MarielaGarcia Arredondomarielagarcia794@gmail.comhttps://orcid.org/0000-0001-8591-364XZoeCardonzcardon@mbl.eduhttps://orcid.org/0000-0001-8725-7842YilinFangyilin.fang@ppnl.govhttps://orcid.org/0000-0003-1969-9889MarcoKeiluweitmarco.keiluweit@unil.chhttps://orcid.org/0000-0002-7061-8346Steve B.Yabusakiyabusaki@pnnl.govMorrisJonesjonesm@franklinpierce.eduhttps://orcid.org/0000-0001-5418-9329U.S. DOE > Office of Science > Biological and Environmental Research (BER)http://dx.doi.org/10.13039/100006206fundingOrganization2024Here we share the data in column format via csv files for pH, redox, and dissolved oxygen collected at hourly resolution from microelectrodes. Dissolved organic carbon concentrations collected from TOC are also provided in a similar format but are composited samples from hourly microdialysis collection. This provided resolution of diel rhizosphere dynamics belowground. Plant physiological data was also collected at every 5 min for 24 hr cycles in order to capture diel dynamics aboveground. This data was used to parameterize the reaction transport model eSTOMP-ROOTS, which examines the rhizosphere biogeochemistry of a growing Vicia faba plant. The aim was to investigate plant activity and belowground biogeochemical processes, particularly their impact on mineral-organic associations in the rhizosphere. We combined in-situ rhizosphere microsensor and plant physiological measurements with a 3-D plant-soil reactive transport model to explore the behavior of dissolved organic carbon (DOC) in the rhizosphere. Over several days, microdialysis probes placed at the root-soil interface in live soil showed distinct daily patterns of DOC concentration in the pore water. Spikes in DOC concentrations during the day aligned with peaks in leaf-level photosynthesis, accompanied by decreasing redox potential and dissolved oxygen levels, and increasing pH in the rhizosphere. This new mechanistic modeling framework, which integrates aboveground plant physiological data with non-destructive, high-resolution monitoring of rhizosphere processes, offers significant potential for studying the factors that control carbon storage in soils.EARTH SCIENCE > LAND SURFACE > SOILSCATEGORICAL:GCMDEARTH SCIENCE > AGRICULTURE > SOILS > SOIL PHEARTH SCIENCE > AGRICULTURE > SOILS > CARBONVARIABLE:GCMDESS-DIVE File Level Metadata Reporting FormatCATEGORICAL:NONE

Garcia Arredondo, M., Fang, Y., Jones, M., Yabusaki, S., Cardon, Z., Keiluweit, M., 2023. Resolving dynamic mineral-organic interactions in the rhizosphere by combining in-situ microsensors with plant-soil reactive transport modeling. Soil Biology and Biochemistry 184, 109097. https://doi.org/10.1016/j.soilbio.2023.109097

This work is licensed under the Creative Commons Attribution 4.0 International License. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/.2019-06-132023-03-04A fertile grassland soil classified as fine-loamy, mixed, superactive Ustic Haplocryolls was collected at a 15 cm depth at the Rocky Mountain Biological Laboratory (Gothic, CO).-106.98777718989066-106.9877771898906638.958717083478438.9587170834784MarielaGarcia Arredondomarielagarcia794@gmail.comhttps://orcid.org/0000-0001-8591-364XSticky Roots — Implications of Altered Rhizodeposition (Driven by Cryptic, Viral Infection of Plants) for the Fate of Rhizosphere Mineral–Organic Matter Associations in Natural EcosystemsSoils were brought to 40% water holding capacity using tap water and packed into the rhizobox to achieve field bulk density (1.5 g cm 3). A single V. faba (Windsour variety) seedling with a tap root of ~3 cm length was transferred into the rhizobox. Soil moisture was maintained using a cotton wick connected to an amber glass flask containing tap water placed along the side of the rhizobox. The polycarbonate portions of the rhizobox were covered with aluminum foil to limit light exposure, leaving the top open for the plant to grow, and placed in a rack where it was held at a 30◦ angle to facilitate plant root growth along the sensor array. Rhizoboxes were maintained in a growth chamber set to 16 h light and 8 h dark cycle with a maximum of 500 μmol photons m 2 s 1 at 20% humidity for light hour settings and 25 ◦C. light cycle temperature and 15 ◦C dark cycle temperature. Plants entered the second nodal stage by the time their roots arrived at sensor array within the rhizoboxes.Microelectrodes for pH, EH, and DO concentration measurements were installed along the side port along which single roots were channeled and were constructed in-house within 1/16” PEEKTM tubing. pH microelectrodes were designed with modifications from Perley (1939). These were calibrated versus a Saturated Calomel Electrode or solid-state Ag/AgCl electrode in pH buffers. Microprobe redox electrodes were built following previous reporting in Vepraskas (2002) with modifications to fit the rhizobox side ports. Changes in DO were monitored using solid Hg/Au voltametric microelectrodes. Hg/Au microelectrodes were constructed as previously reported in the literature (Luther et al., 2008) and used with stationary Ag/AgCl reference and platinum (Pt) counter electrodes. All measurements were made with a DLK-70 potentiostat (Analytical Instruments Systems, Ringoes, NJ) laboratory electrochemical analyzer which communicated via standard web browser on a laboratory laptop positioned outside of the growth chamber with connector cables extending outside from the analyzer to the micro-electrodes inside the rhizobox within the growth chamber. DO was quantified by linear sweep voltammetry (LSV), between 0.1 and 1.9 V at a scan rate of 100 mV s 1 with 10s of deposition at 0.1 V. For DO, a two-point calibration was conducted using oxygen-saturated NaCl saline (20 mM) water (100% saturation) and a deoxygenated NaCl saline (20 mM) water (0% saturation). To maintain reproducibility during analysis, the Au/Hg amalgam electrodes ran cyclic voltammetry before each LSV scan. The redox and pH probes were similarly maintained by running initial scans before collecting stable final data points at each time step in the experiments.Along the interior wall of each individual rhizobox, a 10 mm long microdialysis probe (CMA 20 Harvard Apparatus) was installed. The positioning of the microdialysis probe allowed continuous measurements of the root-soil interface along individual roots. Because the soft original tubing for the microdialysis was noticeably leaking DOC, the original soft tubing. Rhizobox and associated microsensor setup. Magnified panel shows microsensor array containing microelectrodes controlled by the potentiostat as well as microdialysis probe connected to water pump and fraction collector. Rhizboxes are tilted at an angle of 30◦ degrees to facilitate root growth over the microsensor array. Detailed rhizobox design and dimensions included found in publication referenced in this report. Conceptualization of the biogeochemical reactions included in the numerical model to capture the measured diel dynamics in the rhizosphere. Rhizosphere biogeochemical parameters directly measured using microsensors are shown in white. Specific reactions incorporated into multicomponent reaction transport model (i.e., eSTOMP) are shown in yellow. We assumed that, during light periods (daytime), photosynthetic and transpiration activity are at their highest, causing high water uptake and exudation rates at the root-soil surface. Root exudates would increase DOC concentrations, with DOC being subject to microbial consumption (via respiration) and sorption to minerals. Microbial consumption of DOC was expected to lead to declines in DO and thus EH. Declines in EH and DO were assumed to lead to declines in aerobic respiration and relative to anaerobic respiration (Fe(III) oxide reduction). We anticipated that Fe(III) oxide reduction would re-mobilize Fe-bound OC. was removed and replaced by more inert tubing (PEEKTM, 0.65 mm OD x 0.12 mm ID, Sigma-Aldrich). This tubing fed into a fraction collector (CMA 142 Fraction Collector Harvard Apparatus, Hollister, MA) with previously sterilized capped glass vials that allowed for the collection of 300 μL h 1 per rhizobox. Ultrapure water was pumped through 10 mL airtight glass syringes to the microdialysis system by an auto-pump (Standard Infuse/Withdraw PHD, 2000 Syringe Pump, Harvard Apparatus, Hollister, MA). These syringes had three-port microvalves attached that were fastened to the ends of the PEEKTM tubing to the microdialysis system. The microdialysis tubing once affixed to the rhizobox was flushed with ultrapure water for 24 h. Once placed inside the growth chamber, rhizoboxes and microdialysis system were allowed to equilibrate for at least 24 h to adjust from soil rewetting after seedlings were transplanted into the rhizobox system. Once the tap root emerged approximately 1 cm from the top of the microdialysis probe, sampling was initiated to collect background soil pore water. Sampling continued as the root reached the probe, slowly growing along its surface. Samples were collected hourly (300 μL h 1) and subsequently combined to provide composite samples of 4-h resolution. Composites were split, with half flash frozen and the other half refrigerated for immediate DOC analysis. Microdialysis sampling was conducted over three days whereas microelectrode measurements were conducted over five days. DOC concentrations were analyzed using the TOC-L/V- liquid manual smallvolume sample injection unit for the total organic carbon analyzer (Shimadzu TOC-L CPH with an ASI-L, Shimadzu Scientific Inst., Columbia, MD, USA) with a glass airtight syringe following Stubbins and Dittmar (2012). Sample aliquots of 100 μL were manually injected and found to provide the best accuracy and precision with the smallest sample volume.Photosynthesis and transpiration rates were measured on four replicate rhizoboxes treated identically to those described earlier. Measurements were taken on leaves on the second node using a LiCOR 6400XT gas exchange system (LI-COR Biosciences, Lincoln, NE) and leaf internal concentration (ci between 221 and 248 μmol CO2 mol 1 in the day) provided by a blue/ red LED light source, temperature-controlled leaf cuvette set to 25 ◦C and light levels at 500 P AR, with an atmospheric CO2 buffer attached (tubing attached to LiCOR 6400XT that fed into 20 L carboy). Measurements were taken over a 24-hr period and recorded every 5 min to record diel plant responses in photosynthesis, stomatal conductance, and transpiration. In addition, we collected leaf area and leaf count from V. faba at the second node stage. This information was used to parameterize photosynthetic and transpiration rates for the entire plant system in the model.ZoeCardonMarine Biological Laboratory, Woods Hole, MAzcardon@mbl.eduprincipalInvestigatorDOE:DE-SC0021093RTM_dd.csvtext/csvPhotosynthesisData.csvtext/csvfaba_2_redox.csvtext/csvfaba_2_pH.csvtext/csvfaba_2_DOC.csvtext/csvfaba_2_DO.csvtext/csvRTMdata_flmd.csvtext/csv