DOI: 10.5061/dryad.xpnvx0kp0
title: Data from: Nitrifier controls on soil NO and N2O emissions in three chaparral ecosystems under contrasting atmospheric N inputs

description: Nitrogen saturation theory predicts high rates of atmospheric N deposition
 can increase ecosystem N availability and stimulate ecosystem N losses via
 soil nitric oxide (NO; an air pollutant at high concentrations) and
 nitrous oxide (N2O; a strong greenhouse gas) emissions. However, it
 remains unclear whether theories developed in mesic ecosystems apply to
 drylands, where plant and soil N availability are not always coupled in
 dry soils. NO and N2O are often produced in soils during the oxidation of
 ammonia by ammonia oxidizing archaea (AOA) or ammonia oxidizing bacteria
 (AOB) during nitrification. AOB are thought to emit more NO and N2O during
 nitrification than AOA and may be favored in N-rich relative to N-limited
 environments, suggesting high rates of atmospheric N deposition might
 produce positive feedback sending more of the N to the atmosphere. To
 assess how high rates of atmospheric N deposition affect AOB- and
 AOA-derived N trace gas emissions, we selectively inhibited AOA and AOB
 nitrifiers and measured NO and N2O emissions from soils collected from
 three dryland sites exposed to relatively low (3.8 kg ha-1 = Low N) or
 high (11.8 kg ha-1 = High N1; 15.6 kg ha-1 = High N2) rates of atmospheric
 N inputs. We found that while the High N2 deposition site had the lowest
 AOA:AOB ratio (2.33 ± 0.57), consistent with expectations, this site did
 not emit the most NO and N2O. Rather, AOA emitted between 21–78% of the NO
 from our sites, with higher AOA-derived NO emissions from relatively
 coarse-textured soils in the Low N deposition site. In addition to
 nitrification, denitrification also contributed to NO and N2O emissions,
 especially in the Moderate N deposition site (where
 denitrification-derived NO and N2O emissions were 2.0 – 3.7 time greater
 than the other sites), which had finer textured soils that may favor
 denitrification. Interactions between soil texture and N availability,
 therefore, appears to be the primary mechanism determining whether
 atmospheric N deposition is retained in the ecosystem or reemitted to the
 atmosphere as NO or N2O.
description: Study Site We collected soils from three sites
 exposed to a range of atmospheric N deposition rates in southern
 California: a Low N deposition site and two high N deposition sites
 (hereafter, High N1 and High N2; Table 1; EPA, 2021). Vegetation in all
 sites was dominated by chamise (<em>Adenostoma
 fasciculatum</em>). Soils from the Low N site are fine sandy loams
 from the Sheephead series and are classified as shallow Entic
 Haploxerolls. Soils from the High N1 site are coarse loams from the
 Shepherdsaddle series and are classified as Ultic Haploxeralfs. Finally,
 soils from the High N2 site are fine sandy loams from the Trigo series and
 are classified as shallow Typic Xerorthents (Table S1; Soil Survey Staff,
 Natural Resources Conservation Service, United States Department of
 Agriculture, 2019). The climate at all sites is Mediterranean with hot,
 dry summers and cool, wet winters. Annual precipitation ranges from 500 to
 670 mm and average monthly temperatures range from 8 to 40°C.
 In the year leading up to the experiment, the High N2 deposition
 site had the highest ambient atmospheric concentrations of
 NO<sub>x</sub> (NO<sub>x</sub> = nitric oxide +
 nitrogen dioxide; 3.60 ± 1.51 ppm; Figure S1), while the High N1
 deposition site had the highest ambient atmospheric concentrations of
 ammonia (NH<sub>3</sub>; 2.98 ± 2.28 ppm), suggesting that
 they may receive different forms of N deposition. The three sites also
 have slightly different textures: the High N1 site is more enriched in
 clay (38.3 ± 2.57 %) compared to the Low N (20.4 ± 2.92 %) or High N2
 sites (15.8 ± 4.93 %; Table 1). Given that soil texture and the form of
 deposited N varied among the sites, we cannot isolate the effects of N
 deposition and, instead, aim to: i) understand what controls gaseous N
 losses from dryland soils and ii) whether NO and
 N<sub>2</sub>O emissions derived from AOB consistently
 associate with high atmospheric N inputs across remote sites varying in
 soil properties.  Experimental design
 At each of the three sites, we collected dry soils (0 – 10 cm
 depth; A horizons) from underneath five <em>Adenostoma fasciculatum
 </em>shrubs across a ~50 m transect (each shrub was roughly 10 m
 apart) in September 2021. We collected soils in September because this is
 near the end of the dry season at our sites, allowing us to assess the
 contribution of AOA and AOB to N emissions after experimentally rewetting
 soils in the lab. Soils were transported back to the lab where they were
 sieved to 2 mm and stored at 4 °C until the experiment began (soils were
 refrigerated for less than one month). Soils were removed from the
 refrigerator two days prior to beginning the experiment and were incubated
 at lab temperature (~22
 <em><strong>°</strong></em>C). Two soil samples
 were analyzed at a time. Each soil sample was split into three 50 g
 subsamples and transferred to 118 mL canning jars. The subsamples in each
 canning jar were exposed to one of three treatments: AOB inhibition, total
 nitrifier inhibition, or a control. To inhibit NH<sub>3</sub>
 oxidation by autotrophic AOB, 1-octyne was added to one subsample to bring
 the headspace in the jar to 4 µmol L<sup>-1</sup>. The
 1-octyne was prepared by adding 40 µL of liquid 1-octyne to a 125 mL
 bottle fitted with a butyl stopper, over pressurizing the bottle with 100
 mL of air, and, once the liquid 1-octyne evaporated, removing 2.7 mL of
 the bottle headspace to inject into the subsample. To inhibit
 NH<sub>3</sub> oxidation by all autotrophic nitrifiers,
 acetylene was added to one subsample to bring the jar headspace to 6 µmol
 L<sup>-1</sup>. The acetylene was prepared by first bubbling
 acetylene through a sulfuric acid trap to remove impurities, diluting the
 purified acetylene 10-fold with air, and then injecting 0.28 mL into the
 subsample. The third jar was treated as a control and was incubated under
 ambient lab conditions. All jars were incubated for 24 h.
 Following the 24 h incubation, NO and N<sub>2</sub>O
 emissions were measured from each jar after experimentally wetting soils.
 To prevent the growth of new AOB after wetting, the jar that was treated
 with 1-octyene was wet with a solution containing the antibiotic kanamycin
 at a concentration of 220 µg g<sup>-1</sup> soil. To prevent
 the growth of any new nitrifying bacteria or archaea, the jar that was
 treated with acetylene was wet with a solution containing kanamycin (220
 µg g<sup>-1</sup> soil), the archaeal protein synthesis
 inhibitor fuscidic acid (800 µg g<sup>-1</sup> soil), and the
 nitrification inhibitor nitrapyrin (200 µg g<sup>-1</sup>
 soil). The control jar was wet with deionized water only. All jars were
 wet with enough solution to reach 100% water holding capacity. NO and
 N<sub>2</sub>O emissions were measured from 6 of the jars
 (representing 2 samples) every 2 hours for 46 hours after wetting. Net
 nitrification and net N mineralization rates were measured as the
 difference in NO<sub>3</sub><sup>-</sup>
 (nitrification) or NO<sub>3</sub><sup>-</sup> and
 NH<sub>4</sub><sup>+</sup> (N mineralization)
 between the start and end of the 46-hour incubation. This process was
 repeated twice per week until all 45 jars were analyzed (3 sites   3
 treatments = 45 jars total). The contribution of AOB to
 NO and N<sub>2</sub>O emissions was determined by subtracting
 how much NO or N<sub>2</sub>O was emitted from the AOB
 inhibition treatment (i.e., treating soils with 1-octyne and kanamycin)
 from how much NO or N<sub>2</sub>O was emitted from control
 soils (wetted with water only). The contribution of AOA to NO and
 N<sub>2</sub>O emissions was determined by subtracting how
 much NO or N<sub>2</sub>O was emitted from the total nitrifier
 inhibition treatment (i.e., treating soils with acetylene, kanamycin,
 nitrapyrin, and fuscidic acid) from how much NO or
 N<sub>2</sub>O was emitted from the AOB inhibition treatment.
 Finally, heterotrophic NO and N<sub>2</sub>O emissions were
 assumed to be equal to how much NO or N<sub>2</sub>O was
 emitted from soils under the total nitrifier inhibition treatment;
 acetylene does not inhibit N<sub>2</sub>O production from
 heterotrophic denitrification. NO and
 N<sub>2</sub>O emissions            
 Immediately after wetting, six jars (2 samples x 3 treatments) were
 connected to a recirculating sample loop joining a multiplexer (LI-8150,
 LI-COR Biosciences), an infrared CO<sub>2</sub> analyzer
 (IRGA; LI-8100, LI-COR Biosciences, Lincoln, NE), and an
 N<sub>2</sub>O laser analyzer (Model 914-0027, Los Gatos
 Research, Inc., Mountain View, CA). To measure N<sub>2</sub>O
 emissions from each jar, air was recirculated through the closed sample
 loop at a rate of 1.5 L min<sup>-1</sup>. Soil
 N<sub>2</sub>O emissions were calculated as the linear change
 in N<sub>2</sub>O concentrations over a 9-minute period.
 Because the chemiluminescent NO<sub>2</sub> analyzer (LMA 3D;
 Unisearch Associates, Concord, ON, Canada) consumes NO, the
 NO<sub>2</sub> analyzer was not connected to the sample loop
 during this initial 9-minute incubation. Rather, after 9-minutes, an
 automated 3-way solenoid valve (Parker Hannifin Corp., Series 11/25/26,
 #991-000539-006) activated so that the NO<sub>2</sub> analyzer
 pulled from the sample loop at a rate of 1.5 L
 min<sup>-1</sup>. To replace the air that the
 NO<sub>2</sub> analyzer consumed from each jar (the air from
 the jars was vented out of the NO<sub>2 </sub>analyzer into
 the lab), a second valve activated at the same time to allow zero air
 (Ultra Grade Zero Air, Airgas, Radnor, PA) to enter each jar at a rate of
 1.5 L min<sup>-1</sup>. The NO<sub>2</sub>
 analyzer used a CrO<sub>3</sub> converter to oxidize NO to
 NO<sub>2</sub>; we did not detect NO<sub>2</sub>
 when CrO<sub>3 </sub>was removed from the sample loop
 suggesting our measurements were mostly NO. NO emissions were calculated
 using the following equation:  EQ1: NO flux =
 ([NO]outlet - [NO]inlet) x flow x mass N / soil wt. / R / temp
 Where [NO]<sub>outlet</sub> is the concentration of
 NO leaving the jar headspace, [NO]<sub>inlet </sub>is the
 concentration of NO entering the jar (assumed to be 0), flow is the flow
 rate of the sample loop (1.5 L min<sup>-1</sup>), mass N is
 the molar mass of N in NO, soil wt. is the mass of soil in the jar, R is
 the molar gas constant (0.0821 L atm K<sup>-1</sup>
 mol<sup>-1</sup>), and temp is the room air temperature. We
 measured NO concentrations for 10 minutes while zero air was flowing
 through the jar, allowing NO concentrations to reach equilibrium within
 the sample loop. NO fluxes were calculated using the average NO
 concentrations during the final 30 seconds of the 10-minute incubation.
 After 10 minutes, the sample loop was exposed to ambient lab air for one
 minute to purge zero air from the sample loop. After this 20-minute
 incubation, the multiplexer connected the next jar to the sample loop and
 this process was repeated, allowing us to measure each of the 6 jars once
 every ~2 hours. The continuous stream of dry air decreased soil moisture
 throughout the 46-hour incubation (Figure S2).
             We modified a publicly available script to calculate
 N<sub>2</sub>O and NO emissions
 (https://github.com/handr003/TraceGasArray). N<sub>2</sub>O
 emissions were calculated as the change in N<sub>2</sub>O
 concentrations over the last 7 minutes of the incubation when the NO
 analyzer was not connected to the sample loop. Emissions were considered 0
 if the linear relationship between time and N<sub>2</sub>O
 concentrations was not statistically significant (p &gt; 0.05). NO
 emissions were calculated using EQ1. Both the N<sub>2</sub>O
 and the NO analyzer recorded trace gas concentrations every second. We
 used the trapezoidal integration to calculate cumulative
 N<sub>2</sub>O and NO emissions over the 46-hour period
 post-wetting (final units were ng N g dry soil<sup>-1</sup>
 hr<sup>-1</sup>). Net nitrification and net
 N mineralization rates             We measured soil
 extractable NO<sub>3</sub><sup>-</sup> and
 NH<sub>4</sub><sup>+</sup> before wetting soils
 and immediately after the 46-hour incubation to calculate net
 nitrification and net N mineralization rates. Briefly, 3 g of soil (dry
 weight equivalent) were extracted in 30 mL 2M KCl for 1 hour, filtered
 (Whatman 42 filter paper; 2.5 µm pore size), and the extracts were then
 frozen until analysis. Extracts were analyzed for
 NO<sub>3</sub><sup>-</sup> (SEAL method EPA-136-A)
 and NH<sub>4</sub><sup>+</sup> (SEAL method
 EPA-129-A) using colorimetric assays. Net nitrification and net N
 mineralization rates were calculated as the difference in inorganic N
 (NO<sub>3</sub><sup>- </sup>for nitrification and
 NO<sub>3</sub><sup>- </sup>plus
 NH<sub>4</sub><sup>+</sup> for mineralization)
 before and after the incubation divided by the length of the incubation
 (~46 hours). We also measured soil gravimetric water content by drying
 soil samples (~10 g) at 104 °C for 24 hours. We estimated 100% soil water
 holding capacity (WHC) as the amount of water held by soils after
 saturating them with water and allowing them to drain in an air-tight
 container (to limit evaporation) for 8 hours.  
 <em>amoA</em> gene quantification
             A subsample (~5g) of each soil sample (n = 15) was
 frozen (-20 °C) as soon as soils arrived in the lab. Within one month of
 freezing, DNA was extracted from 0.25g of each subsample using a DNA
 extraction kit (Qiagen DNeasy PowerSoil Pro, Hilden, Germany) following
 the manufacturer’s guidelines after an overnight incubation to enhance DNA
 extraction (700µl CD1 + 100µl ATL at 4°C; Qiagen). Quantitative polymerase
 chain reaction (qPCR) was used to estimate the abundance of bacterial and
 archaeal <em>amoA </em>genes; the AmoA1F/amoA2R primer set was
 used for bacteria and the Arch-amoAF/ArchamoAR primer set was used for
 archaea. Each qPCR was run in 10 µL reactions containing 5 µL master mix
 (Forget-Me-Not EvaGreen qPCR Master Mix, Biotium, Inc., Fremont, CA), 0.8
 µL of 2 mM MgCl<sub>2</sub>, 0.25 µL of 0.5 mg ml<sup>-1
 </sup>bovine serum albumin, 0.125 µL of 0.25 µM forward and reverse
 primer, 2.5 µL H<sub>2</sub>O, and 1.2 µL sample DNA.
 Bacterial <em>amoA</em> was amplified using the following
 protocol: 5 minutes at 95 °C, followed by 40 cycles of 45 seconds at 95
 °C, 30 seconds at 56 °C and 60 seconds at 72 °C (CFX384 Touch Real-Time
 PCR Detection System, Bio-Rad, Hercules, CA). Archaeal
 <em>amoA</em> was amplified using the following protocol:
 4 minutes at 95 °C, followed by 40 cycles of 30 seconds at 95 °C,
 45 seconds at 53 °C and 60 seconds at 72°C. The standard sequences were
 chosen from well-known archaeal (crenarchaeota genomic fragment 54d9) and
 bacterial (<em>amo</em>A gene of <em>Nitrosomonas
 europaea</em> ATCC 19718) ammonia oxidizing microorganisms. Standard
 curves were prepared using serial dilutions for bacterial
 <em>amoA</em> (10<sup>6</sup> to
 10<sup>2</sup> copies) and archaeal <em>amoA</em>
 (10<sup>7</sup> to 10<sup>3</sup> copies). The
 bacterial <em>amoA </em>standards had efficiencies of 83.5%
 (R<sup>2</sup> = 0.998) and archaeal <em>amoA</em>
 standards had efficiencies of 66.9% (R<sup>2</sup> =
 0.997).
description: # Data from: Nitrifier controls on soil NO and N2O emissions in three
 chaparral ecosystems under contrasting atmospheric N inputs
 [https://doi.org/10.5061/dryad.xpnvx0kp0](https://doi.org/10.5061/dryad.xpnvx0kp0) This dataset includes the data presented in the manuscript: "Nitrifier controls on soil NO and N2O emissions in three chaparral ecosystems under contrasting atmospheric N inputs". Briefly, we collected soils from the chaparral ecosystems in southern California and measured NO and N2O emissions from nitrification and denitrification. We treated soils with nitrification inhibitors to assess the relative contribution of ammonia-oxidizing bacteria (AOB) versus ammonia-oxidizing archaea (AOA) to N trace gas emissions. We also measured inorganic N pools to asses net rates of nitrification and net N mineralization. All soils were wet to 100% water holding capacity and were allowed to dry during the 46-hour incubation period. Finally, we present data on the abundance of AOA and AOB nitrifiers as measured on dry soils using quantitative polymerase chain reaction. ## Description of the data and file structure Combined_Fluxes.csv: This file contains the soil nitric oxide (NO) and nitrous oxide (N2O) flux data from soils wetted to 100% water holding capcity in a lab incubation. Methods for flux measurements and flux calculations are described in the manuscript text. Moisture.csv: This file contains soil moisture data that are presented in the manuscript. Net_N_rates.csv: This file contains initial soil nitrate and ammonia concentrations as well as net rates of nitrification and nitrogen mineralization from the three sites examined in this study. The treatments refer to lab treatments used to selectively inhibit ammonia oxidizing bacteria or all heterotrophic nitrifiers. Empty cells indicate missing data. qPCR.csv: This file contains data on the abundance of ammonia oxidizing archaea (AOA) and ammonia oxidizing bacteria (AOB) in dry soils collected from our three sites. The abudnance of AOA and AOB was assessed using quantitative polymerast chain reaction (qPCR).