Data Package Metadata   View Summary

SYNOPTIC SAMPLING: Sites 1 - 20 were sampled once in the winter of 2022 (January 12, 2022 and February 4, 2022). At each SCM site, 3 soil sampling locations were selected to capture spatial variation in SCM soil processes. The first was within 1 meter of the inlet structure, referred to as “Inlet”. The second location was in the approximate center of the SCM, deemed “Center” and the third location was within 1 meter of the SCM outlet structure, deemed “Outlet”. At wetland sites, we sampled soils within areas of current inundation, referred to as “Inundated Soil", and nearby floodplain soils that were not currently inundated, deemed “Dry Soil". At each sampling location, a 1-inch soil push probe was used to sample soils to a depth of 40 cm. In cases where compact clay soils prevented the full 40 cm from being sampled, such as wet pond soils, the depth of soil collected was recorded. We also collected soil bulk density samples using a 2 inch x 2 inch steel cylinder attached to a slide hammer. If there was standing water present at the time of sampling, surface water was filtered into acid washed, pre-combusted 40 mL amber vials using a pre-combusted 0.7 μm GFF filter and into 100 mL HDPE bottles using a 0.45 μm PES filter. We also collected grab samples of leaf litter or biomass if present on the soil surface of the SCMs and wetlands. Soil, biomass, and surfacewater samples were transported back to Virginia Tech on ice and stored at 4 °C until analysis.

MONTHLY SAMPLING: The two bioretention SCM’s (Sites 21 and 22) located on the Virginia Tech campus were sampled with the same procedure described in "SYNOPTIC SAMPLING" on a monthly basis from February 2022 to February 2023. The “Center" sample was omitted for monthly sample collection. There were no months during this period where surface water was present in the SCMs at the time of soil sampling. The outlet of SCM 2 was unable to be sampled in August and September of 2022 due to maintenance work being performed at this site. Soils were immediately placed in the fridge at 4 °C after sampling.

SOIL PROPERTIES: To determine soil properties of the different SCM types, non-WSOM extracted soil samples were sent to the Virginia Tech Soil Testing Laboratory (Maguire & Heckendorn, 2019). Prior to Soil Testing Lab analyses, soils were air-dried and crushed to pass through a 10 mesh (2 mm) sieve. P, K, Ca, Mg, Zn, Mn, Cu, Fe, B, and Al were measured using a Mehlich extraction procedure and elemental ICP analysis. pH was determined by mixing soil with water at a 1:1 (vol/vol) ratio for 10 minutes and then measuring water pH using a pH probe. Buffer pH was calculated by stirring in a Mehlich buffer solution into the soil-water mixture previously used to determine pH and allowing it to sit for 30 minutes before using the pH probe to collect a reading. Cation exchange capacity (CEC, reported as meq / 100 g soil), was calculated by summing extractable bases (Ca, Mg, and K) and buffer pH. Soil organic matter (%) was measured using loss on ignition testing where soils are heated at 360 degrees C for 2 hours to determine the weight loss of the soil.

WSOM PROCEDURE: The water-soluble organic matter (WSOM) extraction procedure was developed using methods from Duston (2020), Jones and Willett (2006), Gabor et al. (2006) and Rennert et al. (2007). An additional description of the method can be found in Wardinski, et. al (2022). On the date of extraction, field-moist soil samples were removed from the refrigerator and sorted to remove any large rocks, roots, and debris. 30 g of field moist soil was mixed with 150 mL of 0.01 M calcium chloride in a flask. Each sample was duplicated to produce the desired volume of extracted solution. The flasks were then placed on a shaker table for one hour at a speed of 200 rpm at room temperature (22 °C). An additional 15 g of soil was measured out and placed in an oven at 110 °C for 24 hours to determine moisture content of each soil sample. Immediately following shaking, the well-mixed soil-extract solution was divided into four 50 mL Falcon centrifuge tubes and centrifuged at 4,000 rpm (relative centrifugal force ~3,580 g) for 10 minutes. The centrifuged solution was then filtered into two acid washed, pre-combusted 40 mL amber vials for carbon analysis using an acid washed plastic syringe and pre-combusted 0.7 μm Whatman GF/F filter and the remaining centrifuged solution was filtered into a plastic 150 mL vial for nutrient analysis using a 0.45 μm PES filter. The vials were stored at 4 °C until further analysis. SCM biomass samples were also extracted using this procedure.

INSTRUMENTAL ANALYSES AND SPECTRAL METRICS: Filtered extracted soil solution (WSOM), extracted biomass solution, and surface water samples were analyzed for Non-Purgeable Organic Carbon (NPOC) using a Shimadzu TOC-Vcph Carbon Analyzer. Samples were analyzed for Total Dissolved Nitrogen, Nitrate, Ammonium, Total Dissolved Phosphorus, and Orthophosphate on a SEAL AutoAnalyzer 3. All samples were analyzed for absorbance on a Shimadzu UV Spectrophotometer immediately followed by fluorescence analysis on a Horiba FluoroMax-4 Spectrofluorometer. The spectrophotometer collected absorbance data from 190 to 850 nm in 1 nm increments and samples were blank-corrected during post-processing. To reduce inner-filter effects during fluorescence analysis, samples with absorbance values greater than 0.2 (1/cm) at 240 nm were diluted so that they fell within 0.02 – 0.2 (1/cm) (Ohno, 2002). Excitation-Emission Matrices (EEMs) were collected with an excitation wavelength range of 240-450 nm in 5 nm increments and emission wavelength range of 300-600 nm in 2 nm increments. During post-processing, EEMs were corrected for inner-filter effects, Raman normalized, and blank-corrected (Cory et al., 2010, Hounshell, et al., 2021, McKnight et. al, 2001). Spectral metrics including Specific Ultraviolet Absorbance at 254 nm (SUVA254), Fluorescence Index (FI), Humification Index (HIX), and Spectral Slope Ratio (SR) were calculated after post-processing of absorbance and fluorescence data to characterize organic matter composition. SUVA254 normalizes absorbance at 254 nm to NPOC concentration (Weishaar et al., 2003). FI is the ratio of emission at wavelengths 470 nm and 520 nm at an excitation wavelength of 370 nm (McKnight, 2001). HIX is the ratio of emission (435 nm – 480 nm) / (300 – 345 nm) measured at 254 nm excitation (Ohno, 2002, Gabor et. al, 2015, Zsolnay et al., 1999). SR is the ratio of (275 nm – 295 nm) to (350 nm – 400 nm) absorbance values (Helms et al., 2009). Additionally, fluorescence intensity at Peaks T, A, C, M, and N were determined in WSOM, surface water, and biomass samples (Cobble, 2006, Fellman et al., 2010). Parallel factor analysis (PARAFAC) decomposes the fluorescence signature of EEMs to identify the underlying organic matter signals (Murphy et al 2013). Cory and McKnight (2005) developed a thirteen component PARAFAC model using whole water samples from a wide range of aquatic environments. Using MATLAB (ver. 2019b), we assessed WSOM, biomass, and surface water sample loadings across the thirteen components from the Cory and McKnight (2005) PARAFAC model. Using sample loadings, we calculated Percent Protein (summation of protein-like component loadings) and Redox Index (ratio of reduced quinones to total quinones) (Cory and McKnight, 2005, Mladenov et al., 2006, Miller et al., 2006).

MONTHLY WATER BALANCE: We input NOAA monthly precipitation totals and average monthly temperatures for Blacksburg, VA, into a water balance model code developed by Gannon and McGuire (2022). This monthly water balance model uses temperature and precipitation along with soil water holding capacity to estimate monthly evapotranspiration, water storage within the soil, and surplus water available for recharge or runoff. In this application of the water balance, we applied monthly precipitation and temperature data from 2021, 2022, and 2023 into the code. Each year was analyzed individually due to the iterative nature of the water balance code. We don't calculate exact soil storage or surplus water volumes for the 2 SCMs, rather we use the generalized conditions to explore potential drivers of monthly variations in WSOM and nutrients. The code used to run the water balance is shared courtesy of J.P. Gannon and can be found at: https://github.com/jpgannon/Water-Balance-App. Further information about the water balance can be found at: https://cuahsi.shinyapps.io/WaterBalance/_w_cc50c2f8/#tab-1143-1.

REFERENCES:

Coble, P. G. (2007). Marine optical biogeochemistry: The chemistry of ocean color. Chemical Reviews, 107(2), 402–418. https://doi.org/10.1021/cr050350

Cory, R. M., & McKnight, D. M. (2005). Fluorescence spectroscopy reveals ubiquitous presence of oxidized and reduced quinones in dissolved organic matter. Environmental Science and Technology, 39(21), 8142–8149. https://doi.org/10.1021/es0506962

Cory, R. M., Miller, M. P., Mcknight, D. M., Guerard, J. J., & Miller, P. L. (2010). Effect of instrument-specific response on the analysis of fulvic acid fluorescence spectra. Limnology and Oceanography: Methods, 8(2), 67–78. https://doi.org/10.4319/lom.2010.8.67

Duston, S. A. (2020). Capturing and Characterizing Soluble Organic Matter Dynamics in Soil Formation Processes. Virginia Tech.

Fellman, J.B., E. Hood, and R.G.M. Spencer. 2010. Fluorescence spectroscopy opens new windows into dissolved organic matter dynamics in freshwater ecosystems: A review. Limnology and Oceanography 55, 2452-2462. https://doi.org/10.4319/lo.2010.55.6.2452

Gabor, R. S., Burns, M. A., Lee, R. H., Elg, J. B., Kemper, C. J., Barnard, H. R., & McKnight, D. M. (2015). Influence of leaching solution and catchment location on the fluorescence of water-soluble organic matter. Environmental Science and Technology, 49(7), 4425–4432. https://doi.org/10.1021/es504881t

Gannon, J. P., & McGuire, K. J. (2022). An interactive web application helps students explore water balance concepts. Frontiers in Education, 7, 873196. https://doi.org/https://doi.org/10.3389/feduc.2022.873196

Helms, R. J., Stubbins, A., Ritchie, J. D., Minor, E. C., Kieber, D. J., & Mopper, K. (2009). Absorption spectral slopes and slope ratios as indicators of molecular weight, source, and photobleaching of chromophoric dissolved organic matter (Limnology and Oceanography 53 955-969). Limnology and Oceanography, 54(3), 1023. https://doi.org/10.4319/lo.2009.54.3.1023

Hounshell, Alexandria G., Thai, Rose H., Peeler, Kelly A., Scott, Durelle T., Carey, C. C. (n.d.). Time series of optical measurements (absorbance, fluorescence) for Beaverdam and Falling Creek Reservoir in Southwestern Virginia, USA 2019-2020 ver 1. Environmental Data Initiative. https://doi.org/https://doi.org/10.6073/pasta/d1062b14ed1df86507414afe8d45dc75

Jones, D. L., & Willett, V. B. (2006). Experimental evaluation of methods to quantify dissolved organic nitrogen (DON) and dissolved organic carbon (DOC) in soil. Soil Biology and Biochemistry, 38(5), 991–999. https://doi.org/10.1016/j.soilbio.2005.08.012

Maguire, R. O., & Heckendorn, S. E. (2019). Laboratory procedures: Virginia Tech soil testing laboratory.

McKnight, D. M., Boyer, E. W., Westerhoff, P. K., Doran, P. T., Kulbe, T., & Andersen, D. T. (2001). Spectrofluorometric characterization of dissolved organic matter for indication of precursor organic material and aromaticity. Limnology and Oceanography, 46(1), 38–48. https://doi.org/10.4319/lo.2001.46.1.0038

Miller, M. P., McKnight, D. M., Cory, R. M., Williams, M. W., & Runkel, R. L. (2006). Hyporheic exchange and fulvic acid redox reactions in an alpine stream/wetland ecosystem, Colorado front range. Environmental Science and Technology, 40(19), 5943–5949. https://doi.org/10.1021/es060635j

Mladenov, N., Huntsman-Mapila, P., Wolski, P., Masamba, W. R. L., & McKnight, D. M. (2008). Dissolved organic matter accumulation, reactivity, and redox state in ground water of a recharge wetland. Wetlands, 28(3), 747–759.

Murphy, K. R., Stedmon, C. A., Graeber, D., & Bro, R. (2013). Fluorescence spectroscopy and multi-way techniques. PARAFAC. Analytical Methods, 5(23), 6557–6566. https://doi.org/10.1039/c3ay41160e

National Oceanic and Atmospheric Administration. Climate Data Online: Global Historical Climatology Network - Daily. Retrieved from https://www.ncdc.noaa.gov/cdo-web/datasets

Ohno, T. (2002). Fluorescence inner-filtering correction for determining the humification index of dissolved organic matter. Environmental Science and Technology, 36(4), 742–746. https://doi.org/10.1021/es0155276

Rennert, T., Gockel, K. F., & Mansfeldt, T. (2007). Extraction of water-soluble organic matter from mineral horizons of forest soils. Journal of Plant Nutrition and Soil Science, 170(4), 514–521. https://doi.org/10.1002/jpln.200625099

Wardinski, K., Scott, D., McLaughlin, D., Hotchkiss, E., Jones, C. N., & Strahm, B. (2022). Water soluble organic matter from Delmarva Bay soils. Environmental Data Initiative. https://doi.org/https://doi.org/10.6073/pasta/e1f97936004a0af9a6b8cd0710d56046

Weishaar, J. L., Aiken, G. R., Bergamaschi, B. A., Fram, M. S., Fujii, R., & Mopper, K. (2003). Evaluation of specific ultraviolet absorbance as an indicator of the chemical composition and reactivity of dissolved organic carbon. Environmental Science and Technology, 37(20), 4702–4708. https://doi.org/10.1021/es030360x

Zsolnay, A., Baigar, E., Jimenez, M., Steinweg, B., & Saccomandi, F. (1999). Differentiating with fluorescence spectroscopy the sources of dissolved organic matter in soils subjected to drying. Chemosphere, 38(1), 45–50. https://doi.org/10.1016/S0045-6535(98)00166-0