Deborah A Repert Isiah R Castro Mark M Dornblaser James J Duncker David J Fazio Jennifer B. Sharpe Richard L Smith Kimberly Wickland 20250219 tabular digital data Boulder, CO U.S. Geological Survey https://doi.org/10.5066/P13DSK7B Data on carbon and nutrient processing in key locations and over seasonal time scales can provide critical information about the concentrations and potential utilization rates of bioavailable constituents that are known to have a role in cyanobacteria harmful algal blooms (cyanoHABs). Laboratory experiments were performed to measure seasonal nutrient processing rates of carbon, nitrogen and phosphorus in river water and bottom sediments collected just upstream of the Next Generation Water Observing System (NGWOS) location at Starved Rock Lock and Dam USGS gaging station number 05553700 on the Illinois River. This site was chosen based on previous development of cyanoHABs. Sediments and unfiltered river water were collected during the spring (April), summer (August), and fall (November) of 2022 then shipped to Boulder, CO for laboratory experiments. River water alone and river water combined with < 2-millimeter sieved sediments were assessed under ambient nutrient conditions and nitrogen amended conditions to determine rates of production or consumption for the following processes: 1) aerobic carbon dioxide production, oxygen consumption, and aqueous nutrient transformations, and 2) anaerobic denitrification, carbon dioxide production, methanogenesis, and aqueous nutrient transformations. Modifications to certain experiments were made in November 2022 based on results from the April and August 2022 experiments and are noted in the metadata steps. Background sediment total carbon and nitrogen content, deionized water and potassium chloride extractable sediment nutrient concentrations, and river water nutrient concentrations were measured during each collection. Sequential extractions of the sieved sediment were performed using deionized water, sodium bicarbonate, sodium hydroxide, and hydrochloric acid to measure phosphorus content of the sediment during the various steps. The purpose of the data release is to provide data on nutrient cycling processing rates that can be applied to various models to assess potential drivers of cyanobacteria harmful algal blooms. 20220411 20230518 ground condition Complete None planned World -89.02393 -88.96454 41.34357 41.30051 ISO 19115 Topic Category imageryBaseMapsEarthCover biota None nitrogen carbon phosphorus USGS Metadata Identifier USGS:6658923ed34ef3137d35f887 None Starved Rock Lock and Dam Illinois River Basin Public Any use of trade, firm, or product names is for descriptive purposes only and does not imply endorsement by the U.S. Government. Although this information product, for the most part, is in the public domain, it also contains copyrighted materials as noted in the text. Permission to reproduce copyrighted items must be secured from the copyright owner whenever applicable. The data have been approved for release and publication by the U.S. Geological Survey (USGS). Although the data have been subjected to rigorous review and are substantially complete, the USGS reserves the right to revise the data pursuant to further analysis and review. Furthermore, the data are released on the condition that neither the USGS nor the U.S. Government may be held liable for any damages resulting from authorized or unauthorized use. Although the data have been processed successfully on a computer system at the U.S. Geological Survey, no warranty expressed or implied is made regarding the display or utility of the data on any other system, or for general or scientific purposes, nor shall the act of distribution constitute any such warranty. The U.S. Geological Survey shall not be held liable for improper or incorrect use of the data described and/or contained herein. Users of the data are advised to read all metadata and associated documentation thoroughly to understand appropriate use and data limitations. Deborah A Repert U.S. Geological Survey Microbiologist mailing and physical

3215 Marine St. E127

Boulder CO 80303 USA 303-541-3020 darepert@usgs.gov Funding for this project was provided by the U.S. Geological Survey (USGS) Water Mission Area Water Quality Processes program and the USGS Next Generation Water Observing System (NGWOS) initiative. We are grateful to Paul Bliznik, Josie Marquez, David Roth, and Elizabeth Tomaszewski for helping with lab experiments and providing analytical support. Quality Assurance and Quality Control For dissolved organic carbon (DOC) analyses, a six-point curve calibration curve was constructed. The curve was a linear fit with a zero shift, and a successful calibration was achieved with r2 > 0.995. Two deionized water (DIW) blanks follow each calibration curve and after every 3 samples to ensure minimal carry over. Check standards (10 milligrams per liter (mg/L) and 20 mg/L DOC) were analyzed consistently throughout the run to ensure that the instrument was not drifting. In addition, U.S. Geological Survey standard reference water samples (SRWS) (http://bqs.usgs.gov/srs/) and Environment Canada standards (https://nrc.canada.ca/en/certifications-evaluations-standards/certified-reference-materials/list) were used to check the analytical methods. For ion chromatograph (IC) analyses of major anions, including nitrate and nitrite, and major cations, including ammonium, standards purchased from High Purity Standards (HPS, https://highpuritystandards.com) were used to construct low and high range calibration curves. The anions and ammonium standard curves were fit with a quadratic equation, major cations were fit with a linear equation, and successful calibration was achieved with r2 > 0.995. To ensure the accuracy and precision of the instrument DIW blanks, SRWSs, check standards, and check samples, including a Thermo Scientific DionexTM Five Anion combined standard and DionexTM Six Cation-II combined standard, were analyzed periodically throughout each run. For discrete analyses of ammonium, nitrate plus nitrite, nitrite, and phosphate standards purchased from HPS were used to construct calibration curves. All curves were fit with a linear equation, and successful calibration was achieved with r2 > 0.995. To ensure the accuracy and precision of the instrument DIW blanks, SRWSs, check standards, and check samples, including a Thermo Scientific DionexTM Five Anion combined standard and DionexTM Six Cation-II combined standard, were analyzed periodically throughout each run. For TDN analyses using the Skalar FormacsTN analyzer, standards purchased from HPS were used to construct five-point and eight-point calibration curves. Calibration standards were acidified with 6 N HCl to match the matrix of the preserved samples. The curves were fit with a linear equation and successful calibration was achieved with r2 > 0.995. To ensure the accuracy and precision of the instrument, acidified DIW blanks, SRWSs, check standards, and check samples (which included glutamic acid and nicotinic acid) were analyzed periodically throughout each run. For metal analyses using inductively coupled plasma optical emission spectroscopy (ICP-OES), DIW blanks and analysis of SRWSs were utilized. Blanks were analyzed along with the samples as unknowns. Lab blanks were unfiltered and preserved in the same manner as the samples. Along with blanks, several SRWSs were analyzed as unknowns multiple times during each analytical run to check for accuracy. The National Institute of Standards and Technology (NIST) standard NIST1643d (NIST, 1999) was used to check the accuracy of trace metals determined by ICP-OES. For CHN analysis, an acetanilide standard was used to create a single point calibration then replicates run every 9-12 injections as unknowns to verify the calibration. Empty nickel sleeves plus tin capsules were used as instrument blanks. An Ottawa sand standard (20-30 mesh) was used as a sample blank and run every 9-12 injections. To ensure the accuracy and precision of the instrument a National Institute of Standards and Technology standard reference material 1216 carbon modified silica was analyzed at the beginning and end of each run (U.S. Dept. of Commerce, 2012). For gas analyses, nitrous oxide standard curves were generated using certified standards purchased from Air Liquide. The curves were a fit with a quadratic equation when using an HNU Gas Chromatograph (GC) for the April and August 2022 experiments and a linear equation fit when using a Shimadzu GC for the November experiments. Successful calibration was achieved with r2 > 0.995. Carbon dioxide and methane standard curves were generated using certified standards purchased from Praxair. The curves were fit with a linear equation and successful calibration was achieved with r2 > 0.995. Standards for all gas analyses were run periodically to ensure stability of the calibration curve. Data are consistent and fall within expected ranges 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. No formal positional accuracy tests were conducted, but latitude and longitude measurements are expected to be within 10 meters of the sample site. No formal positional accuracy tests were conducted. Sediment and river water collection, processing, and characterization: River bottom sediments were collected just offshore and upstream of the dam at the Starved Rock location where sediments pool, but water still flushes through, using a shovel to remove the top approximately 0- to 10-centimeter (cm) of bottom material. Several collections of the sediment material were made and deposited in a 2-gallon acid-washed bucket. The bucket was filled completely with sediment and unfiltered (UF) river water (RW), sealed tightly, then shipped on ice overnight to USGS in Boulder, CO. Upon receipt in Boulder, the sediment was mixed well with a large, autoclaved spoon and sieved through a 2‐millimeter (mm)‐mesh polypropylene screen. Sediment subsamples of the < 2 mm sized fraction were collected and preserved for percent moisture, carbon, and nitrogen content by drying at 50 degrees Celsius (deg C), for phosphorus extractions by air drying or freezing at -20 deg C, and for potential future 16s DNA sequencing by freezing at -50 deg C. The remaining < 2 mm sized fraction was retained in baked glass containers and used for preparing oxic and anoxic experiments as described in process steps 2 and 3. UF RW for the laboratory experiments was collected from the same location at the same time as the sediment, stored in 6 clean, field-rinsed 2-liter (L) plastic bottles and shipped on ice overnight to Boulder, CO. Upon receipt in Boulder, subsamples of the UF RW were analyzed for pH and specific conductance (SpC). UF RW subsamples were filtered through a DIW rinsed 0.22 micrometer (µm) Millipore SterivexTM PVDF filter and preserved for analysis of major anions (including nitrate (NO3), nitrite (NO2), and phosphate (PO4)) by freezing (-20 deg C), major cations (including ammonium (NH4)) by acidification with 18-normal sulfuric acid (2 microliters (µL) acid per 1 milliliter (mL) sample), dissolved organic carbon (DOC) and ultraviolet absorbance (UVAbs) at 254 nanometers (nm) by chilling (4 deg C), total dissolved nitrogen (TDN) by acidification with 6-normal hydrochloric acid (3 µL acid per mL sample), trace metals by acidification (1 mL sample plus 9 mL 1% nitric acid), and alkalinity by chilling (4 deg C). The filter was frozen and archived at -50 deg C for potential future 16s dna sequencing. Measurements were made at the field site location at the time of water collection for pH, specific conductance (SpC), dissolved oxygen (DO) concentration, water temperature (T), barometric pressure (BP), turbidity, total algae phycocyanin, and chlorophyll using a YSI EXO2 handheld multiparameter sonde in April and November 2022. Data recorded by sensors located downstream at the USGS Next Generation Water Observing System (NGWOS) Illinois River at Starved Rock gaging station (05553700) were retrieved in April, August, and November 2022 from https://waterdata.usgs.gov/monitoring-location/05553700/#dataTypeId=continuous-00010-82063&period=P7D&showMedian=true. YSI EXO2 data were not available in August 2022 thus field parameters from downstream sensors only were reported for that collection. To prepare water for anoxic experiments, UF RW and DIW were transferred to autoclaved 2 L glass media bottles, sealed with caps modified with blue butyl rubber stoppers and tubing extending through stoppers to the bottom of the bottles, then deoxygenated by bubbling with argon (Ar) gas while stirring for several hours (h). The stoppered caps were then quickly replaced with solid caps and the bottles transferred to an anaerobic chamber (nitrogen, hydrogen, and carbon dioxide atmosphere) overnight. The following morning, the bottles were opened in the anaerobic chamber, stirred to remove residual oxygen, and the anoxia of the water confirmed using CHEMetrics oxygen 3 ampoules. To determine background available nutrients loosely bound to the sediment, extractions were performed on the sieved sediment using 2 molar (M) potassium chloride (KCl) and DIW. For the KCl extractions, 30 grams (g) wet sediment was combined with 90 g 2 M KCl, in triplicate 250 milliliter (mL) centrifuge bottles. For the DIW extractions, 50 g wet sediment was combined with 100 mL DIW, in triplicate 250 mL centrifuge bottles. Control blanks were prepared containing 2 N KCl and DIW only. Bottles were placed on a reciprocal shaker for 2 h at 180 revolutions per minute (rpm) then centrifuged at 10,000 rpm for 8 minutes (mins). The KCl extraction samples were filtered (0.45 µm 25 mm Whatman GD/X glass microfiber syringe filter) and preserved as previously described for NH4 and NO3 plus NO2 analysis. The DIW extraction samples were filtered and preserved as previously described for major cations, major anions, trace metals, and DOC analysis. 20221118 Deborah A Repert USGS - WATER Microbiologist mailing and physical

Boulder NRP,University of Colorado - Building 6

Boulder CO 80303 303-541-3020 darepert@usgs.gov Oxic Experiments: The below experiments were initiated within 0-48 hours of receiving the water and sediment in the Boulder, CO laboratory. 1) Oxic Aqueous-phase Nutrients: Rates of ammonium oxidation to nitrate (nitrification) were measured in 250 mL centrifuge bottles containing the following treatments: 1) 60 g well-mixed < 2 mm sieved wet sediment plus 90 mL oxic UF RW, in triplicate per timepoint (tp), 2) 60 g well-mixed < 2 mm sieved wet sediment plus 90 mL oxic UF RW amended to 250 micromolar (µM) NH4, in triplicate per tp, 3) 90 mL oxic UF RW, in duplicate per tp, 4) 90 mL oxic UF RW amended to 250 µM NH4, in duplicate per tp, and 5) 90 mL oxic DIW, 1 replicate per tp. Bottles were mixed on an end-over-end rotator at room temperature (20-22 deg C) and serially sacrificed after approximately 0.1, 1, 2, 3, and 5 day-time points. Five mL of sample was removed from each bottle for pH and SpC measurements. Bottles containing sediment were allowed to settle for 5 mins prior to this step then centrifuged at 10,000 rpm for 8 mins. The supernatant was filtered through a 0.22 µm Sterivex filter and preserved for analysis of major anions, major cations, DOC, UVabs, and TDN as previously described. Remaining sediment in the sediment plus RW bottles was mixed with a sterile spatula, 4-5 g added to a sterile 5 mL plastic vial, then frozen at -50 deg C. Nutrient concentrations (milligrams per liter) over time were plotted and the linear portion of the curves used to determine rates of nitrate, ammonium, and phosphate production or reduction, standard error (SE) of the slopes, and coefficients of variation (R squared). 2) Oxygen Consumption: Rates of oxygen consumption were measured in 60 mL serum bottles with attached oxygen sensors and contained the following treatments: 1) 20 g well-mixed < 2 mm sieved wet sediment plus 30 mL oxic UF RW, in triplicate, 2) 20 g well-mixed < 2 mm sieved wet sediment plus 30 mL oxic UF RW amended to 250 µM NH4, in triplicate, 3) 30 mL oxic UF RW, in duplicate, 4) 30 mL oxic UF RW amended to 250 µM NH4, in duplicate, and 5) 30 mL oxic DIW, 1 rep only. Bottles were stoppered with blue butyl rubber stoppers, crimped, and an additional 25 mL of air added to each bottle using a 30 mL syringe and 22 gauge (G) needle to ensure sufficient oxygen availability. Bottles were mixed on an end over end rotator at room temperature (20-22 deg C) and measured for oxygen concentration and temperature 8 - 10 times over the course of approximately 1 week using a PreSens Fibox 4 trace oxygen meter plus Sensor spot SP-PSt3 plus polymer optical fiber (https://www.presens.de/products/o2/featured-o2-systems/fibox-4-trace-sp-pst6-pof). Oxygen concentration (micromoles per liter) over time were plotted and the linear portion of the curve used to determine rate of production, standard error (SE) of the slope, and coefficient of variation (R squared). 3) Oxic Carbon Dioxide production: Rates of carbon dioxide (CO2) production were measured in 60 mL serum bottles containing the following treatments: 1) 20 g well-mixed < 2 mm sieved wet sediment plus 30 mL oxic UF RW, in triplicate, 2) 20 g well-mixed < 2 mm sieved wet sediment plus 30 mL oxic UF RW amended to 250 µM NH4, in triplicate, 3) 30 mL oxic UF RW, in duplicate, 4) 30 mL oxic UF RW amended to 250 µM NH4, in duplicate, and 5) 30 mL oxic DIW, 1 replicate only. Bottles were stoppered with blue butyl rubber stoppers, crimped, and an additional 25 mL of air was added to each bottle using a 30 mL syringe and 22 G needle to generate an internal positive pressure for sampling. Bottles were mixed on an end over end rotator at room temperature (20-22 deg C) in between samplings. Headspace gas samples were removed daily for 4-5 days using a gastight syringe plus 22 G needle, injected into a LICOR Model LI-6262 infrared CO2 analyzer with nitrogen carrier gas, and analyzed for CO2 concentration daily for 5 days. At the end of the experiment, the pressure remaining in the bottle was measured using a Crystal Model 31 Pressure Calibrator with attached 22 G needle. The bottle was then opened, and 5 mL of liquid removed for pH measurement. Concentration (micromole per liter) over time was plotted and the linear portion of the CO2 gas concentration curve used to determine CO2 production rates. Carbon dioxide concentration (micromoles per liter) over time were plotted and the linear portion of the curve used to determine rate of production, standard error (SE) of the slope, and coefficient of variation (R squared). *Note: Due to a shortage of lab personnel to assist in November 2022 some adjustments were made to these experiments. Results from April and August 2022 indicated no significant difference in CO2 production rates for the unamended UF RW + Sed and the UF RW + Sed + NH4 treatments, thus the unamended UF RW + Sed + NH4 treatment was omitted in November 2022. In addition, the DIW and UF RW + NH4 treatments were omitted given the very low or negligible rates of CO2 production during the incubation time period in April and August 2022. The UF RW treatment was retained as a control. 4) Oxic Dissolved Inorganic Carbon: To determine dissolved inorganic carbon (DIC) change over the course of the oxic experiments, 60 mL serum bottles were prepared containing the following treatments: 1) 20 g well-mixed < 2 mm sieved wet sediment plus 30 mL oxic UF RW, duplicates for beginning (T0 – time zero) and end (Tf – time final), 2) 30 mL oxic UF RW, duplicates for T0 and Tf, and 3) 30 mL oxic DIW, 1 replicate for T0 and Tf. Bottles were stoppered, crimped, 25 mL additional air added, then mixed on an end-over-end rotator at room temperature (20-22 deg C). After an equilibration period of approximately 2-3 h, T0 samples were collected from the stoppered serum bottle by carefully removing approximately 15 mL liquid using a 30 mL syringe and 18 G inch needle. The needle was removed and a 0.45 µm filter with attached 22 G needle was connected to the syringe. The contents were then filtered into a previously prepared stoppered, crimped, Ar-flushed 30 mL serum bottle containing 400 µL 18 N sulfuric acid to convert all aqueous DIC to CO2. Sample bottles were analyzed at a later date for headspace CO2 concentration as previously described. The pH was measured on the remaining water and sediment in the original 60 mL serum bottle. This procedure was repeated at Tf, corresponding to the end of the oxic respiration experiments. *Note: DIC change was measured in August and November 2022 only. 20221118 Deborah A Repert USGS - WATER Microbiologist mailing and physical

Boulder NRP,University of Colorado - Building 6

Boulder CO 80303 303-541-3020 darepert@usgs.gov Anoxic Experiments: The below experiments were initiated within 24-48 hours of receiving the water and sediment in the Boulder, CO laboratory. 1) Anoxic Aqueous-phase Nutrients: Rates of nutrient change in the aqueous phase were measured in 125 mL serum bottles containing the following treatments: 1) 60 g well-mixed < 2 mm sieved wet sediment plus 90 mL anoxic UF RW, in triplicate per tp, 2) 60 g well-mixed < 2 mm sieved wet sediment plus 90 mL anoxic UF RW amended to 500 µM NO3, in triplicate per tp, 3) 90 mL anoxic UF RW, in duplicate, 4) 90 mL anoxic UF RW amended to 500 µM NO3, in duplicate per tp, and 5) 90 mL anoxic DIW. Sediment was weighed into the bottles aerobically on the lab bench in small sets then transferred to an anaerobic chamber immediately to minimize air exposure and anoxic RW or DIW added. Bottles were stoppered, removed from the anaerobic chamber, crimped, flushed with Ar for 15-20 mins then an anoxic 10 mM NO3 stock solution was added to appropriate bottles using a 5 mL syringe and 22 G needle. The volume of nitrate stock solution added was adjusted seasonally to account for the nitrate initially present in the RW and to achieve a final concentration of 500 µM in the sample bottles. Bottles were mixed on an end-over-end rotator at room temperature (20-22 deg C) and serially sacrificed after approximately 0.1, 1, 2, 3, and 5 days. Five mL of sample was removed from each bottle for pH and SpC measurements. Bottles containing sediment were allowed to settle for 5 mins prior to this step then transferred to a 250 mL centrifuge bottle and centrifuged at 10,000 rpm for 8 mins. Supernatant was filtered through a 0.22 µm Sterivex filter and preserved for analysis of major anions, major cations, DOC/UV, and TDN as previously described. The remaining sediment in the sediment plus RW bottles was mixed with a sterile spatula, 4-5 g added to a sterile 5 mL plastic vial, then frozen at -50 deg C. A second set of bottles were prepared the following day for sampling the aqueous phase on a shorter timescale to capture the rapid nitrate reduction rates. For this bottle set, rates of nutrient change in the aqueous phase were measured in 60 mL serum bottles containing the following treatments: 1) 20 g well-mixed < 2 mm sieved wet sediment plus 30 mL anoxic UF RW, in triplicate per tp, and 2) 30 mL anoxic UF RW, T0 and Tf only. Bottles were prepared as previously described for the original bottle set but sampled after approximately 1, 2, 3, and 4 h for major anions and major cations only. For both the long-term and short-term experiments, nutrient concentrations (milligrams per liter) over time were plotted and the linear portion of the curves used to determine rates of nitrate, ammonium, and phosphate production or reduction, standard error (SE) of the slopes, and coefficients of variation (R squared). *Note: Nitrate was accidentally omitted from the first timepoint for the UF RW + NO3 aqueous sampling only bottle during the April 2022 experiment. 2) Anoxic Denitrification: Rates of nitrate reduction were measured in 60 mL serum bottles containing the following treatments: 1) 20 g well-mixed < 2 mm sieved wet weight sediment plus 30 mL anoxic UF RW, in triplicate, 2) 20 g well-mixed < 2 mm sieved wet weight sediment plus 30 mL anoxic UF RW amended to 500 µM NO3, in triplicate, 3) 30 mL anoxic UF RW, in duplicate, 4) 30 mL anoxic UF RW amended to 500 µM NO3, in duplicate, and 5) 30 mL anoxic DIW. Sediment was weighed into the bottles aerobically on the lab bench in small sets then transferred to the anaerobic chamber immediately to minimize air exposure and anoxic RW or DIW added. Bottles were stoppered, removed from the anaerobic chamber, crimped, flushed with Ar for 15-20 mins then an anoxic 10 mM NO3 stock solution was added to appropriate bottles using a 5 mL syringe and 22 G needle. The volume of nitrate stock solution added was adjusted seasonally to account for the nitrate initially present in the RW and to achieve a final concentration of 500 µM in the sample bottles. The acetylene block method for the determination of denitrification (Yoshinari and Knowles, 1976) was employed by adding 5 mL of anoxic, hydrogen-free acetylene (generated from calcium carbide) to the serum bottles after equilibration. An additional 10 mL of Ar was added to each bottle to generate an internal positive pressure for sampling. The bottles were equilibrated at 20 deg C with mixing for approximately 15-45 mins. Headspace gas samples were removed 6-10 times over the course of a few hours with a Hamilton gastight syringe and analyzed for nitrous oxide (N2O) concentration using a gas chromatograph (GC) equipped with an electron capture detector (ECD). Sample bottles were mixed on an end-over-end rotator at room temperature in between samplings. An HNU model 301 GC was used to analyze N2O for the April and August 2022 experiments but a Shimadzu model GC-17a was used for the November 2022 experiment due to problems with the HNU. The Shimadzu GC was not equipped with a backflush valve to remove acetylene thus a modification to the sampling method was necessary. A 0.5 mL headspace gas sample was removed from the 60 mL sample bottle using a gastight syringe and injected into a 10 mL Ar-flushed serum bottle containing several pellets of palladium, then an additional 5 mL of Ar added anaerobically to provide sufficient pressure for sampling. This procedure converted the acetylene to ethylene via the palladium catalyst, as well as diluted the nitrous oxide gas sample to within the standard curve range. Three timepoints were collected from each of the original sediment and water bottles over the course of 3 h. Nitrous oxide concentration (micromole per liter) over time were plotted and the linear portion of the curve (typically the first 4-6 h) used to determine the rate of production, standard error (SE) of the slope, and coefficient of variation (R squared). *Note: Due to a shortage of lab personnel to assist in November 2022 some adjustments were made to these experiments. Results from April and August 2022 indicated no significant difference in N2O production rates for the unamended UF RW + Sed versus the UF RW + Sed + NO3 treatments, thus the unamended UF RW + Sed treatment was omitted in November 2022. In addition, the DIW, RW, and RW + NO3 treatments were omitted given the very low or negligible rates of N2O production during the incubation time period in April and August 2022. 3) Anoxic Carbon Dioxide production: Rates of carbon dioxide (CO2) production or reduction were measured in 60 mL serum bottles containing the following treatments: 1) 20 g well-mixed < 2 mm sieved wet sediment plus 30 mL anoxic UF RW, in triplicate, 2) 20 g well-mixed < 2 mm sieved wet sediment plus 30 mL anoxic UF RW amended to 500 µM NO3, in triplicate, 3) 30 mL anoxic UF RW, in duplicate, 4) 30 mL anoxic UF RW amended to 500 µM NO3, in duplicate, and 5) 30 mL anoxic DIW. Sediment was weighed into the bottles aerobically on the lab bench in small sets then transferred to the anaerobic chamber immediately to minimize air exposure and anoxic RW or DIW added. Bottles were stoppered, removed from the anaerobic chamber, crimped, and flushed with Ar for 15-20 mins. An anoxic 10 mM NO3 stock solution was added to appropriate bottles using a 5 mL syringe and 22 G needle and an additional 25 mL of Ar added to each bottle using a 30 mL syringe and attached needle. The volume of NO3 stock solution added was adjusted seasonally to account for the NO3 initially present in the RW and to achieve a final concentration of 500 µM in the sample bottles. Headspace gas samples were removed daily for 4-5 days using a Hamilton gastight glass syringe with 22 G needle and injected into a LICOR Model LI-6262 infrared CO2 analyzer with N2 carrier gas. At the end of the experiment, the pressure remaining in the bottle was measured using a Crystal Model 31 Pressure Calibrator with attached 22 G needle. The bottle was then opened, and 5 mL of liquid removed for pH measurement. Concentration (micromole per liter or micromole per gram dry sediment) over time was plotted and the linear portion of the CO2 gas concentration curve used to determine CO2 production rates. Carbon dioxide concentration (micromoles per liter) over time were plotted and the linear portion of the curve used to determine rate of production, standard error (SE) of the slope, and coefficient of variation (R squared). *Note: Due to a shortage of lab personnel to assist in November 2022 some adjustments were made to these experiments. Results from April and August 2022 indicated no significant difference in CO2 production rates for the unamended UF RW + Sed versus the UF RW + Sed + NO3 treatments, thus the unamended UF RW + Sed + NO3 treatment was omitted in November 2022. In addition, the DIW and RW + NO3 treatments were omitted given the very low or negligible rates of CO2 production during the incubation time period in April and August 2022. The UF RW treatment was retained as a control. 4) Methanogenesis: Rates of methane (CH4) production were measured in April 2022 by removing headspace samples from the anoxic respiration bottles using a 10 mL glass syringe and 22 G needle at T0, Tmid (middle timepoint) and Tf timepoints only (days 1, 3, and 5) and analyzed for methane concentration using a Hewlett Packard Model HP5890 gas chromatograph equipped with a flame ionization detector (FID). In August and November 2022, separate bottle sets were prepared for the CH4 experiments as described for the anoxic carbon dioxide production experiments above and sampled daily for ~5-6 days. At the end of the experiment, the pressure remaining in the bottle was measured using a Crystal Model 31 Pressure Calibrator with attached 22 G needle. The bottle was then opened, and 5 mL of liquid removed for pH measurement. Methane concentration (micromoles per liter) over time were plotted and the linear portion of the curve used to determine rate of production, standard error (SE) of the slope, and coefficient of variation (R squared). *Note: Due to a shortage of lab personnel to assist in November 2022 some adjustments were made to these experiments. Results from April and August 2022 indicated a potential lag in CH4 production rate when NO3 was added to the UF RW + Sed treatment, thus both the unamended UF RW + Sed and UF RW + Sed + NO3 treatments were included in November 2022. However, the DIW, RW, and RW + NO3 treatments were omitted given the very low or negligible rates of CH4 production during the incubation time period in April and August 2022. 5) Anoxic Dissolved Inorganic Carbon: To determine dissolved inorganic carbon (DIC) change over the course of the anoxic experiments, 60 mL serum bottles were prepared containing the following treatments: 1) 20 g well-mixed < 2 mm sieved wet sediment plus 30 mL anoxic UF RW, duplicates for beginning (T0) and end (Tf), 2) 30 mL anoxic UF RW, duplicates for T0 and Tf, and 3) 30 mL anoxic DIW, 1 replicate for T0 and Tf. Bottles were stoppered, crimped, 25 mL additional Ar added, then mixed on an end-over-end rotator at room temperature (20-22 deg C). After an equilibration period of approximately 2-3 h, T0 samples were collected from the stoppered serum bottle by carefully removing approximately 15 mL liquid using an 18 G needle and 30 mL syringe. The needle was removed and a 0.45 µm filter with attached 22 G needle was connected to the syringe. The contents were filtered into a previously prepared stoppered, crimped, Ar-flushed 30 mL serum bottle containing 400 µL 18 N sulfuric acid to convert all aqueous DIC to CO2. Sample bottles were analyzed at a later date for headspace CO2 concentration as previously described. The pH was measured on the remaining water and sediment in the original 60 mL serum bottle. This procedure was repeated at Tf, corresponding to the end of the CO2 production incubation. *Note: DIC change was measured in August and November 2022 only. 20221120 Deborah A Repert USGS - WATER Microbiologist mailing and physical

Boulder NRP,University of Colorado - Building 6

Boulder CO 80303 303-541-3020 darepert@usgs.gov Gas calculations: mL @ STP in headspace = nL/mL x (1 mL/106 nL) x mL hdsp x (Bldr hPa/1013 hPa) x ((hdsp mL + (Ar mL + Acet mL – gas removal mL))/hdsp mL) x (273 deg K/(273 deg K + lab deg C)) mL @ STP in water = nL/mL x (1 mL/106 nL) x mL water x (Bldr hPa/1013 hPa) x ((hdsp mL + (Ar mL + Acet mL – gas removal mL))/hdsp mL) x β Total mL in bottle = mL @ STP in headspace + mL @ STP in water uMolar Concentration = (total mL in bottle/mL water) x (1 mmol/22.414 mL) x (1000 mL/L) x (1000 umol/mmol) where, STP = standard temperature and pressure, nL = nanoliters, mL = milliliters, hdsp = headspace, Bldr = Boulder CO, hPa = hectopascals, Ar = argon, acet = acetylene, deg = degrees, K = Kelvin, C = Celsius, β = Bunsen coefficient, mmol = millimoles, 1/22.414 = ideal gas constant, umol = micromoles. 20221120 Deborah A Repert USGS - WATER Microbiologist mailing and physical

Boulder NRP,University of Colorado - Building 6

Boulder CO 80303 303-541-3020 darepert@usgs.gov Sequential Extraction of Sediment for Phosphorus and other metals content: The sequential phosphorus extraction method employed was a modification of the Wang et al. (2021) published method. Subsamples of the air-dried sediment were ground with a mortar and pestle and 0.5 g weighed into triplicate acid-washed 50 mL centrifuge vials. Extractions of the sediment were performed by adding 30 mL of the extractant to the centrifuge vial on sequential days in the following order: 1) deionized water (DIW), 2) 0.5 M sodium bicarbonate (NaHCO3), 3) 0.1 M sodium hydroxide (NaOH), then 4) 1 M hydrochloric acid (HCl). The vials were mixed on an end-over-end rotator for 16 h. Vials were then centrifuged at ~3000 rpm for 10 mins at 5 deg C, filtered through a 25 mm 0.45 µm Supor filter, being careful not to lose sediment in the process, and the next extraction step commenced. To determine inorganic P (Pi), 15 mL subsamples of the NaHCO3 and NaOH extracts were acidified with 5.4 mL of 0.9 M H2SO4 to precipitate extracted organic matter then analyzed for Pi, while subsamples of the H2O and HCl extracts were analyzed directly for Pi using a discrete analyzer and an ICP-OES. To determine total phosphorus (Pt), the H2O, NaHCO3, NaOH, and HCl extracts were further digested by adding 5 mL of sample to 50 mL culture tubes containing 10 mL of 0.9 M H2SO4 and 0.5 g ammonium persulfate [(NH4)2S2O8]. The culture tubes were mixed well with a vortex then autoclaved at 103.4 kPa, 121°C for 1 hr. Samples were then analyzed for Pt using an ICP-OES. The difference between Pt and Pi was estimated to be the organic P (Po) fraction (Tiessen and Moir, 1993). Residual P was determined after the last sequential extraction using a modified method by Thomas et al. (1967). The remaining sediment in the centrifuge tubes was dried at 35-40 °C, then 250 milligrams (mg) weighed into 250 mL phosphoric acid volumetric flasks. Five mL of concentrated H2SO4 was added to each flask then heated on a hotplate at 120-125 °C for 20 mins. The flasks were cooled, then 0.5 mL of peroxide added, and flasks digested at 120-125 °C for 10 mins. The peroxide addition was repeated 6 times until no color remained in the sediment. The digestion temperature was then increased to 250-255 °C for 45 mins to ensure all H2O2 was removed. The solution was cooled then diluted to the 250 mL line with DIW and analyzed for residual P using an ICP-OES. *Note: Given the expense of ICP analyses, not all samples were analyzed for each sequential extraction step. The data reported in this data release include samples that were deemed most important. The replicate numbers are tied to the original sample replicate for these sequential extractions. Thus, some replicate numbers may be missing if the samples were not analyzed. 20230518 Deborah A Repert USGS - WATER Microbiologist mailing and physical

Boulder NRP,University of Colorado - Building 6

Boulder CO 80303 303-541-3020 darepert@usgs.gov Analytical Instrumentation details: 1) Dissolved Organic Carbon, UV absorbance, and SUVA: Shimadzu TOC-L: The filtered background and experimental samples were collected in 40 mL baked glass vials and preserved with HCl (total organic carbon (TOC) only) or sulfuric acid (DOC and total nitrogen (TDN)) to pH < 2 to remove inorganic carbon and allow for better recovery of CO2 during detection of non-purgeable organic carbon (NPOC). Vials were chilled at (< 6 deg C ). The sample is drawn into the autosampler syringe and mixed with 1 N HCl to convert any inorganic carbon to CO2, then sparged with zero grade air and lost to the atmosphere, along with any purgeable organic carbon. The sample is then injected into the combustion furnace where it is heated to 680 deg C along with a platinum catalyst. Through decomposition the sample is converted to CO2, cooled, then sent to a non-dispersive infrared detector (NDIR) for measurement. Teledyne Tekmar Fusion Total Organic Carbon (TOC) UV/Persulfate analyzer: The filtered DIW extraction samples were collected in 40 mL baked glass vials and chilled (< 6 deg C). DOC samples were analyzed using a Teledyne Tekmar Fusion TOC analyzer which utilizes ultraviolet persulfate oxidation to CO2 and pressurization of the sample gas to increase sensitivity from the NDIR detector (EPA 415.1- 415.3). Agilent Model 8453 UV-visable Spectrophotometer: Filtered, unacidified samples were collected for ultraviolet absorbance (UVabs) in 40 mL glass vials and chilled at < 6 °C in the dark. UVabs was measured at 250 nanometers (nm) using an Agilent Model 8453 UV-visable Spectrophotometer (Helms et al., 2008; Potter and Winsatt, 2005). Specific UV absorbance was calculated by dividing the UVabs@254 by the concentration of DOC then multiplying by 100 (Aiken et al., 1992). 2) Total Dissolved Nitrogen: Background and experimental samples were filtered through 0.22 µm DIW rinsed Sterivex filters, collected in 40 mL baked glass vials, and preserved with 6 N HCl (3 µL per 1 mL sample). Samples were analyzed for total dissolved nitrogen (TDN) using a Skalar FormacsTMTN Total Nitrogen Analyzer equipped with an ND25 Nitrogen Detector (Repert et al., 2014). Samples were injected into a high temperature reactor (set at 900 deg C) where all chemically bound nitrogen (organic N + inorganic N) was converted to nitric oxide (NO). The catalysts (cerium oxide and chromium) in the reactor catalyze the oxidation to completion. In the reaction chamber of the chemiluminescent ND25 detector, NO combines with ozone to form excited dinitrogen gas, emitting light during fast decay which was then measured by a photomultiplier tube (PMT). The electrical signal from the PMT was amplified and conducted to a computer where the signal was calculated as concentration. 3) Major Anions by IC: Background and experimental samples for major anions, including chloride (Cl), nitrate (NO3), nitrite (NO2), phosphate (PO4), sulfate (SO4), were filtered through 0.22 µm DIW rinsed Sterivex filters, collected in 5 mL DIW rinsed plastic centrifuge vials, and preserved by freezing (-20 deg C). For the April and August 2022 experiments, samples were analyzed using a Thermo-Fisher Model ICS-5000 ion chromatograph (IC) equipped with AS4A 4-mm analytical and AG4A 4-mm guard columns, an AERS-500 4 mm suppressor, 1.8 millimolar (mM) sodium carbonate plus 1.7 mM sodium bicarbonate isocratic mobile phase, and a CD20 conductivity detector (Smith et al., 2019). For the November 2022 experiments and all the DIW extractions, samples were analyzed using a Thermo-Fisher Model ICS-5000 IC equipped with AS22/AG22 4-mm analytical and guard columns, an AERS-500, 4 mm suppressor, 4.5 millimolar (mM) sodium carbonate/1.4 mM sodium bicarbonate isocratic mobile phase, and a CD20 conductivity detector. 4) Major Cations and Ammonium by IC: Background and experimental aqueous samples for major cations, including sodium (Na), ammonium (NH4), potassium (K), magnesium (Mg), and calcium (Ca), were filtered through 0.22 µm DIW rinsed Sterivex filters, collected in 5 mL DIW rinsed plastic centrifuge vials, and preserved with 18 N sulfuric acid (2 µL per 1 mL sample) then chilled (4 deg C). QAQC acidified DIW blanks were prepared and stored in a similar manner. Samples and blanks were analyzed using a Thermo-Fisher Model ICS-5000 IC equipped with CS12A 4-mm analytical and CG12A 4-mm guard columns, a CERS-500 4-mm suppressor, 20 mM methanesulfonic acid isocratic mobile phase, and a CD20 conductivity detector (Smith et al., 2019). A subset of samples was also run on the ICP for major cations. Values from the ICP were < 5% different than IC values, thus values from the IC only were reported in the data release. 5) Nutrients by discrete analysis: Samples for nitrite from the experiments, nitrate plus nitrite and ammonium from the sediment KCl extractions, and phosphate from the sediment sequential extractions were analyzed colorimetrically using a SEAL AQ300 Discrete Analyzer. Samples for nitrite and nitrate plus nitrate in KCl were filtered through 0.22 µm DIW rinsed Sterivex filters, collected in either 5 mL plastic vials or 60 mL plastic bottles, and preserved by freezing (-20 deg C). Samples for ammonium in KCl were filtered through 0.22 µm DIW rinsed Sterivex filters, collected in 40 mL glass vials, and preserved with 18 N sulfuric acid (2 µL per 1 mL sample) then chilled (4 deg C). Nitrate plus nitrite concentration was measured using a cadmium coil reduction method followed by sulfanilamide reaction in the presence of N-(1-napthylethylenediamine) dihydrochloride (Method Number AGR-231-A Rev 0) with a method detection limit of 0.015 mg N/L. The indophenol blue method (Method Number AGR-210-A Rev. 1) was used for the determination of ammonium concentration in the acidified KCl extracts with a method detection limit of 0.04 mg N/L. Nitrite concentration was measured following the EPA method (Method number EPA-115-D Rev. 1A) for analysis of nitrite in water samples with a method detection limit of 0.008 mg N/L. Phosphate concentration in the extracts from the sediment sequential extractions was measured using method number AGR-203-A Rev. 4 for analysis of phosphate in Olsen’s P extract of soil with a method detection limit of 0.01 mg P/L. 6) Trace Metal elements by ICP: Background, DIW extracted, and sequential extraction samples for trace metal concentrations were collected in 125 mL acid-washed bottles. A 3 mL sample was diluted and preserved with 27 mL 1 percent nitric acid (HNO3). Concentrations of dissolved major cations, silica, and trace metals (Arsenic (As), Boron (B), Barium (Ba), Cadmium (Cd), Chromium (Cr), Cobalt (Co), Copper (Cu), Iron (Fe), Lithium (Li), Manganese (Mn), Nickel (Ni), Phosphorus (P), Selenium (Se), Silica (SiO2), Strontium (Sr), Titanium (Ti), Zinc (Zn)) were determined using inductively coupled plasma– optical emission spectroscopy (ICP–OES) on a PerkinElmer 7300 DV ICP–OES and the software utilized was PerkinElmer WinLab 32 for ICP (Version 5.1.2.0549) (Roth et al., 2022). 7) Sediment CHN Characterization: Subsamples of the < 2 mm sieved core material were dried at 50 deg C for percent moisture and analysis of carbon and nitrogen content. Dried samples were ground using a shatterbox or mortar and pestle then analyzed for percent carbon and nitrogen using an Exeter CE-440 Elemental Analyzer. Samples are weighed in consumable tin capsules, injected into a high temperature furnace, and combusted in pure oxygen at 980 °C. The resulting combustion products pass through tubes containing specialized regents to produce carbon dioxide (CO2), water (H2O) and nitrogen (N2) and oxides of nitrogen. The gases then pass through a reduction tube containing copper to reduce nitrogen oxides to elemental nitrogen. The mixture then passes through a series of high-precision thermal conductivity detectors, each containing a pair of thermal conductivity cells to differentially measure hydrogen and carbon dioxide. Nitrogen is measured against a helium reference (https://www.eai1.com; Zimmerman et al., 1997). 8) Headspace Gases: For the April and August 2022 denitrification experiments, gas samples for nitrous oxide analysis were removed from the headspace of the sample bottles using a gastight syringe plus 22 G needle and immediately injected into an HNU model 301 gas chromatograph (GC) equipped with a Porapak N column, P-5 carrier gas (95% argon, 5% methane), a Valco Instruments Ni-63 electron capture detector (ECD), and a backflush valve to prevent acetylene from reaching the detector [(Brooks et al., 1992)]. For the November 2022 denitrification experiment, gas samples were removed from the headspace then injected into Ar-flushed 10 mL serum bottles containing palladium catalyst to reduce the acetylene. Five mL of Ar was then added to the bottles to provide pressure for analysis. The next day these bottles were analyzed for nitrous oxide using a Shimadzu model GC-17a equipped with a Porapak N column, N2 carrier gas, and Ni-63 ECD (Repert et al., 2014)]. For the CO2 and DIC production, gas samples were removed from the headspace of the sample bottles for measurement of CO2 using a gastight syringe plus 22 G needle, then injected into a LICOR Model LI-6262 infrared CO2 analyzer with nitrogen carrier gas. For the methanogenesis experiments, gas samples were removed from the headspace of the sample bottles for measurement of CH4 using a gastight syringe plus 22 G needle, then injected into an HP5890 series II gas chromatograph, equipped with a Porapak N column, N2 carrier gas, and flame ionization detector (FID) (Yamamoto et al., 1976). 9) KCl extractable N species Sediment samples extracted with KCl were analyzed for NH4 and NO3+NO2 concentration using a SEAL Analytical AQ300 discrete analyzer (https://www.seal-analytical.com). The indophenol blue method (Method Number AGR-210-A Rev. 1) was used for the determination of ammonium concentration in the acidified KCl extracts. NO3+NO2 concentration was measured using a cadmium coil reduction method followed by sulfanilamide reaction in the presence of N-(1-napthylethylenediamine) dihydrochloride (Method Number AGR-231-A Rev 0). 20250121 Deborah A Repert USGS - WATER Microbiologist mailing and physical

Boulder NRP,University of Colorado - Building 6

Boulder CO 80303 303-541-3020 darepert@usgs.gov References: Note: Some of the references listed below are found only in the IRB_DataDictionary file. Aiken, G.R., McKnight, D.M., Thorn, K.A., & Thurman, E.M., 1992, Isolation of hydrophilic acids from water using macroporous resins, Organic Geochemistry, v. 18, pp. 567-573. https://doi.org/10.1016/0146-6380(92)90119-I Barringer, J.L., and Johnsson, P.A., 1996, Theoretical considerations and a simple method for measuring alkalinity and acidity in low-pH waters by Gran titration: U.S. Geological Survey Water Resources Investigations Report 89-4029, 36 p. Brooks, M. H., Smith, R. L., & Macalady, D. L., 1992, Inhibition of existing denitrification enzyme activity by chloramphenicol. Applied and environmental microbiology, 58(5), 1746-1753. Fishman, M.J., and Friedman, L.C., 1989, Methods for determination of inorganic substances in water and fluvial sediments: Techniques of Water-Resources Investigations of the U.S. Geological Survey, Book 5, Chapter A1, p. 55-56, 55-56 p. Helms, J. R., Stubbins, A., Ritchie, J. D., Minor, E. C., Kieber, D. J., & Mopper, K., 2008, Absorption spectral slopes and slope ratios as indicators of molecular weight, source, and photobleaching of chromophoric dissolved organic matter. Limnology and oceanography, 53(3), 955-969. https://doi.org/10.4319/lo.2008.53.3.0955 NIST, 1999, National Institute of Standards and Technology, Certificate of Analysis, Standard Reference Sample 1643d, Trace Elements in Water, https://www-s.nist.gov/srmors/certificates/archive/1643d.pdf. Potter, B. B., & J. C. Wimsatt, 2005, Method 415.3 - Measurement of total organic carbon, dissolved organic carbon and specific UV absorbance at 254 nm in source water and drinking water. U.S. Environmental Protection Agency, Washington, DC. Repert, D. A., Underwood, J. C., Smith, R. L., & Song, B., 2014, Seasonal nitrogen cycling processes and relationship to microbial community structure in bed sediments from the Yukon River at Pilot Station, Journal of Geophysical Research: Biogeoscience, 119, 2328-2344. https://doi.org/10.1002/2014JG002707 Roth, D.A., Johnson, M.O., McCleskey, R.B., Riskin, M.L., & Bliznik, P.A., 2022, Evaluation of preservation techniques for trace metals and major cations for surface waters collected from the U.S. Geological Survey's National Water Quality Network Sites: U.S. Geological Survey data release, https://doi.org/10.5066/P9SMPZ3M Smith, R. L., Repert, D. A., Stoliker, D. L., Kent, D. B., Song, B., LeBlanc, D. R., McCobb, T.D., Bohlke, J.K., Hyun, S.P., & Moon, H.E., 2019, Seasonal and spatial variation in the location and reactivity of a nitrate contaminated groundwater discharge zone in a lakebed, Journal of Geophysical Research: Biogeosciences, 124(7), pp. 2186-2207. https://doi.org/10.1029/2018jg004879 Thomas, R. L., Sheard, R. W., & Moyer, J. R., 1967, Comparison of conventional and automated procedures for nitrogen, phosphorus, and potassium analysis of plant material using a single digestion 1. Agronomy Journal, 59(3), 240-243. Tiessen, H. & Moir, J. O., 1993, Characterization of available phosphorus by sequential extraction. In: Carter MR (ed) Soil sampling and methods of analysis. Canadian Society of Soil Science, Lewis Publishers, Boca Raton, pp 75–86. U.S. Dept. of Commerce, 2012, National Institute of Standards and Technology. Accessed 5-22-2012, at http://www.nist.gov/index.html. Wang, Y. T., Zhang, T. Q., Zhao, Y. C., Ciborowski, J. J., Zhao, Y. M., O'Halloran, I. P., ... & Tan, C. S., 2021, Characterization of sedimentary phosphorus in Lake Erie and on-site quantification of internal phosphorus loading, Water Research, 188, 116525. https://doi.org/10.1016/j.watres.2020.116525 Weiss, R.F. & Price, B.A., 1980, Nitrous oxide solubility in water and seawater, Marine chemistry, 8(4), pp. 347-359. Yamamoto, S., Alcauskas, J. B., & Crozier, T. E., 1976, Solubility of methane in distilled water and seawater. Journal of Chemical and Engineering Data, 21(1), 78-80. Yoshinari, T., & Knowles, R. (1976). Acetylene inhibition of nitrous oxide reduction by denitrifying bacteria. Biochemical and biophysical research communications, 69(3), 705-710. Zimmerman, C. F., C. W. Keefe, & J. Bashe, 1997, Method 440.0 Determination of Carbon and Nitrogen in Sediments and Particulates of Estuarine/Coastal Waters Using Elemental Analysis. U.S. Environmental Protection Agency, Washington, DC, EPA/600/R-15/009, 1997. 20250121 Deborah A Repert USGS - WATER Microbiologist mailing and physical

Boulder NRP,University of Colorado - Building 6

Boulder CO 80303 303-541-3020 darepert@usgs.gov Site details, background sediment and water data, experimental data, and sequential extraction data can be found in the following 9 .csv tables: 1. IRB_BkgWater&SedimentData.csv 2. IRB_BkgExtractionData.csv 3. IRB_ExptAqueousData.csv 4. IRB_ExptAnoxicGasData.csv 5. IRB_ExptOxicGasData.csv 6. IRB_ExptDICGasData.csv 7. IRB_ExptDataloggerData.csv 8. IRB_SeqExtractionData.csv 9. IRB_ExptRates.csv Definitions for column headers and characteristic name descriptions in the dataset tables are located in IRB_DataDictionary.xlsx. Repert, D. A., Castro, I. R., Dornblaser, M. M., Duncker, J. J., Fazio, D. J., Sharpe, J. B., Smith, R. L., Wickland, and K. P., 2025, Data from laboratory experiments to assess seasonal carbon, nitrogen, and phosphorus processing in riverbed sediments from the Illinois River near Starved Rock Lock and Dam, 2022, U.S. Geological Survey data release, https://doi.org/10.5066/P13DSK7B. GS ScienceBase U.S. Geological Survey mailing address

Denver Federal Center, Building 810, Mail Stop 302

Denver CO 80225 United States 1-888-275-8747 sciencebase@usgs.gov Unless otherwise stated, all data, metadata and related materials are considered to satisfy the quality standards relative to the purpose for which the data were collected. Although these data and associated metadata have been reviewed for accuracy and completeness and approved for release by the U.S. Geological Survey (USGS), no warranty expressed or implied is made regarding the display or utility of the data on any other system or for general or scientific purposes, nor shall the act of distribution constitute any such warranty. Digital Data https://doi.org/10.5066/P13DSK7B None 20250219 Deborah A Repert U.S. Geological Survey Microbiologist mailing and physical

3215 Marine St. E127

Boulder CO 80303 USA 303-541-3020 darepert@usgs.gov FGDC Biological Data Profile of the Content Standard for Digital Geospatial Metadata FGDC-STD-001.1-1999