Abertura

Substrates were collected from two locations: sediments from Harts Creek, a shallow spring- fed, gaining stream in the Canterbury plains region of New Zealand (43°46’S, 172°16’E), and

submerged paddy soils from the International Rice Research Institute experimental farm (IRRI) in the Philippines (14º1’N, 121º15’E). Locations were selected to provide a snapshot of global (cross-biome) v. local (km) scale variation in freshwater εdenit. Within Harts Creek, multiple sites were sampled to span variations in water depth, temperature, and organic matter content, factors known to influence

kdenit (Garcia-Ruiz et al. 1998, Alexander et al. 2000) and denitrifier activity and abundance (Findlay 2010, Findlay et al. 2011). Sites within IRRI were selected based on changes in land-use / cultivation history, known to control kdenit and denitrifier community dynamics in submerged soils (Buresh et al. 2008, Bannert et al. 2011, Ahn et al. 2012).

Sample collection

In Harts Creek, streambed sediments were collected from four reaches at increasing distance from the source spring: 1.69 km (A), 3.69 km (B), 7.75 km (C), and, near the river mouth, 9.61 km (D). Prior to sediment collection, dissolved oxygen (DO) and water temperature were measured in-situ at each reach using a portable hand-held meter (550A YSI, Yellow Springs, OH). Stream depth was measured using a metre stick at upstream sites (A and B) and installed gauges at downstream sites (C and D). Harts Creek climate conditions were evaluated using data from the Leeston Harts Creek

climate station (43°46’1.99”S, 172°16’57.79”E) (<1 km from site A) (CliFlo: NIWA’s Climate Database Online (http://cliflo.niwa.co.nz). Data retrieved 06-Jun-2011.)

Paddy soils were collected from three fields with different cultivation histories and clay contents (which reflects cultivation intensity (Kogel-Knabner et al. 2010)) at IRRI: E (two irrigated rice crops a year for >20 years, 61% clay), F (zero to two irrigated rice crops a year for ~10 years, 33% clay), and G (uncultivated soil within 500 m of E). Floodwater depth and soil temperature were recorded prior to sampling. Climate data was obtained from the on-farm climate station and soil characteristics from survey data (W. Lorenzo and R.J. Buresh, unpubl.).

At each location, 20 sediment samples (10 cm depth x 5 cm diameter) were collected (locations determined using a pre-prepared randomised grid) and bulked together to ensure that samples were representative of each site (Bissett et al. 2010). Samples were stored on ice until returned to the lab (<2 h), whereupon they were sieved through a 0.5 mm mesh to remove larger particles and soil fauna. Sub-samples (~50 g) from each location were stored at 4ºC until chemical analyses (water-holding capacity (WHC), total organic carbon (TOC), pH, and total C and N).

Intrinsic ε

incubations

Streambed sediments were incubated in 30 ml amber glass bottles (Alcom) sealed with Teflon- lined rubber septa. Three replicate incubations were made for each sampling time x treatment, plus an additional control. Each bottle was filled with 25 ml of sediment slurry (100% WHC), which were kept continuously mixed (i.e., no diffusive limitation to denitrification) using a rotary shaker table. For each paddy soil, the equivalent of 200 g dry weight was added to a 2 l mesocosm Pyrex jar and brought up to 1 kg with deionised H2O (volume selected to ensure that the substrate surface area did not change significantly over time with sub-sampling), sealed, and then kept continuously mixed using a magnetic stirrer. Two replicate mesocosms were set-up for substrate from each site, each fitted with two platinum electrodes and one hydrogen probe for redox measurements (using an Ag/AgCl

reference electrode), which were corrected based on hydrogen probe readings (simultaneously used to measure soil pH) (as per Johnson-Beebout et al. (2009)).

All incubations were continuously flushed with N2 or He gas, and outflows from the

incubations were bubbled through air-tight 12 ml Exetainer® filled with 10 ml of deionised water to ensure no air backflow. After 48 h of pre-incubation, C (as glucose; 1mM) and NO3- _{(25 mg N l}-1 _as KNO3) were added to treatment incubations to ensure that the initial denitrification period was not substrate limited. Slurries were collected +1, 3, 6, 12, 24, and 36 h after substrate additions. For the Harts Creek sediments, sampling was destructive, consisting of opening mesocosms within an air-free bag (purged with He), where the slurries were transferred into a sterile 50 ml centrifuge tube for extraction. For paddy soils, three 50 ml sub-samples were collected from each mesocosm using a sterile syringe. These slurries were immediately injected into acid-washed centrifuge tubes. Once sediments and soils were in the centrifuge tubes, deionised water was added at a 5:1 w/w ratio and samples were centrifuged at 3500 rpm for 20 min, and supernatents pumped through GF/F (Whatman)

filter paper to remove particulates. Air temperature was measured continuously, and paddy slurry redox potential and pH were recorded at each sampling interval.

Layered incubations (ε

)

Site C sediments were also used in incubations testing the impact of diffusion and nitrification on εeff. The sediments were placed in sterile 30 ml plastic vials (2.5 cm diameter x 6.0 cm height) and then brought to 100% WHC, and then overlain with either 0 cm (L0), 2 cm (L1) or 4 cm (L2) of acid- washed quartz sand plus an additional 6 ml of deionised water. Treatments were replicated four times (including one control without added N or C) for each sampling interval. After 48 h, NO3- _{(25 mg N l}-1 as KNO3) and C (1 mM glucose) were slowly injected into the base of the sediment layer using a 1 ml syringe fitted with a 23 g (0.337 mm) needle. Incubations were destructively sampled 4, 8, 24, 48, and 96 h after substrate additions: surface water (SW) was removed using a 15 ml sterile syringe, sediment porewater (PW) extracted as in previous incubations, and both SW and PW samples passed through GF/F filter paper (Whatman).

Vertical profiles (500 μm depth intervals) of O2 and N2O concentrations in the sediments, sand, and water, were measured following the procedure of Elberling et al. (2010). Briefly, O2 was determined using a miniaturised Clark-type O2 micro-sensor (OX10, Unisense, Science Park, DK- 8000 Aarhus, Denmark) equipped with an internal reference and a guard cathode (linearity was confirmed by recording the output current (pA) in water sparged with N2, air and pure O2). Standard micro-sensors were used to measure N2O concentrations and substrate diffusivity (Elberling et al. (2010) and references therein).