Soil drying–wetting cycles are widespread and can stimulate large emissions of both nitrous oxide (N2O) and nitric oxide (NO) (Scholes et al. 1997; Homyak et al. 2016; Eberwein et al. 2020) with profound implications for Earth’s climate, regional air quality, and ecosystem N retention. This is because N2O is a potent greenhouse gas (Ciais et al. 2013), NO is a precursor to tropospheric ozone (Crutzen 1979), and both NO and N2O represent important pathways for ecosystem N loss (Peterjohn and Schlesinger 1990). While the production of NO and N2O is governed by both biological and chemical processes upon wetting dry soil, the magnitude of the emissions vary as a function of aridity (Wang et al. 2014; Liu et al. 2017; von Sperber et al. 2017), with some drylands recording among the highest NO and N2O emission pulses globally (Eberwein et al. 2020). However, how these emissions vary across ecosystems experiencing drying–wetting cycles and the biogeochemical processes producing them are still not well characterized. Identifying the underlying processes producing pulsed NO and N2O emissions is necessary to predict how ecosystem N cycling may respond to global change factors including high rates of atmospheric N deposition (Fenn et al. 2006), rising temperatures, and changing precipitation regimes (Dai 2013).

Multiple biological and abiotic processes regulate NO and N2O emissions after dry soils are wetted. Biological processes include nitrification, the aerobic oxidation of ammonia (NH3) to nitrate (NO3) with NO and nitrite (NO2) as intermediates (Caranto and Lancaster 2017; Prosser et al. 2019), and denitrification, the sequential anaerobic reduction of NO3 to N2 gas with NO2, NO, and N2O as obligate intermediates (Knowles 1982); both of these processes can release NO and N2O as byproducts. In low oxygen environments, some nitrifiers use NO2 as the electron acceptor during the oxidation of NH3 and produce NO and N2O via nitrifier denitrification (Prosser et al. 2019). Chemodenitrification—an abiotic non-enzymatic process—can also produce NO and N2O through the chemical reduction of NO2 and hydroxylamine (NH2OH) (Venterea and Rolston 2000; Zhu-Barker et al. 2015; Heil et al. 2016), which can accumulate in dry soils (Homyak et al. 2016). Because both biological and abiotic processes can occur simultaneously, it has been challenging to determine the contribution of individual processes to pulsed N emissions.

To advance understanding of the processes producing NO and N2O, the “hole-in-the-pipe” conceptual framework relates the factors that control N emissions to the processes that control N transformation rates (Firestone and Davidson 1989). Under this framework, N transformations are represented by changes in the diameter of the pipe—the diameter varies in proportion to process rates—whereas the factors controlling how much NO or N2O leak out of the pipe are represented by the holes (e.g., edaphic or environmental factors such as pH). In this sense, wetting soils could stimulate NO and N2O emissions by promoting nitrification, (Placella and Firestone 2013; Homyak and Sickman 2014), denitrification (Parker and Schimel 2011; Soper et al. 2016), or abiotic reactions (McCalley and Sparks 2009; Zhu-Barker et al. 2015; Homyak et al. 2017), thereby increasing the diameter of the pipe. However, the magnitude and timing of pulsed N emissions may vary as a function of environmental and edaphic factors that mediate which gaseous N intermediates are released to the atmosphere (i.e., the holes in the pipe). Understanding how process rates interact with the factors that control how much NO and N2O are emitted can help determine how N emissions may vary under future global change scenarios.

Two major challenges have limited progress identifying controls over soil NO and N2O emission pulses: (i) multiple biological and abiotic processes occur simultaneously making them difficult to separate, and (ii) traditional static chamber headspace experiments offer low temporal resolution, limiting understanding of the timing and magnitude of N trace gas emissions. To this end, isotope tracers are powerful tools that can help determine which N transformations produce NO and/or N2O (Van Groenigen et al. 2015). Moreover, isotope tracers can be coupled with laser-based isotope analyzers and automated soil chambers to detect the incorporation of 15N tracers into N2O in-situ and at high resolution (e.g., one measurement per second). While similar instruments do not yet exist for NO, the incorporation of 15N tracers into NO can be detected using passive samplers (Homyak et al. 2016). By pairing high temporal resolution measurements of N emissions with stable isotopes, we assess the importance of increasing N availability (here used as a proxy for increasing the diameter of the pipe) relative to the factors that control how much NO and N2O is released (i.e. the holes in the pipe). Specifically, we ask: (1) what processes are contributing to pulsed NO and N2O emissions after wetting dry soils, and (2) how does N availability (both the amount and chemical form) affect the magnitude of pulsed emissions?

To answer these questions, we monitored N emissions at two dryland sites (desert and chaparral) in Southern California characterized by pronounced and frequent transitions from dry-to-wet soils. We chose two sites with contrasting environmental conditions (Table 1) to understand whether meteorological and edaphic factors would overrule the effects of increasing N supply and the form of nitrogen added, nitrate (NO3) or ammonium (NH4+). We hypothesized that N trace gas emissions are limited by soil N availability, resulting in pulsed NO and N2O emissions proportional to the amount of added N. To infer which processes contributed to NO and N2O emissions, we added 15N labeled NO3 or NH4+ and used an automated chamber system connected to a NO and an isotope N2O analyzer. We also measured NH3 emissions using passive samplers as a relative index of the amount of NH3 in soil pore space that may be available to nitrifiers. We predicted that if pulsed N emissions were from nitrification, then added 15N–NH4+ would be captured as NO and/or N2O; if they were from denitrification, then added 15N–NO3 would be captured as NO and/or N2O; and if they were from the rapid transformation of accumulated nitrification intermediates (e.g., NO2), then no 15N label would be incorporated in N emissions.

Table 1 Soil chemical properties and meteorology prior to beginning experiments in the desert and chaparral sites (n = 8)


Sites description

We studied two drylands in Southern California in August 2018 (the end of the summer dry season) with contrasting soils and vegetation. Our chaparral site was located in the Box Springs Reserve (33° 58′ 16.4″ N, 117° 17′ 53.4″ W), a transitional zone between coastal sage scrub and chaparral dominated by chamise (Adenostoma fasciculatum). Our desert site was located in the Boyd Deep Canyon Desert Research Center (33° 38′ 54.7″ N, 116° 22′ 39.4″ W), and was dominated by creosote (Larrea tridentata). Both sites are part of the University of California Natural Reserve System. Since 1980, the chaparral site has received an average of 28 cm of rain per year with an average maximum August daily temperature of 35 °C. During this same time, the desert site received an average of 11.4 cm of rain per year with an average maximum August daily temperature of 39.3 °C. The chaparral soils are sandy loams classified as thermic Typic Haploxeralfs within the Fallbrook series. The desert soils are stony sands classified as hyperthermic Typic Torriorthents within the Carrizo series. Both sites received no rain in the month before our experiments.

The soils at the two sites differed in several ways (Table 1). Soil NO2 was over seven times greater in the desert (0.58 ± 0.64 µg N g−1) than in the chaparral (0.08 ± 0.03 µg N g−1, p < 0.05), while extractable NO3 and NH4+ did not differ between sites (Table 1). Total C and N concentrations were both higher in the chaparral (2.03 ± 0.35% C, 0.15 ± 0.02% N) than in the desert (0.92 ± 0.50% C, 0.08 ± 0.03% N). Desert soils were more alkaline (8.4 ± 0.19) than the chaparral (5.8 ± 0.50, p < 0.05). In the hour before starting our experiment, relative humidity was higher in the chaparral (80.7 ± 15.0%) relative to the desert (10.7 ± 1.04%, p < 0.001), while soil temperature was higher in the desert (30.0 ± 0.68 °C) than in the chaparral (16.5 ± 2.15 °C, p < 0.001).

Experimental design

We measured N trace gas emissions from underneath eight chamise shrubs in the chaparral and eight creosote shrubs in the desert. Interspace soils were not sampled as dryland shrubs are considered to be “islands of fertility” where soil nutrients are concentrated (Schlesinger et al. 1990). All shrubs were located within a 10-m radius and were separated from one another by at least one meter. Under each of the eight shrub canopies, we installed two pairs of PVC collars (4 collars, each 20 cm diameter × 10 cm height; inserted 5 cm into the ground) at least 48 h prior to the start of our measurements; the collar pairs were separated from each other by at least 50 cm to avoid cross contamination of isotope tracer and within 50 cm from the base of the shrubs. One pair of collars was wetted with NO3 solution, while the other was wetted with NH4+ solution. Within each pair, one collar was used to measure N emissions, while the other was used to measure soil temperature, moisture, and inorganic N to minimize disturbances to the collars from which we measured emissions.

We wetted soils inside the collars with 500 mL of deionized water; this amount corresponds to about seven mm of rainfall, which is within the range of historically occurring rain events (Boyd Deep Canyon Desert Research Station, During wetting, we added eight levels of N spike corresponding to 0, 2, 4, 6, 8, 10, 12, or 15 kg-N ha−1 as either NO3 or NH4+, covering a range of annual N deposition in Southern California drylands (Eberwein et al. 2020, Fenn et al. 2006). The nitrogen added was isotopically enriched to 2 atom percent 15N. The labeled NO3 was added to two of the collars underneath each shrub starting at approximately 9 am. Soil NO and N2O emissions were measured from one collar underneath each shrub every 30 min beginning 15 min prior to wetting. After 24 h, this process was repeated with the NH4+ label using the remaining collars underneath each shrub.

A separate group of four shrubs was used to measure the emission of NH3 as well as the isotopic composition of NO. Emissions of NH3 were used as an index of substrate availability to nitrifiers. These measurements were made using passive samplers (Ogawa pads; Ogawa USA, Pompano Beach, FL) that required soil chambers to be permanently closed, prohibiting integration with our automated chambers. The passive sampling pads are chemically pretreated so that they would collect either NOx, NO2, or NH3 and have been demonstrated to work well under warm and humid conditions expected inside our soil chambers (Coughlin et al. 2017). We did not detect NO2 on the NO2 pads, indicating any N accumulation on the NOx pads was mostly NO. Two collars underneath each of the shrubs were wetted with 500 mL of either NO3 or NH4+ solution (2 atom percent 15N) at a concentration corresponding to 15 kg-N ha−1. The remaining two collars underneath each of the four shrubs were wetted with deionized water only. Chamber lids were installed immediately after wetting and pads were switched out at the following time intervals: 0 to 15 min, 15 min to 12 h, and 12 to 24 h post-wetting to capture NO and NH3 during periods when we expected N emissions to be high.

NO and N2O emissions

We used an automated chamber system to simultaneously measure NO and N2O emissions from one of the collars under each of the eight shrubs sequentially over a 24-h period post-wetting. Collars were equipped with automated chambers (8100-104/C, LI-COR Biosciences, Lincoln, NE) connected to a multiplexer (LI-8150, LI-COR Biosciences) to sequentially measure emissions from each of the eight collars. We measured gas concentrations for two minutes, during which time gas from the chamber was recirculated through a sample loop connecting the multiplexer, an infrared gas analyzer (IRGA; LI-8100, LI-COR Biosciences), an isotope N2O analyzer (Model 914-0027, Los Gatos Research, Inc., Mountain View, CA), and a NO analyzer (Model 410 and Model 401, 2B Technologies, Boulder CO). The IRGA, N2O analyzer, and NO analyzer all sampled air from the recirculating sample loop, and each instrument, except for the NO analyzer, returned air back into the sample loop. Since the NO analyzer consumed NO, this air was vented to the atmosphere at a rate of 0.75 L min−1. While this open system dilutes the concentration of trace gases emitted from the soil with atmospheric air, flux rates are not appreciably affected after accounting for our chamber volume (~ 6 L) and the short incubation period (Davidson et al. 1991). All instruments were housed inside an air-conditioned box made of five cm thick housing insulation (5 × 2 × 2 m). To prevent condensation in the lines, the sample loop included a water trap to remove moisture by cooling the hoses with ice water. Soil temperature (Model 8150-203, LI-COR Biosciences) and moisture sensors (Model 8150-205, LI-COR Biosciences) were installed under each shrub and were connected to the IRGA, which also measured relative humidity.

Fluxes of NO and N2O were calculated as the linear change in trace gas concentrations inside the chamber headspace over the last 90 s of the two-minute incubation (script available on This timeframe was chosen to allow for even mixing of chamber air throughout the sample loop. The N2O analyzer recorded concentrations once every second and the NO analyzer recorded every ten seconds. If the linear correlation between time and trace gas concentration was not statistically significant (p > 0.1), the net flux was reported as zero. The change in NO concentration over time was highly linear over the 90 s window for all measurements (R2 = 0.96). The change in N2O concentration over time was close to linear for all measurements (R2 = 0.56) and was highly linear when N2O fluxes were greater than 10 ng N–N2O m−2 s−1 (R2 = 0.97).

Flux values were corrected for the volume in the sample loop, soil temperature, and chamber volume. The isotopic N2O analyzer also recorded [δ15N]N2O, which requires five minutes of averaging time to report δ15N values within 1-sigma precision. Given the short incubation period of our measurements (2 min) and the fact that our measurements were diluted with ambient air, we do not attempt to calculate absolute [δ15N]N2O values. Rather, we report *[δ15N]N2O as an index of when 15N tracer was incorporated into N2O after wetting dry soils. *[δ15N]N2O was calculated as the average δ15N value during the final 10 s of each incubation—across all measurements the standard deviation of [δ15N]N2O during this 10 s interval averaged 4.95 ‰. We also refrain from reporting isotopomer values for these same reasons—two-minute chamber closures were not sufficient to ensure isotopic accuracy and precision. The isotope N2O analyzer was referenced against a commercially available standard (Airgas, 5000 ppm N2O, δ15N = − 0.3 ‰) and a cylinder of medical grade air analyzed for N2O and isotopic composition at the UC Davis Stable Isotope Facility (0.44 ± 0.02 ppm N2O; δ15N = 5.76 ± 0.15 ‰).

NO isotopes and soil NH3 emissions

We used the bacterial denitrifier method to measure the [δ15N]NO and [δ18O]NO of NO captured on the NOx pads (Coplen et al., 2012). Briefly, the Ogawa pads were extracted in 8 mL of deionized water and shaken overnight to extract NO as NO2; no NO3 was detected in the filtered extracts. The NO2 was then converted to N2O using Pseudomonas aureofaciens (Sigman et al. 2001). δ15 N and δ18O values were measured using a Thermo Delta V isotope ratio mass spectrometer (Thermo Fisher Scientific, Woltham, MA) at the Facility for Isotope Ratio Mass Spectrometry (FIRMS; at the University of California, Riverside. Due to isotopic fractionation associated with NO collection with passive samplers, isotopic fractionation associated with the denitrifier method, exchange of oxygen atoms between NO2 and water (Casciotti et al. 2007; Dahal and Hastings, 2016; Yu and Elliott 2017), and potential interactions between volatile organic compounds (VOC) with NO (Walters and Michalski 2016) it is unlikely we measured the actual [δ15N]NO and [δ18O]NO emitted from soil. However, these fractionation and oxygen exchange effects are generally uniform across samples (Dahal and Hastings, 2016) and even if NO and VOCs interacted within our chambers, the passive samplers can still inform when 15N tracers added to soils are detected as NO (Homyak et al. 2016), altogether helping to preserve a [δ15N]NO and [δ18O]NO signal from the Ogawa pads, hereafter referred to as *[δ15N]NO and *[δ18O]NO. To help preserve the [δ18O]NO signal from the Ogawa pads, we used the same water source to prepare all isotope tracers and analyzed all samples in a single batch. Furthermore, our samples did not have NO3—the NO was extracted as NO2—reducing bias in the final isotope measurement (Casciotti et al. 2007).

We used Ogawa pads to measure NH3 emissions as an index of NH3 availability in soil pore space that may be available to nitrifiers. The NH3 pads were extracted in 8 mL of deionized water and shaken overnight to extract NH3 as NH4+. Extracted NH4+ was quantified using a colorimetric assay (SEAL methods Environmental Protection Agency (EPA)-126-A) using a SEAL AQ-2 discrete analyzer (SEAL analytical, Mequon, WI).

Soil chemical properties

We measured soil extractable NH4+ and NO3 prior to wetting, two hours after wetting, and 24 h after wetting. NO3 and NH4+ were measured by extracting soils (5 g) in 2 M KCl (30 mL). Soil solutions were shaken for one hour, filtered (Whatman 42 filter paper; 2.5 µm pore size), and frozen until analysis. We also measured NO2 prior to wetting; NO2 was extracted in deionized water to minimize its loss via gaseous N products (Homyak et al. 2015). We used colorimetric assays to measure soil extractable NH4+ (SEAL method EPA-126-A), NO3 (SEAL method EPA-129-A), and NO2 (SEAL method EPA-137-A). Additionally, we measured total C, total N, and pH in dry soils (0–10 cm depth) collected from underneath each shrub prior to adding water or N. Soil total C and total N was measured in an elemental analyzer (Flash EA1112; Thermo Scientific, Woltham, MA) at the Environmental Sciences Research Laboratory at the University of California, Riverside ( Soil pH was measured in a 1:1 soil to water ratio with a pH meter (Orion VersaStar Pro; Thermo Scientific, Woltham, MA).

Statistical analyses

All statistics were conducted in R version 3.6.1 (R core development team, 2019). We used linear regression to evaluate the relationship between the amount of added N and soil NO emissions, N2O emissions, and peak *[δ15N]N2O. This was accomplished by first calculating the cumulative NO or N2O emissions measured at each shrub using the “trapz” function. Peak *[δ15N]N2O was calculated as the highest *[δ15N]N2O value recorded from underneath each shrub. We then used the “lm” function to determine the linear relationship between the amount of added N and cumulative NO or N2O emissions and peak *[δ15N]N2O. We report the R2 of each linear regression where p < 0.10 to avoid type II error associated with high spatial variation in field experiments. However, we consider linear regressions with p > 0.05 as “weak” and include alternative explanations for these relationships. A block in the sample loop prevented us from measuring fluxes from two of the collars in the chaparral (2 and 10 kg-N NO3 ha−1) and these data were omitted from our analyses.

We used mixed effects models to determine when 15N tracers were detected in NO collected using passive samplers. The models included *[δ15N]NO as the response variable, collection time as the predictor variable, and a random effect to account for measuring the same collar repeatedly. Models were run using the “nlme” package in R. We used the anova.lme function to determine if time was a significant model term and Tukey corrected multiple comparisons to determine which times differed compared to ambient *[δ15N]NO. We used the same approach to determine if *[δ15N]NO and *[δ18O]NO changed in collars that were amended with water only.


Soil N2O emissions

In the desert, peak N2O emissions averaged 529 ± 469 ng N–N2O m−2 s−1 after wetting soils with NO3 and NH4+ amended solutions, and returned to prewetting levels within four hours (Fig. 1a,c). However, desert N2O emissions did not increase in proportion to adding more NO3 (p = 0.12) or NH4+ (p = 0.89, Table 2). In contrast to the desert, peak chaparral N2O emissions averaged only 38.0 ± 72.0 ng N–N2O m−2 s−1 after wetting with NO3 and NH4+amended solutions (Fig. 1b,d). As observed in the desert, N2O emissions did not increase in proportion to adding NO3 (p = 0.25) or NH4+ (p = 0.10, Table 2).

Fig. 1
figure 1

Soil N2O emissions (ng N–N2O m−2 s−1) over 24 h from the desert (a, c) and chaparral (b, d) following wetting of dry soils with nitrate (NO3a, b) or ammonium (NH4+c, d) solutions. Each black dot represents flux measurements over a 2-min interval for each of the chambers

Table 2 Linear relationship between the added amount of NO3 or NH4+ and cumulative N2O or NO emissions over the course of 24-h from either the desert or chaparral

Rapid reduction of NO3 produced pulsed N2O emissions at both sites. In response to 15 kg ha−1 equivalent NO3 addition, *[δ15N]N2O reached 1953 ‰ in the desert and 124 ‰ in the chaparral (Fig. 2a,b), whereas the *[δ15N]N2O of from soils amended with water only did not surpass 32.4 ‰ at either site. Peak *[δ15N]N2O increased in proportion to NO3 addition in the desert (R2 = 0.62, slope = 107 ‰ (kg N ha−1)−1, p = 0.013) and to a smaller degree in the chaparral (R2 = 0.31, slope = 6.2 ‰ (kg N ha−1)−1, p = 0.089, Table S1). *[δ15N]N2O remained under 75 ‰ in NH4+-amended soils at both sites (Fig. 2c,d). Peak *[δ15N]N2O was positively correlated to NH4+ addition in the desert only (R2 = 0.50, slope = 2.85 ‰ (kg N ha−1)−1, p = 0.03), but the slope of this relationship was over 37 times smaller compared to NO3 addition (Table S1).

Fig. 2
figure 2

Isotopic composition (*[δ-15N]N2O) of N2O emitted over 24 h from the desert (a, c) and chaparral (b, d) following wetting of dry soils with nitrate (NO3a, b) or ammonium (NH4+c, d) solutions. Each black dot represents the average isotopic composition of N2O measured over the last 30 s from each chamber

Soil NO and NH3 emissions

Prior to wetting, NO emissions were greater in the chaparral (14.7 ± 4.31 ng N–NO m−2 s−1) than in the desert (0.39 ± 9.27 ng N–NO m−2 s−1) (Table 1). In the desert, NO emissions steadily increased for 10 h post-wetting, and remained elevated for the remainder of the experiment (Fig. 3a); peak NO emissions averaged 221 ± 269 ng N–NO m−2 s−1 in NO3-amended soils (Fig. 3a) and 254 ± 200 ng N–NO m−2 s−1 in NH4+-amended soils (Fig. 3c). In contrast to the desert, chaparral NO emissions reached their peak within only 5 h of wetting and decreased at faster rates; NO emissions averaged 114 ± 204 ng N–NO m−2 s−1 in NO3 amended soils (Fig. 3b) and 202 ± 154 ng N–NO m−2 s−1 in NH4+ amended soils (Fig. 3d).

Fig. 3
figure 3

Soil NO emissions (ng N–NO m−2 s−1) over 24 h from the desert (a, c) and chaparral (b, d) following wetting of dry soils with nitrate (NO3a, b) or ammonium (NH4+c, d) solutions. Each black dot represents flux measurements over a 2-min interval for each of the 8 chambers

In contrast to N2O, isotopically labeled NH4+ and NO3 were both incorporated into the NO emitted at both sites. The 15N labeled NO3 was rapidly converted to NO in the chaparral (F3,9 = 93.8, p < 0.0001), enriching *[δ15N]NO from − 13.2 ± 1.82 ‰ to 388 ± 27.8 ‰ within 15 min of tracer addition (p < 0.0001, Fig. 4b). In the desert, the 15N–NO3 label was detected in NO (F3,9 = 1.7, p = 0.001) but not at 15 min (*[δ15N]NO = 23.1 ± 7.33 ‰, p = 1.00); it was detected between 0.25 and 12 h, when *[δ15N]NO reached 745 ± 202 ‰ (p = 0.003; Fig. 4a). The 15N–NH4+ label took between 0.25 and 12 h to become incorporated into NO at both sites; *[δ-15N]NO reached 949 ± 152 ‰ in the desert (F3,9 = 12.1, p = 0.002; Fig. 4c) and 754 ± 132 ‰ in the chaparral (F3,9 = 60.4, p < 0.001; Fig. 4d).

Fig. 4
figure 4

Isotopic signature (*[δ-15N]NO) of NO emitted from the desert (a, c) and chaparral (b, d) over 24 h after wetting dry soils with nitrate (NO3a, b) or ammonium (NH4+c, d) solutions. Lines represent the mean [*δ-15N]NO (n = 4) from each treatment within each site. Dots represent individual measurements using passive Ogawa samplers. Asterisks indicate if the mean for a given time differed from the control (*p < 0.1, **p < 0.05, ***p < 0.01)

The natural abundance δ15N– and δ18O–NO values emitted from soils amended with only deionized water decreased over the course of the experimental incubation (Fig. 5). The *[δ18O]NO decreased from approximately 10 ‰ prior to wetting to − 15 ‰ 24 h after wetting in both the chaparral (F3,21 = 7.35, p = 0.002) and the desert (F3,21 = 11.5, p = < 0.001). Similarly, *[δ15N]NO decreased from approximately − 10 ‰ to − 40 ‰ over the course of the incubation in both the chaparral (F3,21 = 5.29, p = 0.007) and the desert (F3,21 = 5.15, p = 0.01).

Fig. 5
figure 5

Dual isotope plot of NO (*[δ-15N]NO and *[δ-18O]NO) produced after wetting dry soils from the desert (a) and chaparral (b) with 500 mL water. Colors correspond to timing of the isotopic signature of NO collected from ambient air and from soils after wetting. Each dot represents the isotopic composition of NO measured at each of the shrubs (n = 8). Isotopic NO composition is presented from shrubs receiving water-only additions

Soil NO emissions increased in proportion to incremental NH4+ additions in the chaparral (R2 = 0.58, p = 0.03, Fig. 6d), whereas in the desert, the relationship was positive but weak (R2 = 0.45, p = 0.07, Fig. 6c). Adding NO3 may have increased NO emissions in the chaparral, but the relationship was weak (p = 0.09, Table 2); adding NO3 did not increase NO emissions in the desert (p = 0.28, Table 2).

Fig. 6
figure 6

Cumulative soil NO emissions (µg N–NO m−2) from the desert (a, c) and chaparral (b, d) over 24 h after wetting dry soils with nitrate (NO3a, b) or ammonium (NH4+c, d) solutions. Lines show the linear regression between added N and cumulative NO emissions. Shaded gray areas represent the 95% confidence interval for statistically significant linear regressions (p < 0.1)

Soil NH3 emissions increased immediately after wetting both sites but remained higher in the desert relative to the chaparral (Fig S1). In the desert, NH3 emissions averaged 27.3 ± 24.6 µg N–NH3 m−2 h−1 between 0.25 and 12 h in NO3 amended soils; the NO3 treatment did not increase NH3 emissions compared to soils amended with only water (Fig S1a). In NH4+ amended desert soils, NH3 emissions averaged 52.5 ± 45.0 µg N–NH3 m−2 h−1 between 0.25 and 12 h, compared to 16.7 ± 10.6 µg N m−2 h−1 in soils amended with only water (Fig S1c).


We investigated the dynamics of and mechanisms driving pulsed NO and N2O emissions during drying–wetting cycles in two contrasting drylands. We found that soil NO emissions increased in proportion to the amount of NH4+ added in both sites, although this relationship was weaker in the desert, partially supporting the hypothesis that increasing biological process rates would increase N emissions and suggesting that nitrification may control NO emission magnitude in these coarse-textured soils. In contrast, increasing N supply did not increase N2O emissions at either site, which does not support the hypothesis that N2O emissions are limited by NO3 or NH4+. While N addition did not stimulate N2O emissions, N2O was produced in part by the near-instantaneous reduction of NO3, raising questions as to the mechanisms driving NO3 reduction in these dryland soils and how factors controlling these emissions could help explain variation in N emissions across ecosystems.

N2O emissions: controls and dynamics

While we expected N2O to increase within minutes after wetting to produce large emission pulses (Eberwein et al. 2020), the incorporation of 15N–NO3 tracer into N2O within 15 min of adding water was unexpected (Fig. 2a,b)—denitrification is an anaerobic process not thought to dominate in well-aerated coarse-textured soils during dry summer months (Werner et al. 2014). Possibly, rapid onset of microbial respiration (Birch 1958; Jenerette and Chatterjee 2012) consumed sufficient O2 to stimulate N2O production via denitrification immediately after adding water, or soil aggregates may have sustained a viable denitrifier population within anoxic microsites throughout the hot and dry summer (Sexstone et al. 1985). Indeed, laboratory studies show denitrification enzyme activity can be maintained in dry soils (Peterjohn 1991; Parker and Schimel 2011), perhaps suggesting this process is viable in deserts. However, because NO is produced as an obligate intermediate during denitrification, and our 15NO3 tracer was not incorporated into NO within 15 min post-wetting in the desert (Fig. 4a), denitrification may not have contributed to rapid N2O emissions. Besides biological processes, chemodenitrification can produce N2O (Zhu-Barker et al. 2015; Heil et al. 2016; Harris et al. 2021), but the abiotic reduction of NO3 has only been reported under manipulated laboratory settings (Davidson et al. 2003; Matus et al. 2019) and is yet to be demonstrated to occur in-situ (Colman et al. 2007, 2008). The detection of the 15N–NO3 label in N2O within 15 min of wetting dry soils at our site shows that dryland soils have the capacity to reduce NO3 immediately after wetting and argues for additional work identifying which processes contribute to rapid N2O emissions.

Even though 15N–NO3 was rapidly reduced to N2O, adding more NO3 did not increase the magnitude of pulsed N2O emissions. This suggests that the processes reducing NO3 to N2O are not limited by soil N availability (i.e., the size of the pipe), and that other factors regulate the magnitude of N2O emissions. For example, more 15N–NO3 tracer was reduced to N2O in the desert (Fig. 2; Table S1), where soils had higher pH and warmer temperature compared to the chaparral (Table 1). These soil properties and environmental conditions can determine which N intermediates are released to the atmosphere, potentially explaining variation in the magnitude of N2O emissions between sites. For example, higher pH desert soils may have increased denitrification rates (Knowles 1982), or warmer temperatures in the desert may have favored abiotic reactions that can produce N2O (McCalley and Sparks 2009; Zhu-Barker et al. 2015). Average peak N2O emissions from the desert were slightly higher compared to emissions measured in tropical forests (66.4 ng N–N2O m−2 s−1; Hall and Matson 2003) and temperate agricultural systems (355 ng N–N2O m−2 s−1; Smith et al. 1994), which are thought of as denitrification hotspots. In addition to differences in pH and temperature between sites, variation in soil properties underneath each shrub could override any effect of experimental N addition on N2O emissions. Indeed, N2O emissions are notoriously difficult to predict since they are often driven by high rates of microbial activity within microsites where soil C and N are concentrated (Sey et al. 2008; Harris et al. 2021). As such, greater replication may be needed to detect effects of N addition over the inherent variability in N2O emissions. Despite this variation, documenting the rapid reduction of NO3 to form N2O is an important step in identifying controls over dryland N2O emissions.

NO emissions: controls and dynamics

Nitrification produced NO at our sites as supported by the detection of 15N–NH4+ in NO (Fig. 4c,d) and the positive response of NO emissions to adding NH4+ (Fig. 6c,d). In addition to nitrification, denitrification also produced NO at both sites; 15N–NO3 was reduced to NO 12 h after wetting dry soils in the desert, and within 15 min in the chaparral (Fig. 4a,b). Denitrifiers can initiate NO3 reduction within hours of decreasing soil O2 concentrations (Liu et al. 2019) and maintain this activity once aerobic conditions return (Roco et al. 2016). We also observed a simultaneous decrease in *[δ18O]NO and *[δ15N]NO over the course of the incubation at both sites, perhaps suggesting other processes produced NO (Fig. 5). While changes to *[δ18O]NO and *[δ15N]NO could have been caused by interactions between NO and VOCs (Walters and Michalski 2016), these observations may also indicate nitrifier denitrification activity as observed in a Mediterranean grassland (Homyak et al. 2016). Nitrifier denitrification produces NO from NO2, which contains O from both water and air, whereas nitrification produces NO from NH2OH, which contains only O from air (Andersson and Hooper 1983; Buchwald et al. 2012; Medinets et al. 2015; Boshers et al. 2019). As such, the change in *[δ18O]NO may reflect incorporation of 18O from the NO2 produced prior to and after wetting these dry soils (Homyak et al. 2016). Furthermore, biological NO production pathways—including nitrifier denitrification and nitrification—fractionate against 15N by 28–60 ‰ (Robinson 2001), consistent with the simultaneous decrease in *[δ15N]NO observed throughout the incubation. Abiotic reactions may have also contributed to soil NO efflux by converting nitrification intermediates—such as NO2 or NH2OH—to NO (McCalley and Sparks 2009; Heil et al. 2016; Homyak et al. 2017). Regardless of the mechanism, our work suggests that multiple pathways, including those requiring anaerobic conditions, produce NO after wetting these dry coarse-textured soils.

Soil NO-producing pathways were likely limited by soil N availability, since adding more N was associated with higher NO emissions. The positive response of cumulative NO emissions to adding NH4+ is consistent with N limitation of N trace gas production via nitrification (Davidson et al. 2000; Vourlitis et al. 2015; Prosser et al. 2019), as has been observed in other drylands (Hartley and Schlesinger 2000; Eberwein et al. 2020). However, other factors besides N limitation likely contributed to the magnitude of the NO pulse since NO emissions diverged between sites; the tracers were reduced to NO more quickly in the chaparral (Fig. 4), while cumulative NO emissions had a larger positive relationship with NH4+ addition in the desert (Fig. 6). These differences between sites may be explained by background microbial activity. For example, chaparral soils were exposed to fog (Table 1) and were already producing NO before we added water, whereas desert soils were not (Fig. 3b,d). In this sense, non-rainfall water inputs via fog (McHugh et al. 2015) may have influenced the magnitude of pulsed N emissions by resuscitating microbes and priming them for the N we added, helping to explain the rapid NO emission pulse (Fig. 3b,d) and the rapid incorporation of 15N–NO3 into NO (Fig. 4b). In contrast to chaparral, microorganisms in the relatively drier desert took hours to activate before producing the more delayed, but relatively long-lasting, NO emission pulse (Fig. 3a,c). In the desert, we measured higher NH3 emissions relative to the chaparral (Fig S1), consistent with higher soil pH favoring NH3 production from the equilibrium between NH3 and NH4+ (pKa = 9.25; Avnimelech and Laher 1977). This suggests the longer NO emission pulse in the desert could have been sustained by greater NH3 diffusion through soil pore space and supply to nitrifiers even as drying soils may have limited nitrifier access to NH4+ in soil pore water (Stark and Firestone 1995). The role of NH3 diffusion to nitrifiers may also help explain why the relationship between NH4+ addition and NO emissions was weaker in the desert; variable background NH4+ concentrations may have supplied NH3 to nitrifiers even when little N was added to soils. Taken together, our observations support the hypothesis that wetting-induced NO emissions are limited by soil N availability but suggest that environmental and edaphic factors contribute to variation in NO production among ecosystems.


We demonstrate that rapid NO3 reduction (within 15 min) can occur even in coarse summer-dry desert soils under temperature extremes to produce N2O. However, the N2O emissions produced were insensitive to experimentally adding N. Identifying the processes that govern the rapid NO3 reduction pathway will help constrain variation in N emissions across dryland soils as these ecosystems expand with expected changes in climate (Huang et al. 2016). In contrast to N2O, NO emissions were governed by N limitation of multiple N cycling processes, suggesting that N-limited NO production pathways may increase in response to higher rates of atmospheric N deposition (Fenn et al. 2006). These wetting induced N trace gas production pathways appear widespread across ecosystems that experience repeated drying–wetting cycles and will likely become increasingly important sources of atmospheric NO and N2O as global precipitation regimes become more variable.