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Taken from Darby et al. (in revision) Soil Biology and Biochemistry, with modest edits for brevity:

Soils were collected in July 2014 from three sites at and near the Hubbard Brook Experimental Forest (see Geographic Location, above). At each site, eight soil cores were collected from within a 900 m² plot; half (n = 4 cores per site) were randomly selected for the microbial analyses described here. The forest floor (Oe/Oa) was collected by removing loose leaf litter (Oi), then using a knife to collect a block of the Oe and Oa horizons from within a 15 x 15 cm wooden frame. A diamond bit rotary corer (Rau et al., 2011) with a 9.5 cm internal diameter enabled quantitative collection of the underlying mineral soil in 10 or 20 cm increments to 50 cm depth (0-10, 10-20, 20-30, and 30-50 cm). All analyses described here were performed on all 60 soil samples (3 sites × 4 cores per site × 5 depths per core), with minor exceptions noted below. Samples were stored on ice packs during transport and at 4 °C in the laboratory until processing. Soils were sieved within 24 hours of collection, using a coarse sieve (4 mm) to quickly homogenize the sample and remove large rocks and roots. Sub-samples of sieved soil were stored at -20 °C for later use in enzyme activity assays. The remaining soil was stored at 4 °C for up to 2 more days for the N cycling assays.

Bulk density and soil C and N stocks were determined from the quantitative soil samples following Rau et al. (2011). Briefly, 10 g subsamples of sieved soil were taken for moisture determination by drying for 1 day at 110 ^oC, and for C and N analysis after grinding to a fine powder with a ball mill (Retsch mixer mill MM200; Verder Scientific, Newtown, Pennsylvania, USA). Soil C and N concentration and isotopic composition were measured at the Cornell Stable Isotope Laboratory in Ithaca, NY, using a Finnigan MAT Delta Plus mass spectrometer following combustion with an elemental analyzer (Carlo Erba NC2500; Thermo Finnigan, San Jose, CA, USA). Surface mineral soil pH was measured with an Accumet AB15 pH meter (ThermoFisher Scientific, Waltham, MA, USA) with a 1:2 ratio of dry soil to water.

Rates of gross N mineralization and nitrification were assessed using the isotope pool dilution method (Davidson et al., 1991; Hart et al., 1994), in which a known amount of ¹⁵NH₄ ⁺ or ¹⁵NO₃ ^- is added to a soil sample, and its rate of dilution by mineralization or nitrification of unlabeled N is measured over 24 hours. To quantify rates of gross N mineralization and ¹⁵NH₄ ⁺ consumption, 7.53 μg N of 98 atom% (¹⁵NH₄)₂SO₄ were added to a pair of 15 g field-moist sub-samples, each in a 125 mL HDPE bottle. The label, along with 1 mL of deionized water, was distributed within each sub-sample using a 250 μg syringe. One sub-sample from each pair was extracted with 50 mL of 2M KCl within 15 minutes of labeling, and the second sub-sample was extracted after incubation in the dark for 24 hours at room temperature. The KCl extracts were stored in 60 mL HDPE bottles and frozen at -20 °C until chemical analysis. The same procedure was used to estimate rates of gross nitrification and ¹⁵NO₃ ^- consumption in another pair of subsamples using 1.63 μg N of 98% K¹⁵NO₃ ^- per subsample.

The KCl extracts were analyzed with colorimetric methods for NH₄ ⁺ (alkaline phenate) and NO₃ ^- + NO₂ ^- (cadmium reduction) concentrations using a Quikchem 8100 flow injection analyzer (Lachat Instruments, Milwaukee, WI, USA) at the Cary Institute of Ecosystem Studies in Millbrook, NY. The N diffusion method (Brooks et al. 1989) was used to determine ¹⁵N in the extract NH₄ ⁺ or NO₃ ^-. Extracts with small concentrations of extractable NH₄ ⁺ or NO₃ ^- were spiked with known amounts (50 or 100 μg N) of unlabeled NH₄ ⁺ or NO₃ ^- to ensure that each sample contained sufficient N for diffusion and ¹⁵N analysis. For the diffusions, two small glass fiber filters were acidified with 20 μL KHSO₄, then sealed in a Teflon tape packet that was floated on each KCl extract in a 125 ml HDPE bottle that was placed on a shaker table for seven days. MgO was added to all samples to increase pH and convert NH₄ ⁺ to NH_3, which is trapped on the acidified filters. Devarda’s alloy was also added to the ¹⁵NO₃ ^--labeled extracts to convert NO₃ ^- to NH₄ ⁺; filters in these extracts collected both NH₄ ⁺ and NO₃ ^-. Filter ¹⁵N contents were analyzed at the Cornell Stable Isotope Laboratory as described above. Pre-spike ¹⁵NH₄ ⁺ and ¹⁵NO₃ ^- extract concentrations were computed using mean ¹⁵N measurements of the unlabeled spike solutions, and assuming that ¹⁵N values of the unlabeled NH₄ ⁺ in the extracts for gross nitrification matched the natural abundance ¹⁵N measured in corresponding soil samples (Högberg 1997).

Gross N cycling rates were calculated using the differences in atom percent ¹⁵N enrichment above background (APE) and in inorganic N concentrations between the pre- and post-incubated samples using the equations in Hart et al. (1994), originally developed by Kirkham and Bartholomew (1954). Lack of ¹⁵N enrichment for four of the 240 diffusions prevented calculation of two each of the 60 gross mineralization and nitrification rates. One-day net N mineralization and nitrification rates were calculated using the pre- and post- incubation extractable inorganic N measurements collected during the gross N mineralization assay. Net N mineralization was calculated as ([NH₄ ⁺-N]+[ NO₃ ^--N])_incubation - ([NH₄ ⁺-N]+[ NO₃ ^--N])_initial and net nitrification was calculated as [NO₃ ^--N]_incubation- [NO₃ ^--N]_initial.

For the enzyme assay, two soil slurries were created for each soil sample, each using two grams of soil in 150 mL of a 50 mM sodium acetate buffer (pH 5.0) homogenized with a hand blender. One slurry was used for measurements of hydrolytic enzyme activity and the other for oxidative enzyme activity. Lack of material precluded analysis of oxidative enzyme activity for one sample at the old-growth site and eight samples at the mature site, including all four of its forest floor samples. For both sets of enzyme analyses, 50 µL of each sample slurry was added to 8 replicate wells in a column of a 96-well plate. Potential activities of six hydrolytic enzymes used for microbial acquisition of C-, N-, and P were quantified with fluorometric assays using the method outlined in German et al. (2011). These enzymes included two used to degrade cellulose, β-glucosidase (BG) and cellobiohydrolase (CB); one to degrade hemicellulose, β-xylosidase (BX); one to acquire phosphorus, acid phosphatase (AP); and two used to acquire N, NAG for amino sugars and LAP for amino acids. A fluorescent substrate specific to each enzyme function was added to the plate, which was then incubated in the dark at room temperature. Fluorescence was measured on a microplate reader set at 365 nm excitation and 450 nm emission. Assays were run alongside a standard curve containing soil homogenate with an increasing concentration of methylumbelliferone (MUB), except for the LAP analyses which used amidomethylcoumarin (AMC) standards (Table 2). Fluorescence was converted to units of potential enzyme activity per unit dry mass (nmol g^-1 h^-1) as in German et al. (2011). Potential activity of the lignolytic oxidative enzyme phenol oxidase (POX), was assessed using two different enzyme substrates, ABTS (2,20-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid)) and L-DOPA (L-3,4-dihydroxyphenylalanine), as suggested by Bach et al. (2013) on account of the variability of these assays. These activities are referred to here as POX_ABTS and POX_LDOPA, respectively. Plates were incubated in the dark for 24 hours at room temperature after adding the L-DOPA or ABTS substrates. Before measurement, 150 µL of each well was transferred onto a clear reading plate with transparent well bottoms.

Bach, C.E., Warnock, D.D., Van Horn, D.J., Weintraub, M.N., Sinsabaugh, R.L., Allison, S.D., German, D.P., 2013 .Measuring phenol oxidase and peroxidase activities with pyrogallol, L-DOPA, and ABTS: effect of assay conditions and soil type. Soil Biology Biochemistry, 67, 183-191.

Brooks, P.D., Stark, J.M., McInteer, B.B., Preston. T., 1989. Diffusion method to prepare soil extracts for automated N-15 analysis. Soil Science Society of America Journal 53, 1707-1711.

Davidson, E.A., Hart S.C., Shanks C.A., Firestone, M.K., 1991. Measuring gross nitrogen mineralization, immobilization, and nitrification by ¹⁵N isotopic pool dilution in intact soil cores, Journal of Soil Science 42, 335-349.

German, D.P., Weintraub, M.N., Grandy, A. S., Lauber, C.L., Rinkes, Z.L., Allison. S.D., 2011. Optimization of hydrolytic and oxidative enzyme methods for ecosystem studies. Soil Biology and Biochemistry 43, 1387-1397.

Hart, S.C., Nason, G.E., Myrold, D.D., Perry, D.A., 1994. Dynamics of gross nitrogen transformations in an old-growth forest: The carbon connection. Ecology 75, 880-891.

Högberg, P. 1997. Tansley Review No. 95: ¹⁵N natural abundance in soil-plant systems. New Phytologist 137, 179-203.

Kirkham, D., Bartholomew, W.V., 1955. Equations for following nutrient transformations in soil, utilizing tracer data. Soil Science Society of America Proceedings 19, 189-192.

Rau, B.M., Melvin, A.M., Johnson, D.W., Goodale, C.L., Blank, R.R., Fredriksen, G., Miller W.W., Murphy, J.D., Todd D.E., Walker, R.F., 2011. Revisiting soil carbon and nitrogen sampling: quantitative pits versus rotary cores. Soil Science 176, 273-279.