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Soil net ammonification and net nitrification rates were measured in-situ using the “buried bag” method (Eno 1960; Westerman and Crothers 1980; Hanselman et al. 2004; Dúran et al. 2012). Two adjacent 10 x 10 cm soil samples were taken by removing the litter layer and sampling the full depth of the organic layer at 0, 15, 30, 60, and 90 meters from the forest edge. Soil was taken exclusively from the organic layer, as most decomposition of organic material and N mineralization occurs there (McNeill and Unkovich 2007). The depth of the organic soil layer was determined in the field with a ruler, according to techniques outlined by the Soil Science Society of America (Vero 2021). Variation in the depth of the soil organic layer was measured and calculated at each plot throughout the edge to interior gradient at each site. For each set of paired samples, one sample was returned to the lab immediately to be analyzed within 24 hours of field sampling (T0; n = 2 at each distance from the forest edge), while the remaining sample (Tf; n = 2 at each distance from the forest edge) was sealed in a gas-permeable plastic bag and placed back into the hole from which it was taken. The hole was covered with the removed leaf litter, and the bagged Tf sample was left to incubate in-situ for 26-35 days. Soils were incubated in the field July 20 - August 15 in 2018 and May 23 - June 2 and July 17 -August 19 in 2019. At the end of the incubation periods, the incubated cores were returned to the lab for analysis within 24 hours of field collection. The site at Blue Hills in Canton, MA was not sampled in 2018.

After field collection, fresh soils were put through a 2 mm metal sieve to remove roots, rocks, and other large debris and homogenized. 5 g of freshly sieved soils was shaken for 1 hour in 60 ml 2 M potassium chloride solution (KCl) to extract ammonium (NH4+) and nitrate (NO3-) ions for analysis. Each extract solution was then filtered using Whatman Grade 2 filters (8 m pore size), and the resulting solutions were stored at 4°C until further analysis.

Soil extracts were analyzed colorimetrically for NH4+ and NO3- concentrations using the microplate method. Soil extracts were pipetted into a 96 well microplate, including ammonium nitrate standards and quality control solutions (ERA Environmental). To quantify ammonium in solution, citrate, sodium nitroprusside, and hypochlorite reagents were added in succession, and the resulting color intensity was measured in a microplate reader (Versa Max tunable microplate reader) at an absorbance wavelength of 667 nanometers (Ali and Lovatt 1995). Nitrate in solution was quantified by adding a vanadium chloride solution, incubating for 16 hours, and running plates through a microplate reader at an absorbance wavelength of 540 nanometers. Net rates of ammonification and nitrification were calculated as the respective differences in ammonium and nitrate concentrations between Tf and T0 samples divided by the duration of the in-situ incubation period. Total net mineralization was calculated as the sum of net ammonification and net nitrification.

Soil moisture and soil organic matter content (SOM) were quantified using a 5-gram subsample of fresh, sieved soil from each buried bag soil sample from all three deployment periods (n = 6 measurements). Fresh soil was dried in an oven at 65 °C, and gravimetric water content was calculated as the difference between wet and dry soil mass, divided by wet mass, and multiplied by 100. SOM was quantified using the loss on ignition method. Each dried subsample used in the moisture analysis was then placed in a muffle furnace at 450 °C for 4 hours to combust organic matter. The combustion temperature of 450 °C was chosen to maximize removal of organic matter while avoiding dehydroxylation of clay minerals in soil samples (Nelson and Sommers 1996). SOM concentration was calculated as the difference between pre- and post-combustion mass divided by pre-combustion mass of the subsample and multiplied by 100.

Soil samples collected in July 2019 were analyzed for pH, conductivity, and bulk density. Soil pH was determined using a 5-gram subsample of freshly sieved soil from the soil cores collected in July 2019 (T0 samples). Soils were mixed with 10 mL distilled deionized (DDI) water and shaken to form a slurry, which was then measured with a pH meter (Mettler-Toledo SevenEasy pH). Separate subsamples of the same fresh soil were mixed with DDI water in a 1:5 soil-to-water ratio and shaken for 1 minute at 10-minute intervals over a period of 30 minutes for conductivity analysis. Making sure to keep the soil suspended in the water, conductivity (soluble salt content) in each sample was measured with a Hach HQ40D Portable Multi Meter with conductivity probe.

A separate set of soils collected in July 2019 at each transect location at each site were analyzed for bulk density. Samples were collected using a 10 cm depth hammer core (2.4 cm radius), and organic and mineral soil horizons were separated and measured for depth in the field. In the lab, soil samples were homogenized using a 2 mm metal sieve and weighed, dried, and reweighed. Roots, rocks, and other debris were separately weighed for each sample, and rocks were then added to 50 mL of water in a graduated cylinder to measure their volume by displacement. Dry and wet bulk density was then calculated for each sample as soil mass per rock-free volume.

Foliage from the mid- and upper canopy was sampled from three oak trees (Quercus spp.) using a manual pole pruner at each of the transect distances at each site in July 2019, preferentially sampling sun-lit leaves from the mid canopy. Samples were air-dried prior to processing. Leaves were ground using a Wiley mill (40 mesh) and then analyzed for C and N concentration using a NC2500 elemental analyzer (CE Elantech, Lakewood, NJ, USA). Foliar N, while it does not consider the nutrient content of belowground plant tissues or microbes, can be used as a proxy for N uptake by trees (Kahmen et al. 2008), and C:N gives a measure of lability of organic tissue.