Site description.

The two-factor experiment (N source and time) was conducted from 9 to 11 Aug 2022 at the US Department of Agriculture–Agriculture Research Service, Application Technology Research Unit in Wooster, OH, USA (lat. 40°46′26.0″N, long. 81°54′42.6″W) in an open-wall, polyethylene-covered hoop house (i.e., high tunnel) to exclude rain (Supplemental Figs. 1 and 2). The N sources were arranged in a randomized block design.

Plant material and potting.

At Last^® Rosa × ‘HORCOGJIL’ (USPP 27,451; Rosaceae family) liners (224; Griffin Greenhouse Supplies, Inc., Tewksbury, MA, USA) were received and held in an open-wall, polyethylene-covered hoop house that received daily irrigation until planting. On 23 May 2022, shrub rose liners were pruned to similar sizes before planting to ensure experimental unit uniformity; then, an individual plant was transplanted in a #2 trade container (5.68 L volume × 21.6 cm height × 22.9 cm width; C600; Nursery Supplies Inc., Chambersburg, PA, USA) with a pine bark substrate (T.H. Blue Inc., Eagle Springs, NC, USA). The pine bark substrate was amended with 1.8 kg·m⁻³ ground dolomitic lime [95.0% CaCO₃ equivalent, 21.6% calcium (Ca), 10.0% magnesium (Mg); Soil Doctor, Atlanta, GA, USA] and 0.45 kg·m⁻³ granular micronutrient fertilizer [6.0% Ca, 3.0% Mg, 12.0% sulfur (S), 0.1% boron (B), 1.0% copper (Cu), 17.0% iron (Fe), 2.5% manganese (Mn), 0.1% molybdenum (Mo), 1.0% zinc (Zn); Micromax; Everris, Dublin, OH, USA). Pine bark was mixed with amendments in a ribbon mixer for 10 min [Twister I (single phase); Bouldin and Lawson, McMinnville, TN, USA] to uniformly incorporate amendments into the substrate.

Plant establishment.

Shrub roses were grown for 7 weeks before experimental initiation (Fig. 1) to allow for establishment of the microbiome comparable to that of common production conditions and practices within the specialty crop industry. During this establishment period, plants received daily predawn (6:00 AM) irrigation delivered by pressure-compensated plum spray stakes (SKU 22500-002030; Netafim, Orbia Inc., Hatzerim, Israel). Approximately every 2 to 3 d, fertigation was applied to plants at 150 mg-N·L⁻¹ with a complete water-soluble fertilizer (20N–5P₂O₅–20K₂O; 8% NH₄-N, 8% NO₃-N, 4% Urea-N; SKU 200285; Turf 2; Harrells LLC, Lakeland, FL, USA) as needed using a fertilizer injector (Dosatron D14MZ2; Ingersoll Rand, Davidson, NC, USA) from a stock solution at a 1:100 dilution, which is a typical practice of nursery production in the eastern United States. Two weeks before initiation of the experiment, plants were fertigated with 100 mg·L⁻¹ potassium nitrate (KNO₃) using a 1:100 diluted stock solution using a dosing pump. The week before initiation of the experiment, the plants were irrigated with water without fertilizer to flush any freely available residual DIN from the experimental units. No nutritional deficiencies of model crop were noted during establishment and experimentation.

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Nitrogen source and application.

The three N source treatments were CH₄N₂O-N (SKU U20225; Research Products International, Mount, Prospect, IL, USA), NH₄⁺-N applied as ammonium phosphate monobasic [(NH₄)H₂PO₄] (SKU A684-3; Thermo Fisher Scientific Inc., Waltham, MA, USA), and NO₃-N applied as KNO₃ (SKU P263-500; Thermo Fisher Scientific Inc.). Additionally, 2 L of 200 mg·L⁻¹ of each N-containing solution was applied to each container; this dose was shown by Cavins et al. (2005) as sufficient for container crop production. Each N source was formulated by combining water with the N source agricultural salt to formulate treatments on 9 Aug 2022 as a single fertilization event; no further N application occurred for the remainder of the study (Fig. 1).

Substrate physical properties.

Pine bark substrate physical properties (n = 4) were determined using the North Carolina State University porometer method (Fonteno and Harden 2010). Stabilized pine bark had a minimum air space of 29.7% by volume [standard error (SEM), ±1.2], maximum water holding capacity of 49.3% by volume (SEM, ±0.6), total porosity of 79.0% by volume (SEM, ±0.9), and a bulk density of 0.18 g·m⁻³ (SEM, ±0.1). Substrate texture (n = 3) was defined by the following three separate particle classes (Altland et al. 2014): coarse particles (diameter > 2.0 mm); medium particles (0.5 mm < diameter ≤ 2.0 mm); and fine particles (diameter < 0.5 mm). Particle size distribution was determined by mechanical agitation of oven-dried substrate for 5 min on a Ro-Tap shaker (Rx-29; W.S. Tyler, Mentor, OH, USA). Particle size distribution was determined by using sieves with appropriate mesh openings. Particle classes (by weight; SEM, ±0.1) were coarse (62.7%), medium (26.3%), and fine (11.0%).

Temperature.

Daily air temperature data were obtained using a HOBO External Temperature/RH Sensor Data Logger (SKU MX2302A; Onset, Bourne MA). The average air temperature during this trial was 23.9 °C, with a maximum temperature of 36.5 °C and minimum temperature of 16.2 °C. Substrate temperature data were collected using HOBO Pendant MX Water Temperature Data Loggers (SKU MX2201; Onset) placed in the center of the containerized substrate. The average substate temperature during this trial was 23.7 °C, with a maximum temperature of 34.5 °C and minimum temperature of 17.8 °C (Supplemental Fig. 3).

Pore water extraction and N analysis.

Pore water extractions (Fig. 1) occurred at 4, 8, 24, and 48 h after initiation (HAI) using the pour-through method (Wright 1986). In brief, we applied 180 mL of deionized water evenly over the substrate surface of each experimental unit (container; n = 4; total = 16). This process resulted in 40 to 60 mL of displaced pore water (substrate bulk solution). The displaced pore water was collected for analysis. Pore water was transferred into a labeled 50-mL conical tube (Falcon^® 50-mL conical tube; Corning, Corning, NY, USA). Samples were separated and analyzed to determine electrochemical properties [pH and electrical conductivity (EC)]; mineral nutrient ions; and total organic N and carbon.

Additional deionized water was added to each experimental unit or container at 23 and 47 HAI to ensure the container was saturated before conducting a pour-through, which requires containers to be at field capacity to obtain a representative sample of plant-available pore water (Wright 1986). Specifically, an additional 200 mL or 300 mL of deionized water was added to each container at 23 HAI and 47 HAI to bring containers to field capacity. At 15 min after application, leachate was collected and reapplied over the top of the substrate surface. Substrates were then allowed to freely leach for 45 min before pour through extraction.

Pore water sample pH was measured using a benchtop meter (MA235 pH/Ion Analyzer; Mettler Toledo, Columbus, OH, USA) with an InLab Expert PRO-ISM pH electrode (Mettler Toledo). The EC was measured using a benchtop conductivity meter (S230 SevenCompact; Mettler Toledo) with an InLab 741-ISM electrode (Mettler Toledo).

Extracted pore water aliquots were filtered with a 0.45-µm syringe filter (Choice™ nylon Syringe Filter; Thermo Fisher Scientific Inc.). A 7-mL aliquot of filtered solution was placed in a 10-mL polystyrene vial (074228; Thermo Fisher Scientific Inc.) for immediate ion chromatography analysis, 10 mL was placed in a 22.2-mL PTFE/SILicone ulined (824030-2385; Neta Scientific, Hainesport, NJ, USA) for an immediate total organic carbon (TOC)/dissolved organic carbon (DOC) total N (TN) analysis, and a 10-mL aliquot was placed in a 15-mL polypropylene vial (339650; Thermo Fisher Scientific Inc.). An additional aliquot was placed in a 10-mL round bottom borosilicate glass tube (14-961-27; Thermo Fisher Scientific Inc.) for urea analysis. The rest of the sample was frozen and stored at −20 °C for reanalysis if necessary.

Ion concentrations were determined via ion chromatography coupled with an AS-AP chilled autosampler (ICS6000; Thermo Fisher Scientific Inc.). An Ion Pac AS19 2-× 250-mm column (062886; Thermo Fisher Scientific Inc.) and Ion Pac CS12a 2-× 250-mm column (046075; Thermo Fisher Scientific Inc.) were used for anion analysis and cation analysis, respectively. Limits of detection of anions nitrite (NO₂⁻-N) and nitrate (NO₃⁻-N) and cation ammonium (NH₄⁺-N) were 0.2 to 125 mg·L⁻¹. The TOC and TN (sum of organic and inorganic N forms) analyses via combustion were performed using a TOC and TN aqueous analyzer with ASI-L Autosampler (Shimadzu, Colombia, MD, USA) with a range of 0 to 100 mg·L⁻¹.

The urea-N analysis was performed using Method 10-206-00-1-A in deionized water using the Flow Injection Analysis System (Lachat Quikchem 8500; Hach Co., Loveland, CO, USA). Samples were diluted to suit detection limits of 0.1 to 20 mg·L⁻¹ CH₄N₂O-N and analyzed in The Ohio State University Service Testing and Research laboratory (STAR Laboratory, Wooster, OH, USA). Diluted samples were reanalyzed immediately if not within detection limits for any measure.

Reactive N gas analysis.

Gaseous emissions (Fig. 1) were sampled at 7, 31, and 55 HAI. A real-time gas analysis was conducted using Fourier transform infrared spectroscopy (FTIR; Terra GT5000; Gasmet, Vantaa, Finland). A closed-loop sampling system was made using a plastic 22.74-L bucket with polyurethane tubing (McMaster-Carr, Cleveland, OH, USA) that was connected to the FTIR apparatus. Chamber integrity was confirmed with CO₂ measurements during experimental sampling. One experimental unit (container with plant) was placed inside the chamber of the closed-loop system, and the gas flux was measured over 5 min (n = 4 per treatment). Then, the FTIR apparatus was returned to ambient gas concentrations for 3 min; ambient gas concentrations were sampled between each experimental unit. Gas flux on a per-container basis over time, f_N·t⁻¹, was calculated using Eq. [1] (Gyawali et al. 2019).$$f_{\mathit{NOx}}=\frac{P_0 V_c}{R{T}_0} \frac{ \mathrm{\Delta }C}{\mathrm{\Delta }t}$$ [1] where $P_0$ is the pressure within the chamber [M·L⁻¹·t⁻²], which is assumed to be equivalent to the atmospheric pressure, V_c is the cumulative volume of the chamber, apparatus internal volume, and tubing volume [L³], R is the ideal gas law constant [M·L²·N⁻¹·T⁻¹·t⁻²], T₀ is the air temperature in kelvin (K) [T], ΔC is the change in concentration of a given gas on a molar basis [N·N⁻¹], Δt, which is then used to compare change in gaseous species concentration over change in time [t]. Gaseous species of interest included N₂O-N, NO-N, NH₃-N, NO₂-N, and the sum of all four N gases as RN. Observations of gaseous flux were stopped before head space saturation of each specific gas species to ensure flux calculations were not underestimating or overestimating gas flux. The FTIR apparatus was calibrated before the initiation of gas data collection for each sampling date.

Plant tissue and substrate N content.

Initial and final harvest of aboveground and belowground biomass and substrate were performed before experiment initiation (0 HAI) and at 48 HAI to determine the N content of four plants per block, resulting in a total of 16 plants (Fig. 1). Roots were cut from shoots at the substrate surface and washed using a high-pressure water stream to remove substrate before drying. Shoot and root tissues were dried (60 °C) until no change in mass could be detected. Bark substrates were dried (100 °C) and weighed until no loss of mass could be detected. Plant tissues and substrates were weighed and then ground to pass through a 2-mm sieve using a laboratory mill (Foss Cyclotec^TM 1093; Foss, Hillerød, Denmark). Once ground, plant tissue and substrate were placed in a labeled coin envelope and mailed to Brookside Laboratories (New Bremen, OH, USA) to determine the N content by combustion (T002 analysis). Plant tissue and substrate N contents were determined by dry weight $\times$ N concentration, and root and shoot N contents were combined to determine total plant N.

Statistical analysis and interpretation.

Data were analyzed using JMP Pro 15.0 (SAS Institute, Inc., Cary, NC, USA). Distribution of all data were examined, and most were determined to have a normal distribution. If data were not normally distributed, then residuals, log, and cube root transformations were conducted to meet the assumptions of normal distribution. The N source and time were subject to a two-way analysis of variance. Pearson correlation coefficients (R² > 0.4) were used to determine relationships. P ≤ 0.05 was used to determine statistical differences. Gaseous data were subject to a multivariate analysis of variance for repeated measures.

Site description.

A glass greenhouse was used to evaluate the effects of a controlled environment (Supplemental Figs. 2 and 4). Unless otherwise stated, plant material and potting, plant establishment, N source, measures of substrate physical properties and temperature, pore water extraction, gas analysis, and data analysis and interpretation were the same as those used in the high tunnel experiment (Supplemental Fig. 4). The mean greenhouse daytime temperature was 24 °C and the mean greenhouse nighttime temperature was 18 °C while maintaining a 16-h photoperiod provided by sunlight with supplemental lighting delivered by high-pressure sodium and metal halide lamps (GLX/GLS e-systems GROW lights; PARsource, Petaluma, CA, USA) (Supplemental Fig. 5).

Roses were allowed to establish for 9 weeks in the high tunnel after potting. On 26 Aug 2022, plants were moved to a heated glass greenhouse to allow acclimation of experimental units for 3 weeks before experiment initiation. Plants were allowed to establish for 12, 13, or 14 weeks. The experiment timeline was as follows: NH₄-N was applied on 13 Sep 2022; data collection concluded on 15 Sep 2022; NO₃-N was applied on 20 Sep 2022; data collection concluded on 22 Sep 2022; CH₄N₂O-N was applied on the 27 Sep 2022; and data collection concluded on 29 Sep 2022. The staggered separation of each N source application reduced the amount of time between sample collection and analysis.

Fertigation was stopped before each N source application. Deionized water was the irrigation water source for 4 d before each N source application to remove pore water DIN. We were unable to analyze samples for TN because of analytical equipment failure; therefore, we excluded these data from analysis.

Aqueous pore water.

Urea-N curvilinearly decreased (P < 0.0001; R² = 0.74) from 44.7 mg·L⁻¹ at 4 HAI to 31.2 mg·L⁻¹ at 8 HAI, and to 1.4 at 24 HAI (Fig. 2). By 48 HAI, CH₄N₂O-N was undetected in the extracted pore water. Ammonium-N exhibited a polynomial fit whereby concentrations averaged 3.0 mg·L⁻¹ at both 4 HAI and 8 HAI, increased to 10.9 mg·L⁻¹ at 24 HAI, and decreased to 6.3 at 48 HAI (P = 0.0004; R² = 0.70) (Fig. 2). Nitrite-N concentrations curvilinearly increased (P < 0.0001; R² = 0.81) throughout sampling from 0.4 mg·L⁻¹ at 4 HAI to 7.4 mg·L⁻¹ at 24 HAI and peaked (14.4 mg·L⁻¹) at 48 HAI. Nitrate-N exhibited a polynomial trend (P = 0.0001; R² = 0.75) whereby concentrations were initially low, averaging 1.8 mg·L⁻¹ for the first 24 HAI; thereafter, they increased to 11.1 mg·L⁻¹ at 48 HAI (Fig. 2). Total N concentrations decreased over the 48-h experimental duration and exhibited a polynomial trend, with a high initial value of 54.2 mg·L⁻¹ that decreased to 23.4 mg·L⁻¹ at 24 HAI (P < 0.0001; R² = 0.81) (Fig. 2). Total N remained stable for the final 24 h of the experiment (Fig. 2). Initial (4 HAI) pore water extract pH was 6.7, and it increased to 7.0 and 7.2 at 24 HAI and 48 HAI, respectively (P = 0.0013) (Fig. 3). The EC remained stable at approximately 0.8 mS·cm⁻¹ over the 48-h experimental period (Fig. 3). The TOC concentration remained relatively stable throughout the experiment, averaging 54 mg·L⁻¹ (Supplemental Fig. 7).

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Gaseous emissions.

There were no measured emissions of NH₃-N or NO₂-N gas after the application of CH₄N₂O-N (Fig. 2). Nitrous oxide-N emission exhibited a polynomial trend whereby concentrations increased from 0.2 at 7 HAI to 4.1 at 55 HAI (P = 0.0007; R² = 0.80) (Fig. 2). Nitric oxide-N emission increased from 0.6 µg·min⁻¹ at 7 HAI to 7.4 µg·min⁻¹ at 55 HAI (P = 0.024). Nitric oxide and nitrous oxide (NO-N and N₂O-N) were the two major contributors to RN emissions regardless of sampling time.

Aqueous pore water.

The aqueous NH₄⁺-N concentration was highest at 14 mg·L⁻¹ at the 4-HAI interval and decreased to 4.5 mg·L⁻¹ at 48 HAI (P = 0.0016) (Fig. 2). Aqueous NO₂⁻-N increased linearly throughout the experiment (P < 0.0001; R² = 0.78) (Fig. 2). Nitrate-N was stable for the first 8 HAI and increased to 7.8 mg·L⁻¹ at 48 HAI, exhibiting a linear trend (P = 0.0022; R² = 0.50) (Fig. 2). Total N concentrations remained stable throughout the experiment, with an average TN concentration of 19.5 mg·L⁻¹. Urea-N was not present in any samples. Pore water pH remained stable at 6.4, whereas EC had a decreasing polynomial fit (P = 0.0328; R² = 0.41) (Fig. 2) whereby values decreased from 1.1 to 0.9 mS·cm⁻¹ over the experiment. The TOC increased linearly by 71% (P = 0.0001; R² = 0.66) over the 48-h sampling period (Supplemental Fig. 7).

Gaseous emissions.

Two major contributors to RN emissions regardless of sampling time were NO-N and N₂O-N. Emissions of N₂O-N increased linearly from 0.2 µg·min⁻¹ at 7 HAI to 1.9 µg·min⁻¹ at 55 HAI (P = 0.0012; R² = 0.67) (Fig. 2). There were no differences in NO-N emissions during the 55-h span (P = 0.8699). A low emission of NH₃-N 0.1 µg·min⁻¹ was detected at 7 HAI; thereafter, no NH₃-N emissions occurred for the remainder of experiment. Nitrogen dioxide-N was not detected at any sampling point during the experiment.

Aqueous pore water.

Aqueous NO₃⁻-N exhibited a decreasing polynomial fit (P = 0.014; R² = 0.48) (Fig. 2) whereby concentrations decreased from 93.8 mg·L⁻¹ at 4 HAI to 58.3 mg·L⁻¹ at 24 HAI. Nitrate-N concentrations remained stable for the final 24 h of sampling (Fig. 2). Nitrite-N remained low hroughout all sampling, averaging less than 0.2 mg·L⁻¹ pooled across all sample collections. Ammonium-N and CH₄N₂O-N remained below detection limits for all samples. Total N decreased from 90.1 mg·L⁻¹ at 4 HAI to 68.6 mg·L⁻¹ at 24 HAI and remained stable for the remainder of the trial (P = 0.0018). The pH of the pore water solution increased from 6.5 to 7.1 over the experiment (P < 0.0001) (Fig. 3). The EC values were higher than those of other N source treatments and decreased from 1.4 mS·cm⁻¹ 4 HAI to 1.1 mS·cm⁻¹ at 48 HAI (P = 0.0002), thus exhibiting a polynomial fit (P < 0.0001; R² = 0.77) (Fig. 3). The TOC concentrations (mg·L⁻¹) showed little change during the first 8 HAI; it decreased from 29 mg·L⁻¹ to 28 mg·L⁻¹ but increased during the remainder of the sampling points, with a final concentration of 42 mg·L⁻¹ at 48 HAI (P = 0.0008) (Supplemental Fig. 7).

Gaseous emissions.

Nitric oxide and N₂O (NO-N and N₂O-N) were the two major contributors to RN emissions regardless of sampling time. Gaseous N₂O-N and NO-N emissions remained stable at all sampling times throughout the experiment (Fig. 2). The application of NO₃⁻-N fertilizer resulted in the lowest RN emissions of all treatments across time. Emissions of NH₃-N and NO₂-N were not measured at any sampling point during experimental sampling.

Aqueous pore water.

Following the application of CH₄N₂O-N, the CH₄N₂O-N concentration was initially 0.1 mg·L⁻¹ at 4 HAI and peaked at 30.4 mg·L⁻¹ at 8 HAI; then, it decreased to 17.0 mg·L⁻¹ at 24 HAI and 0.4 mg·L⁻¹ at 48 HAI, thus exhibiting a polynomial fit (P = 0.0341; R² = 0.41) (Fig. 4). Ammonium-N concentration exhibited a polynomial trend with an initial concentration of 0.9 mg·L⁻¹ at 4 HAI, increasing to 33.5 mg·L⁻¹ at 8 HAI and 38.0 mg·L⁻¹ at 24 HAI; thereafter, NH₄⁺-N concentrations decreased to 19.1 mg·L⁻¹ at 48 HAI (P = 0.0045; R² = 0.56). Nitrite-N concentrations steadily increased throughout sampling with a polynomial fit (P < 0.0001; R² = 0.8319). Initial NO₂⁻-N concentrations were 0 mg·L⁻¹ at 4 HAI, increasing to 0.4 mg·L⁻¹ at 8 HAI and continuing to increase to 4.4 mg·L⁻¹ at 48 HAI, exhibiting a polynomial fit (P < 0.001; R² = 0.83) (Fig. 4). Nitrate-N concentrations were low during the first 8 h; they increased from 0.3 mg·L⁻¹ at 4 HAI to 14.7 mg·L⁻¹ at 48 HAI and were characterized by a polynomial fit (P < 0.0001; R² = 0.9496). The sum of all inorganic N concentrations was initially low, with a concentration of 1.2 mg·L⁻¹ at 4 HAI, which increased to 64.8 mg·L⁻¹ at 8 HAI and then decreased to 57.4 mg·L⁻¹ at 24 HAI and 38.4 mg·L⁻¹ at 48 HAI (Fig. 4). The CH₄N₂O-N treatment EC values did not change over time with this treatment (Fig. 5). The CH₄N₂O-N treatment pore water TOC remained relatively stable throughout the experiment, with an average of 89.2 mg·L⁻¹ (Supplemental Fig. 8).

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Gaseous emissions.

Nitrous oxide-N emission concentrations were initially high at 7 HAI, with a concentration of 2.1 µg·min⁻¹, peaked at 31 HAI at 2.6 µg·min⁻¹, and then decreased to 0.3 µg·min⁻¹ at 55 HAI, thus expressing a polynomial fit (P = 0.0189; R² = 0.5862) (Fig. 4). Nitric oxide-N emissions were 1.4 µg·min⁻¹ at 7 HAI and peaked 31 HAI at 4.9 µg·min⁻¹; however, the values were not statistically significant (P = 0.573). There were no emissions of NO-N at 55 HAI (Fig. 4). Nitric oxide and N₂O (NO-N and N₂O-N) were the two major contributors to RN emissions regardless of sampling time. Minor emissions of NH₃-N were measured at 55 HAI, with a value of 0.04 µg·min⁻¹, while there were no emissions of NO₂-N at any point during the experimental sampling for this treatment (Fig. 4).

Aqueous pore water.

Following the NH₄⁺-N application, NH₄⁺-N concentrations were initially 1.6 mg·L⁻¹ at 4 HAI, increased to and peaked at 24 HAI at 62.4 mg·L⁻¹, and then decreased to 56.1 mg·L⁻¹ at 48 HAI, thus exhibiting a polynomial fit (P < 0.0001; R² = 0.74) Fig. 4). Aqueous NO₂⁻-N exhibited a polynomial fit (P < 0.0001; R² = 0.92) (Fig. 4). The concentrations were low during the first 24 HAI (>0.2 mg·L⁻¹) and then increased to 1.6 mg·L⁻¹ at 48 HAI. The sum of all inorganic N concentrations was initially 3.8 mg·L⁻¹ at 4 HAI and increased to 68.3 mg·L⁻¹ at 48 HAI (Fig. 4). Urea-N was not present at any sampling point for the NH₄⁺-N treatment (Fig. 4). The NH₄⁺-N treatment pore water pH decreased by 0.5 units from 6.6 at 4 HAI to 6.1 at 48 HAI (P = 0.0153) (Fig. 5). The NH₄⁺-N treatment pore water EC (mS·cm⁻¹) remained relatively stable for the entirety of the experiment (Fig. 5). The TOC concentration remained stable throughout the experiment, increasing from 57.8 mg·L⁻¹ at 4 HAI to 65.0 mg·L⁻¹ at 48 HAI (P = 0.322) (Supplemental Fig. 8); however, it exhibited no significant difference.

Gaseous emissions.

The NH₄⁺-N treatment showed a similar trend of N₂O-N emission concentrations as the CH₄N₂O-N treatment, although to a lesser extent, increasing from 0.9 µg·min⁻¹ at 7 HAI to 1.0 µg·min⁻¹ at 31 HAI and decreasing to 0.3 µg·min⁻¹ at 55 HAI (P = 0.1372) (Fig. 4). Nitric oxide-N emission concentrations remained stable with only minor changes during the experiment (P = 0.4049) (Fig. 4). Nitric and nitrous oxide (NO-N and N₂O-N) were the two major contributors to RN emissions regardless of sampling time. There were no measured emissions of NH₃-N and NO₂-N at any sampling point during the experimental sampling (Fig. 4).

Aqueous pore water.

Following the application of NO₃⁻-N fertilizer, NO₃⁻-N concentrations were 47.4 mg·L⁻¹ 4 HAI, peaked at 8 HAI at 79.4 mg·L⁻¹, and decreased to 7.7 mg·L⁻¹ at 48 HAI (P = 0.026) (Fig. 4). Nitrite-N concentrations remained stable during the experiment (Fig. 4). Ammonium-N and CH₄N₂O-N remained below detection limits for all sampling points (Fig. 4). The sum of all inorganic N concentrations was initially 47.5 mg·L⁻¹ at 4 HAI, increased at 8 HAI with a concentration of 79.6 mg·L⁻¹, and decreased to 8.2 mg·L⁻¹ at 48 HAI (Fig. 4). The pH of the pore water when fertilized with the NO₃⁻-N treatment remained stable throughout the experimental sampling (P = 0.7533) (Fig. 5). The EC (mS·m⁻¹) of the pore water solution of the NO₃⁻-N treatment was stable throughout sample collection (P = 0.8785) (Fig. 5). The TOC concentrations (mg·L⁻¹) increased over the experiment from an initial value of 54 mg·L⁻¹ at 4 HAI to 82.1 mg·L⁻¹ at 48 HAI, exhibiting a polynomial fit (P = 0.0361; R² = 0.40) (Supplemental Fig. 8).

Gaseous emissions.

Nitrous oxide-N (N₂O) emission exhibited a polynomial fit (P = 0.0032; R² = 0.72); concentrations of 1.0 µg·min⁻¹ from the NO₃⁻-N treatment were the highest at 7 HAI and then decreased to 0.1 µg·min⁻¹ at 31 HAI and <0.1 µg·min⁻¹ at 55 HAI (Fig. 4). Low emissions of NH₃-N occurred at 31 HAI, with a concentration of 0.1 µg·min⁻¹. However, no emissions of this species occurred at 7 HAI or 55 HAI. There were no emissions of NO-N and NO₂-N during any sample collection.

Urea application.

Urea hydrolysis, conversion of CH₄N₂O-N to NH₄⁺-N, occurred in both the high tunnel and glass greenhouse regardless of environment. The highest emissions of RN gases occurred in the CH₄N₂O-N treatment; this indicated that most gaseous RN species formations appear to stem from CH₄N₂O-N hydrolysis and subsequent nitrification of NH₄⁺-N to NO₃⁻-N, while denitrification, anaerobic reduction of NO₃⁻-N to downstream gaseous species, seems to be a lesser contributor to RN emissions (Figs. 2 and 4). After applying CH₄N₂O-N, NH₄⁺-N increased and by-products of nitrification-mediated reactions that include increased NO₂⁻-N and NO₃⁻-N (Figs. 2 and 4) were observed. We inferred that aqueous NO₂⁻-N formation was brief and resulted in the rapid and complete nitrification of NO₂⁻-N to NO₃⁻-N because the NO₃⁻-N:NO₂⁻-N ratio was greater than one. Initial pH increased because CH₄N₂O hydrolysis during the first 24 HAI occurred as expected (Niemiera et al. 2014); thereafter, pH was stable for the remainder of the high tunnel experiment. We attributed this result to concurrent nitrification reactions (acidic) and denitrification reactions (alkaline) (Figs. 3 and 5). The changes in pH were less prevalent in the glass greenhouse (Fig. 5), likely because of a more stable pH as a result of a less variable environment that may have slowed the reaction rate. Our results indicated that applied CH₄N₂O-N is predominately and completely hydrolyzed within 48 h of application (Figs. 2 and 4). These results are similar to those found by Niemiera et al. (2014).

Ammonium application.

A steady decrease of NH₄⁺-N and a concomitant increase in aqueous NO₂⁻-N and NO₃⁻-N regardless of a high-tunnel or glass greenhouse setting indicated a rapid rate of nitrification as previously shown by Niemiera and Wright (1987a) (Fig. 2). We inferred that nitrification and denitrification occurred concurrently when NH₄⁺-N was applied because of the observed stable pH throughout sampling as previously described (Figs. 3 and 5). A less pronounced decrease in NH₄⁺-N and an increase in NO₃⁻-N were observed when CH₄N₂O-N was applied to containers in the glass greenhouse (Fig. 4). Similar to when NO₃⁻-N was applied, the decreased degrees of NH₄⁺-N and NO₃⁻-N changes were attributed to greater temperature control in the greenhouse lessening temperature extremes and decreased evapotranspiration and subsequent of solute flux. When applying NH₄⁺-N, products of nitrification NO₂⁻-N and NO₃⁻-N were found throughout pore water sampling in increasing concentrations over time (Figs. 2 and 4).

Nitrate application.

In both the high tunnel and glass greenhouse, when NO₃⁻-N was applied, there was a steady decrease of NO₃⁻-N without the presence of any other N intermediate aqueous species (Figs. 2 and 4). Denitrification resulted in lower emissions of RN from the NO₃⁻-N treatments (Figs. 2 and 4). This lack of N intermediaries implied that denitrification is the major N transformation resulting in the formations of N₂ gas (not measured). The decrease in NO₃⁻-N coupled with an increasing pH (Figs. 3 and 5) implicated the alkaline nature of denitrification (Havlin et al. 2014) that occurred after the application of NO₃⁻-N.

Synthesis.

Regardless of the N source applied, or whether the environment was uncontrolled or controlled, the trend of reactions included the formation of NH₄⁺-N via CH₄N₂O hydrolysis and then NO₃⁻-N via nitrification, which resulted in denitrification and the inferred complete denitrification to N₂. Thus, our findings suggested that these reactions proceed in a reaction sequence from CH₄N₂O hydrolysis to nitrification to denitrification, depending on the applied N form (Fig. 6). The formation of aqueous NO₂⁻-N appeared to be associated with nitrification from NH₄⁺-N application or hydrolyzed CH₄N₂O-N and was ephemerally formed via denitrification because of the low concentrations measured from the application of NO₃⁻-N (Figs. 2 and 4). Aqueous NO₂⁻-N formation in the denitrification reaction appeared to be readily transformed into gaseous N species shown by low NO₂⁻-N concentrations in pore water sampling (Figs. 2 and 4).

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Numerous microbial communities can use the applied N in the substrate to form predominately N₂ or lesser losses of RN gaseous intermediates (NO-N, N₂O-N, and NH₃-N) of nitrification and denitrification (Caranto and Lancaster 2017). Regarding applied N fertilizers, gaseous losses have been found to be nearly equivalent to aqueous losses via leaching according to previous research (Brown 2024). Urea resulted in higher RN gaseous emissions than those of the NO₃⁻-N treatment. In contrast, NH₄⁺-N treatment emissions were the same when compared with CH₄N₂O-N and NO₃⁻-N treatments (Figs. 2 and 4). Based on these results, we inferred that the gaseous losses from the NO₃⁻-N treatment were more likely in the form of N₂ than RN gaseous species N₂O-N, NO-N, and NH₃-N. Denitrification appears to be the final reaction of applied N because all processes after application in our system resulted in increasing concentrations of NO₃⁻-N.

Nitrogen inefficiencies in production.

We inferred from these data that using CH₄N₂O-N based fertilizers resulted in greater losses of RN gaseous emission compared with NO₃⁻-N-based fertilizer products. The application and use of NO₃⁻-N-based fertilizers may reduce greenhouse gases emissions from N fertilizers and more readily form the relatively inert N₂ species.

Neither plant N nor substrate-bound N was affected by the applied N source. We attributed the lack of plant N uptake and N substrate sorption to the introduction of N during establishment, the relatively short treatment exposure of 48 h, and analytical sampling sensitivity. These results support previous findings that denitrification resulting in N₂ gas formation is a major, if not the predominant, inefficiency when applying N fertilizers (Pitton et al. 2022). Although these losses may be economically concerning to growers, gaseous losses of N from containerized cropping systems may present less harmful RN gaseous emissions in container-grown crops in the United States than previously hypothesized by Pitton et al. (2022). However, RN losses or emissions on larger scales may result in considerable economic and environmental losses.

Future considerations.

Havlin et al. (2014) noted that the emission of N₂ via denitrification in commercially produced container-grown plants appeared to be a rapid process. Management practices that may have the capability to slow or stabilize applied forms of N or the release of N to the production system should be investigated. However, attempting to alter natural enzymatic or microbially aided processes, as present in N cycle transformations, may produce losses of other forms of N, as noted by Souza et al. (2023). Dinitrogen gas formation from applied fertilizers is a loss of fertilizer economic expenditures to growers; however, this N gaseous species is inert and the primary gas composing the atmosphere (approximately 78%). Fertilizer N not used by the plant or bound to substrate and subsequently lost as N₂ may be the best-case scenario because N₂ has no associated environmental or health-related consequences (Scheer et al. 2020).

As the mechanisms of nutrient release from N-containing controlled-release fertilizers continue to improve, researchers may be tasked with revisiting best management practices coupled with economic research to attempt to reduce inputs and labor costs that may relieve some of the financial and environmental pressures faced by the green industry (Fulcher et al. 2016). By improving fertilizer application, nutrient release, and longevity, N efficiency could be improved.

Products that claim to inhibit transformations of applied N forms and thereby mitigate aqueous and gaseous losses are currently being sold and marketed to the nursery industry; however, in a recent publication by Souza et al. (2023), some of these products have unintended consequences. When using some of these inhibition products, mitigated losses of N were offset by increased RN gas emissions, specifically N₂O-N, and leachate losses of NO₃⁻-N. By working with simulated conditions and investigating fertilizer and irrigation applications, as well as various other management practices, alternative cultural practices may be developed to improve N application and retention without the need for additional chemistries in the container throughout the production cycle.

Limitations within research.

A significant limitation within the scope of the N fate and gaseous emission of N is our inability to directly measure N₂ losses. Currently, analytical techniques are less effective for measuring N₂ emissions compared with many other gaseous species of interest; this is attributed to the use of N₂ in the sampling apparatus or high background atmospheric levels of N2 (Takaya et al. 2003). Until N₂ can be effectively and efficiently measured, skepticism about the applied N fate in a production system will remain a topic for debate.