jens Reeder Joel R Sevinsky Peter J Turnbaugh William A Walters Jeremy Widmann Tanya Yatsunenko Jesse Zaneveld Rob Knight 20100411 publication Nature Methods vol. 7, issue 5 n/a Springer Nature ppg. 335-336 https://doi.org/10.1038/nmeth.f.303 Digital and/or Hardcopy 20100411 publication date Caporaso, J. G., et al, 2010 Methodology used in processing step 5 Jai Ram Rideout Yan He Jose A. Navas-Molina William A. Walters Luke K. Ursell Sean M. Gibbons John Chase Daniel McDonald Antonio Gonzalez Adam Robbins-Pianka Jose C. Clemente Jack A. Gilbert Susan M. Huse Hong-Wei Zhou Rob Knight J. Gregory Caporaso 20140821 publication PeerJ vol. 2 n/a PeerJ ppg. e545 https://doi.org/10.7717/peerj.545 Digital and/or Hardcopy 20140821 publication date Rideout, J. R., et al, 2014 Methodology used in processing step 5 Robert C. Edgar 20100812 publication Bioinformatics vol. 26, issue 19 n/a Oxford University Press (OUP) ppg. 2460-2461 https://doi.org/10.1093/bioinformatics/btq461 Digital and/or Hardcopy 20100812 publication date Edgar, R. C., et al, 2010 Methodology used in processing step 5 Daniel McDonald Morgan N Price Julia Goodrich Eric P Nawrocki Todd Z DeSantis Alexander Probst Gary L Andersen Rob Knight Philip Hugenholtz 20111201 publication The ISME Journal vol. 6, issue 3 n/a Springer Nature ppg. 610-618 https://doi.org/10.1038/ismej.2011.139 Digital and/or Hardcopy 20111201 publication date McDonald, D., et al, 2012 Methodology used in processing step 5 Q. Wang G. M. Garrity J. M. Tiedje J. R. Cole 20070622 publication Applied and Environmental Microbiology vol. 73, issue 16 n/a American Society for Microbiology ppg. 5261-5267 https://doi.org/10.1128/AEM.00062-07 Digital and/or Hardcopy 20070622 publication date Wang, Q., et al, 2011 Methodology used in processing step 5 Andre P Masella Andrea K Bartram Jakub M Truszkowski Daniel G Brown Josh D Neufeld 20120214 publication BMC Bioinformatics vol. 13, issue 1 n/a Springer Nature ppg. 31 https://doi.org/10.1186/1471-2105-13-31 Digital and/or Hardcopy 20120214 publication date Masella, Andre P., et al, 2012 Methodology used in processing step 5 Catherine Lozupone Manuel E Lladser Dan Knights Jesse Stombaugh Rob Knight 20100909 publication The ISME Journal vol. 5, issue 2 n/a Springer Nature ppg. 169-172 https://doi.org/10.1038/ismej.2010.133 Digital and/or Hardcopy 20100909 publication date Lozupone et al, 2011 Methodology used in processing step 5 The University of Texas at Austin, Bureau of Economic Geology 20170810 application/service http://www.beg.utexas.edu/coastal/thscmp/curriculum/sand%20exercise.htm Digital and/or Hardcopy 20170810 publication date Guidelines for Beach Grain Analysis Methodology used in processing step 3 Qiagen 201707 application/service For the isolation of genomic DNA from filtered water samples, including turbid water https://www.qiagen.com/us/resources/resourcedetail?id=bb731482-874b-4241-8cf4-c15054e3a4bf&lang=en Digital and/or Hardcopy 201707 publication date DNeasy PowerWater Methodology used in processing step 3 Qiagen 201705 application/service For the isolation of microbial genomic DNA from all soil types https://www.qiagen.com/us/resources/resourcedetail?id=5a0517a7-711d-4085-8a28-2bb25fab828a&lang=en Digital and/or Hardcopy 201705 publication date DNeasy PowerSoil Methodology used in processing step 3 Field Collections: Various studies with different study designs (e.g., sites, sampling frequency, replicate collection, substrates) were undertaken during the two year span (2016-2017) and are as follows: 2016: Individual water samples were collected 1 day a week (Thursday) for 10 weeks between 6/09/16 and 8/11/16. Samples were collected from two river and six shoreline locations. The two river locations are as follows: 1. Root River at the Root River Environmental Education Community Center (REC) 2. Grand Calumet River at Columbus Drive (GC) The six shoreline locations are as follows: 1. North Beach (NB1 and NB3), Racine, Wisconsin 2. 63rd Street Beach (63-1 and 63-2), Chicago, Illinois 3. Jeorse Park (JP1 and JP2) East Chicago, Indiana All water samples were analyzed for E. coli and host-specific microbial source tracking markers for gull (Gull2) and human (HF183) fecal sources. Additionally in 2016, triplicate water (all locations) and sediment and sand (shoreline locations) samples were collected during three separate events (6/21/16, 7/19/16, and 8/9/16); at Jeorse Park sampling locations included JP1 and an additional location, JPW (near the breakwall), in lieu of JP2. All samples were analyzed for E. coli and 16S rRNA metagenomics for determining bacterial communities. 2017: Individual water samples were collected 1 day (Thursday) a week for 10 weeks between 6/8/17 and 8/10/17 at REC, NB1, and NB3; 63-1 and 63-2; and GC, JP1, and JP2. During this time, sand and sediment samples were collected in triplicate, concurrently with water samples, 3 times (6/22/17, 7/20/17, and 8/10/17) at the shoreline locations. All samples were analyzed for E. coli and host-specific microbial source tracking markers for gull (Gull2) and human (HF183) fecal sources. Sampling techniques for the study were as follows. At each location water samples were collected using sterile Nalgene plastic bottles following protocols outlined in Standard Methods (American Public Health Association 2005), USGS protocols (Whitman and Nevers 2004), and Annapolis Protocol (World Health Organization 1999). Shoreline samples were collected in ~45 cm-deep water, 10cm below the surface, REC samples were collected from a boat launch using a reach pole or sampling line and GC samples were collected from an overhead bridge using a sterile bucket. Sediment collections took place at the same locations as water samples within a 0.5 m2 quadrat frame placed in a different position for each replicate. Within the frame, sub-samples were collected using a sterile, plastic core tube, open on both ends, by swiping along the bottom while covering half of the tube’s opening with the index finger. This allowed the sampler to collect approximately only the top 1 cm of sediment which was transferred into a whirlpack bag. Sand collections took place in the nearshore area, 50 cm away from the highest extension of the waves within a 0.5 m2 quadrat frame placed in a different position for each replicate. Within the frame, sub-samples were collected using sterile, stainless-steel shovels from the top 1-2 cm of surface sand and transferred into a whirlpack bag. Samples were transported to the laboratory in a cooler, on ice, and processed within 6 hr of collection. Citations: American Public Health Association. 2005. Standard Methods for the Examination of Water and Wastewater, 21st Edition. American Public Health Association, Washington, D.C. Whitman, R.L., and M.B. Nevers. 2004. Escherichia coli sampling reliability at a frequently closed Chicago beach: Monitoring and management implications. Environ. Sci. Technol. 38:4241-4246. World Health Organization. 1999. Health-based monitoring of recreational waters: The feasibility of a new approach (The "Annapolis Protocol") WHO/SDE/WSH/99.1. World Health Organization, Sustainable Development and Healthy Environments, Geneva. American Public Health Association, 2005 World Health Organization, 1999 Whitman and Nevers, 2004 20170810 Katarzyna Przybyla-Kelly U.S. Geological, Great Lakes Science Center, Lake Michigan Ecological Research Station Biological Technician mailing address

1574 Kemil Road N 300 E

Chesterton IN 46304 United States 219-926-8336 219-929-5792 kprzybyla-kelly@usgs.gov E. coli analysis: Water samples (generally 100 mL) and sand and sediment elutriate (generally 10-50-fold dilutions) were analyzed for E. coli using the IDEXX Colilert®-18 and Quanti-Tray® 2000 method (IDEXX Laboratories, Westbrook, Maine), a defined substrate technology (Edberg, Allen et al. 1991), with results provided as most probable number (MPN)/100 mL (water) or 1g (sand and sediment). For sand and sediment elutriation, 150-250 mL of phosphate buffer saline solution (PBS) was added to 100-200 g sand or sediment and contents shaken for 2 min on a wrist arm shaker. To determine moisture content of sand and sediment samples, about 10 g of fresh sample was placed in a 115°C drying oven for 24 hours and the weight differential was recorded. Citation: Edberg, S.C., M.J. Allen, and D.B. Smith. 1991. Defined substrate technology method for rapid and specific simultaneous enumeration of total coliforms and Escherichia coli from water: Collaborative study. J. Assoc. Off. Anal. Chem. 74:526-529. Edberg, Allen, et al. 1991 20170810 Dawn A Shively U.S. Geological, Great Lakes Science Center, Lake Michigan Ecological Research Station Biological Technician mailing address

1574 Kemil Road N 300 E

Chesterton IN 46304 United States 219-926-8336 x427 219-929-5792 dshively@usgs.gov Laboratory processing Filtration: All water samples (1000 mL) were pre-filtered through a 5.0 µm nitrocellulose filter followed by a 0.22 µm nitrocellulose filter (EMD Millipore, Billerica, MA), while 2017 sand and sediment elutriates (50 mL) were processed through only a 0.22 µm nitrocellulose filter. All filters were placed into separate PowerWater DNA Bead Tubes (DNeasy PowerWater Kit, Qiagen, Inc), and stored at -80°C until DNA extraction. Contamination blanks for laboratory processing or filtration protocol included method blanks (n=30), or 100 ml of sterile phosphate-buffered water (PBW; pH 7.0 ± 0.2) filtered through a 0.22 µm nitrocellulose filter each week (at each lab) and extracted alongside samples. DNA extraction: Genomic DNA was extracted from each 0.22 µm filter using the DNeasy PowerWater Kit (Qiagen, Inc.) according to instructions with one exception: the final DNA elution was performed twice using 50 μL of DNA elution buffer each time for a final extraction volume of 100 μL. To note, the 2016 water samples for 16S rRNA gene sequencing were co-extracted (both 5 and 0.22 µm) which resulted in supernatant being combined from both filters into one spin column and then treated as all other filters. DNA from the 2016 sand and sediment samples for 16S rRNA gene sequencing was extracted directly from 0.25 +/- 0.025 g of sample. The sample was thawed (stored in -20°C) and homogenized gently for 30 seconds. The subsample was aseptically placed into a PowerBead tube and extracted using the DNeasy PowerSoil Kit (Qiagen, Inc). The manufacturer protocol was followed with slight modifications to the 4°C incubation times, which were extended to 30 and 15 minutes each. Also, as with the PowerWater kit, the final elution step was completed twice using 50 uL of Solution C6 for a final volume of 100 uL. DNA concentration was quantified using a Qubit® dsDNA HS Assay Kit (Invitrogen, ThermoFisher Scientific, Waltham, MA), and quality (260/280 ratio) was measured using a Nanophotometer (Nanophotometer Pearl, Implen Inc, Westlake Village, CA). Extraction blanks (reagents only) were done alongside and throughout extracting DNA to ensure all reagents, techniques, and instruments were free of contamination (n=10). Sand fractionation: Sand and sediment samples were subjected to sand fractionation using a Keck Sand Shaker to determine percent fraction type (gravel, very coarse sand, coarse sand, medium sand, fine sand, very fine sand, and silt; clay was determined by a filtration technique); in 2016, triplicate samples were analyzed individually and in 2017, triplicate samples were homogenized for analysis. Process modified from (Guidelines for Beach Grain Analysis): http://www.beg.utexas.edu/coastal/thscmp/curriculum/sand%20exercise.htm. Guidelines for Beach Grain Analysis DNeasy PowerWater DNeasy PowerSoil 20170810 Ashley M Spoljaric U.S. Geological, Great Lakes Science Center, Lake Michigan Ecological Research Station Laboratory Technician mailing address

1574 Kemil Road N 300 E

Chesterton IN 46304 United States 219-926-8336 219-929-5792 aspoljaric@usgs.gov qPCR for Microbial Source Tracking markers: DNA extracts were used for the host-specific MST markers using the following primer/probe sets: Gull (Gull2, Lu et al., 2008; Griffith et. al., 2013): Forward: 5'- TGCATCGACCTAAAGTTTTGAG, Reverse: 5'- GTCAAAGAGCGAGCAGTTACTA, and TaqMan® probe: [6-FAM]-5'- CTGAGAGGGTGATCGGCCACATTGGGACT-BHQ1 and human (HF183, Green et. al., 2014): Forward: 5'- ATCATGAGTTCACATGTCCG, Reverse: 5'- CTTCCTCTCAGAACCCCTATCC, and TaqMan® probe: [6-FAM]-5'- CTAATGGAACGCATCCC-MGB. Quantitative polymerase chain reaction assays were performed to quantify gene copies associated with targeted fecal sources using real-time PCR platform (Bio-Rad CFX Connect™ Real-time PCR Detection System, Bio-Rad, Hercules, California) in 96-well PCR plates with a reaction volume of 25µl. For all assays, quantitation was determined from standard curves obtained from serial dilutions of gBlocks® Gene Fragments (Integrated DNA Technologies, Coralville, Iowa) specific to MST markers. Results for MST are reported as copy numbers (CN)/100 mL (water) or CN/g (sand and sediment); where copy number is the number of DNA molecules in a cell. DNA extracts were randomly chosen (20%) and analyzed diluted (5X) and undiluted for each qPCR assay to determine inhibition. Samples were considered inhibited if Cq values (diluted and undiluted) were dissimilar relative to dilution. Lower limit of quantification (LLOQ) was determined for each qPCR assay and applied to all corresponding data; this was the average of at least two of the triplicate Cq values detected at the lowest concentration of the standard curve, and high reproducibility was observed. All Cq values generated within 40 cycles were considered positive; a Cq < LLOQ is within the range of quantification (ROQ) and a Cq value > LLOQ was considered as detected but not quantifiable (DNQ) and results were reassigned a quantity of half of the LLOQ (CN/rx = X, HF183 and CN/rx = X, Gull2) and all non-detect (ND) samples were reassigned a quantity of 1/4 of the LLOQ (CN/rx = X): for HF183=4 in 2016 and 5 in 2017 and for Gull2=8 in both 2016 and 2017. Citations: Griffith, J. F., et al. (2013). The California Microbial Source Identification Manual: A Tiered Approach to Identifying Fecal Pollution Sources to Beaches, Technical Report 804, Southern California Coastal Water Research Project, Costa Mesa, CA. Lu, J., J.W. Santo Domingo, R. Lamendella, T. Edge, and S. Hill. 2008. Phylogenetic diversity and molecular detection of bacteria in gull feces. Appl. Environ. Microbiol. 74:3969-3976. Green, H. C., et al. (2014). "Improved HF183 quantitative real-time PCR assay for characterization of human fecal pollution in ambient surface water samples." Applied and Environmental Microbiology 80(10): 3086-3094. Gentry-Shields, J. R., Jakob G, Stewart, Jill R (2012). "HuBac and nifH source tracking markers display a relationship to land use but not rainfall." Water Research 46(18): 6163-6174. Green, H. C., et al, 2014 Lu, J., et al., 2008 Griffith, J. F., et al, 2013 20170810 Ashley M Spoljaric U.S. Geological, Great Lakes Science Center, Lake Michigan Ecological Research Station Laboratory Technician mailing address

1574 Kemil Road N 300 E

Chesterton IN 46304 United States 219-926-8336 219-929-5792 aspoljaric@usgs.gov Illumina 16S rRNA Gene Sequencing: PCR-amplification of the 16S rRNA gene in water and sand DNA extracts was performed using V3-V4 region primers (343-forward TAC GGR AGG CAG CAG and 804-reverse CTA CCR GGG TAT CTA ATC C). PCR protocol and primers tags were following the manufacturer’s suggested step out protocol (Illumina, San Diego, CA). Reactions were carried out using ~10 ng of template DNA in Q5® High Fidelity DNA Polymerase 2X master mix (New England Biolabs). PCR amplicons were purified using AxyPrepMag PCR clean-up kit (Axygen Scientific) and quantified using a Nanodrop 3000 fluorospectrophotometer after staining with the QuantiFluor dsDNA System (Promega). Amplicon were combined in equimolar amounts sequenced (2x250 paired end) on a MiSeq Illumina instrument at the Purdue Genomics core facilities. 16S rRNA gene sequence analysis: Sequences with primer tags removed and paired end reads merged (PANDAseq software), (Masella et al., 2012) were analyzed using the QIIME pipeline (version 1.9.1) (Caporaso et al., 2010). The options in QIIME chosen were “pick_open_otus” (Rideout et al., 2014), uclust (Edgar, 2010) for clustering, PyNAST (Caporaso et al., 2010) for sequence alignment, and RDP classifier (Wang et al., 2007) for taxonomic assignment using the Greengenes data set (version 13_5) (McDonald et al., 2012). Rarefied datasets (lowest number of reads among the samples being compared) were used for beta-diversity comparisons. Metrics tested included both phylogenetic Unifrac distances (weighted and un-weighted) (Lozupone et al., 2011) and non-phylogenetic distances (weighted Jaccard and Bray Curtis). Good’s coverage to assess completeness of OTU representation in each sample. Alpha-diversity measurements were used for richness and evenness (Shannon diversity), richness (ChaoI index, observed-species), Faith’s phylogenetic diversity (PD whole tree) and evenness (Simpson’s e). The raw metagenomic data can be accessed at the NCBI repository under the biproject accession PRJNA578544: https://www.ncbi.nlm.nih.gov/sra/PRJNA578544. References: Masella, Andre P., Andrea K. Bartram, Jakub M. Truszkowski, Daniel G. Brown, and Josh D. Neufeld. "PANDAseq: paired-end assembler for illumina sequences." BMC bioinformatic. 2012, 13 (1): 31. Caporaso, J. G.; Kuczynski, J.; Stombaugh, J.; Bittinger, K.; Bushman, F. D.; Costello, E. K.; Fierer, N.; Pena, A. G.; Goodrich, J. K.; Gordon, J. I.; Huttley, G. A.; Kelley, S. T.; Knights, D.; Koenig, J. E.; Ley, R. E.; Lozupone, C. A.; McDonald, D.; Muegge, B. D.; Pirrung, M.; Reeder, J.; Sevinsky, J. R.; Turnbaugh, P. J.; Walters, W. A.; Widmann, J.; Yatsunenko, T.; Zaneveld, J.; Knight, R., QIIME allows analysis of high-throughput community sequencing data. Nat. Meth. 2010, 7 (5), 335-336. Rideout, J. R.; He, Y.; Navas-Molina, J. A.; Walters, W. A.; Ursell, L. K.; Gibbons, S. M.; Chase, J.; McDonald, D.; Gonzalez, A.; Robbins-Pianka, A.; Clemente, J. C.; Gilbert, J. A.; Huse, S. M.; Zhou, H.-W.; Knight, R.; Caporaso, J. G., Subsampled open-reference clustering creates consistent, comprehensive OTU definitions and scales to billions of sequences. PeerJ 2014, 2, e545. Edgar, R. C., Search and clustering orders of magnitude faster than BLAST. Bioinformatics 2010, 26 (19), 2460-2461. McDonald, D.; Price, M. N.; Goodrich, J.; Nawrocki, E. P.; DeSantis, T. Z.; Probst, A.; Andersen, G. L.; Knight, R.; Hugenholtz, P., An improved Greengenes taxonomy with explicit ranks for ecological and evolutionary analyses of bacteria and archaea. ISME J. 2012, 6 (3), 610-618. Wang, Q.; Garrity, G. M.; Tiedje, J. M.; Cole, J. R., Naive Bayesian classifier for rapid assignment of rRNA sequences into the new bacterial taxonomy. Appl. Environ. Microbiol. 2007, 73 (16), 5261-5267. Lozupone, Catherine, Manuel E. Lladser, Dan Knights, Jesse Stombaugh, and Rob Knight. "UniFrac: an effective distance metric for microbial community comparison." The ISME journal. 2011, 5 (2) : 169. Masella, Andre P., et al, 2012 Caporaso, J. G., et al, 2010 Rideout, J. R., et al, 2014 Edgar, R. C., et al, 2010 McDonald, D., et al, 2012 Wang, Q., et al, 2011 Lozupone et al., 2011 20160809 Cindy H. Nakatsu Purdue University, Department of Agronomy mailing

915 W State Street

Lafayette Indiana 47907 USA 765-496-2997 cnakatsu@purdue.edu Coordinates.csv This table contains information associated with sampling locations, including latitude and longitude. Producer defined Location_ID shortened location name representing the sampling locations Producer defined Indicates the nine sampling locations for the study: REC, NB3 and NB1, 63-1 and 63-2, GC, and JPW, JP1, and JP2 Location_name Formal location name Producer defined Formal location names for the sampling locations. Rivers: REC: Root River at the Root River Environmental Education Community Center, Racine, WI GC: Grand Calumet River at Columbus Drive, East Chicago, Indiana Shoreline locations: NB3 and NB1: North Beach, Racine, WI 63-1 and 63-2: 63rd Street Beach, Chicago, Illinois JPW, JP1, and JP2: Jeorse Park, East Chicago, Indiana. Latitude Latitude in decimal degrees (WGS84) Producer defined Latitude for each location obtained from Google Earth Longitude Longitude in decimal degrees (WGS84) Producer defined Longitude for each location obtained from Google Earth E coli.csv This table contains E. coli densities for each location. Producer defined Date Field sample collection date Producer defined Field sample collections took place: June 09 through August 11, 2016 June 08 through August 10, 2017 Substrate Type of substrate from which samples were collected Producer Defined Substrates are: Water: Water collected from Lake Michigan (shoreline locations), Grand Calumet River, and Root River Sediment: Lake bottom sand collected beneath lake water samples at shoreline locations Sand: Beach sand collected in nearshore area, 50 cm away from the highest extension of the waves at shoreline locations Location_ID Sampling locations Producer defined See table Coordinates.csv for specific information on these sampling locations. Replicate Replicate number Producer defined NA Individual sample, no replicate collected Producer defined During sample collections, individual (NA) or triplicate water, sand, and sediment samples (1-3) were collected at each location, which was dependent upon study period and site. Ecoli Culture-based E. coli measurements. If the data in the substrate type column is water, the units of measure are MPN/100 mL, whereas if the substrate type column indicates sand or sediment, the units of measure are MPN/1 g. Producer defined 0.23 23396.40 Comments Additional sample information Producer defined Pertinent information regarding E. coli detection MST.csv This table contains data related to qPCR assay results. Producer defined Date Field sample collection date Producer defined Field sample collections took place: June 09 through August 11, 2016 June 08 through August 10, 2017 Substrate Type of substrate from which samples were collected Producer Defined Substrates are: Water: Water collected from Lake Michigan (shoreline locations), Grand Calumet River, and Root River Sediment: Lake bottom sand collected beneath lake water samples at shoreline locations Sand: Beach sand collected in nearshore area, 50 cm away from the highest extension of the waves at shoreline locations Location_ID Sampling locations Producer defined See table Coordinates.csv for specific information on these sampling locations Replicate Replicate number Producer defined NA Individual sample, no replicate Producer defined During sample collections, individual (NA) or triplicate water, sand, and sediment samples (1-3) were collected at each location, which was dependent upon study period and site. Assay Target host-specific DNA marker Producer defined Gull2: Gull-specific HF183: Human-specific SQ Starting quantity Producer defined ND ND means "not detected" and refers to no target DNA detected in the qPCR assay. Producer defined Initial amount of DNA present in each sample per amount of genomic DNA added to reaction mixture. Detection DNA marker detection status Producer defined DNQ: Detected but not quantifiable (below the limit of quantification) ND: Not detected ROQ: Detected within the range of quantification CN Amount of target DNA present in DNA copy numbers. For water samples: CN/100 mL=SQ* (100 uL DNA eluted/2 uL DNA per rx)*(100 mL/1000 mL filtered). For sand and or sediment samples: CN/1 g=SQ* (100 uL DNA eluted/2 uL DNA per rx)*dry weight ratio. For statistical analysis purposes, samples that yielded DNQ results were reassigned a quantity of half of the LLOQ (CN/rx = X): for HF183=9 in 2016 and 10 in 2017 and for Gull2=17 in both 2016 and 2017 and ND samples were reassigned a quantity of 1/4 of the LLOQ (CN/rx = X): for HF183=4 in 2016 and 5 in 2017 and for Gull2=8 in both 2016 and 2017. If the data in the substrate type column is water, the units of measure are CN/100 mL, whereas if the substrate type column indicates sand or sediment, the units of measure are CN/1 g. Producer defined 4 202050 Fractionation.CSV This table contains sand and sediment fractionation results. Producer Defined Date Sample collection date Producer Defined Sand and sediment were collected during 3 events each year 2016: 6/21/16, 7/19/16, and 8/9/16 2017: 6/22/17, 7/20/17, and 8/10/17 Location_ID Sampling locations Producer Defined See table Coordinates.csv for specific information on these locations. Substrate Type of substrate from which samples were collected Producer Defined Substrates are: Sediment: Lake bottom sand collected beneath lake water samples at shoreline locations Sand: Beach sand collected in nearshore area, 50 cm away from the highest extension of the waves at shoreline locations Replicate Replicate number Producer Defined NA Triplicate samples homogenized to one individual sample Producer defined 2016: Triplicate sand and sediment samples were collected at each location. 2017: Triplicate sand and sediment samples were collected at each location and in the lab, each replicate sample was homogenized and sub-samples (75g) from each replicate were combined. Gravel Gravel fraction (> 2 mm) at each location Producer Defined 0.0144490773232 99.5272698579 Percent VeryCoarseSand Very coarse sand fraction (1–2 mm) at each location Producer Defined 0.0 32.8133933676 Percent CoarseSand Coarse sand fraction (0.5–1 mm) at each location Producer Defined 0.038736612555 75.5745590593 Percent MediumSand Medium sand fraction (0.25–0.5 mm) at each location Producer Defined 0.0130302147686 97.7870808427 Percent FineSand Fine sand fraction (0.125-0.25 mm) at each location Producer Defined 0.00843131543851 29.8449174561 Percent VeryFineSand Very fine sand fraction (0.0625-0.125 mm) at each location Producer Defined 0.000564639985545 3.51688291476 Percent Silt Silt fraction (0.0039-0.0625 mm) at each location Producer Defined 0.0 4.04812238772 Percent Clay Clay fraction (< 0.0039 mm) at each location Producer Defined 0.0 0.372181923949 Percent GS ScienceBase U.S. Geological Survey mailing address

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