Carbon Availability and Nitrogen Mineralization Control Denitriﬁcation Rates and Product Stoichiometry during Initial Maize Litter Decomposition

: Returning crop residues to agricultural ﬁelds can accelerate nutrient turnover and increase N 2 O and NO emissions. Increased microbial respiration may lead to formation of local hotspots with anoxic or microoxic conditions promoting denitriﬁcation. To investigate the effect of litter quality on CO 2 , NO, N 2 O, and N 2 emissions, we conducted a laboratory incubation study in a controlled atmosphere (He/O 2 , or pure He) with different maize litter types ( Zea mays L., young leaves and roots, straw). We applied the N 2 O isotopocule mapping approach to distinguish between N 2 O emitting processes and partitioned the CO 2 efﬂux into litter- and soil organic matter (SOM)-derived CO 2 based on the natural 13 C isotope abundances. Maize litter increased total and SOM derived CO 2 emissions leading to a positive priming effect. Although C turnover was high, NO and N 2 O ﬂuxes were low under oxic conditions as high O 2 diffusivity limited denitriﬁcation. In the ﬁrst week, nitriﬁcation contributed to NO emissions, which increased with increasing net N mineralization. Isotopocule mapping indicated that bacterial processes dominated N 2 O formation in litter-amended soil in the beginning of the incubation experiment with a subsequent shift towards fungal denitriﬁcation. With onset of anoxic incubation conditions after 47 days, N ﬂuxes strongly increased, and heterotrophic bacterial denitriﬁcation became the main source of N 2 O. The N 2 O/(N 2 O+N 2 ) ratio decreased with increasing litter C:N ratio and C org :NO 3 − ratio in soil, conﬁrming that the ratio of available C:N is a major control of denitriﬁcation product stoichiometry.


Introduction
Returning of crop residues is a common agricultural management strategy to prevent nutrient losses and to increase soil fertility. However, acceleration of N and C cycling processes often lead to increased losses of climate-relevant gases.
Addition of plant litter to soils has been proven to increase CO 2 and N 2 O emissions over a vast range of soil conditions and litter types [1][2][3][4]. Upon degradation, plant litter provides nutrients for decomposing and denitrifying microorganisms. Thus, variations in N 2 O emissions have often been related to litter quality, especially the C:N ratio [1,2]. For litter with C:N < 25:1, mineralization increases soil NO 3 − content leading to increased denitrification [5,6], while for C:N > 25:1, N is immobilized by soil microorganisms to decompose litter C compounds [7] and restricts N 2 O emissions [8]. When litter quality Appl. Sci. 2021, 11, 5309 2 of 21 was analyzed in more detail, easily degradable fractions explained a large share of the variability of N 2 O emissions [9,10], while the lignin content was not relevant [11,12]. Recent studies confirm that the quality of C compounds (especially water-soluble C) from litter is a main driver of denitrification after litter addition [13,14].
Easily degradable C compounds (e.g., sugars, proteins, amino acids, and carbohydrates) control litter decomposition dynamics in the initial phase and subsequent CO 2 efflux from soils [4,15,16]. Furthermore, the quality of organic substrates affects decomposition of soil organic matter (SOM) [17]. Readily accessible high-quality substrates increase SOM decomposition, leading to a positive priming effect in soils [18,19]. When litter and SOM turnover are increased after litter addition, microbial O 2 demand increases with increasing microbial respiration. This may lead to formation of local hotspots with anoxic or microoxic conditions providing favorable conditions for denitrifying soil microorganisms [20]. Accordingly, a recent study reported the highest denitrification-derived N 2 O losses in soils with high SOM priming after addition of labile C substrates (glucose, vanillin) [21]. However, further studies with plant residues are necessary to better understand the interactions between C turnover and denitrification.
The aim of this study was to investigate the effect of litter C quality and SOM turnover on denitrification. We anticipate that increased SOM turnover after litter addition promotes the formation of anoxic hotspots for denitrification and expect higher litter and SOM turnover from litter with high degradability. Thus, we hypothesize that (i) N 2 O fluxes from denitrification increase when C turnover from litter and SOM is high, leading to (ii) higher N 2 O+N 2 losses when litter with a high share of easily degradable C is added, while (iii) the N 2 O/(N 2 O+N 2 ) ratio is controlled by the availability of C org in relation to NO 3 − . Therefore, we setup a laboratory incubation experiment in an artificial N 2 -free atmosphere under fully controlled conditions. We compared different types of maize litter (fresh leaves and roots, straw) and investigated the effect of litter quality on total CO 2 , NO, N 2 O, and N 2 emissions. To trace maize litter (C4 plant) and SOM turnover, we used a grassland soil whose organic C originates solely from C3 vegetation and partitioned the CO 2 efflux into its sources (i.e., litter-and SOM-derived CO 2 ) based on the natural 13 C isotope abundances. In addition, we analyzed the intramolecular distribution of the naturally occurring 15 N and 18 O isotopes in the linear N 2 O molecule and applied the N 2 O isotopocule mapping approach to estimate the contribution of denitrification to N 2 O formation [22,23].

Preparation of Soil and Plant Material
The soil for the experiment was taken from a long-term field experiment at the grassland research station of the University of Gießen (latitude N50 • 32 , longitude E8 • 41.3 , elevation 172 m a.s.l.), sieved to 10 mm, air-dried, and stored at 4 • C. The soil was classified as Fluvic Gleysol of clay loam texture (32% clay, 41% silt, and 27% sand) with a pH (CaCl 2 ) of 5.67. Total soil N content was 0.42%, total soil C content was 4.2%, and δ 13 C was −28.37‰. Prior to the incubation experiment, the soil was pre-incubated in the dark for 5 weeks at 50% water holding capacity (WHC) and 20 • C.
Maize plants (Zea mays L. cv. Ronaldinio) were grown in nutrient solution [24] for 5 weeks. Leaves were cut from stems and left to wilt at room temperature for 4 h. Roots were rinsed with H 2 O dest and carefully dried with paper towels. Leaves and roots were stored at 4 • C and 90% relative humidity until experimental setup. Maize straw was collected from an experimental field site of the University of Göttingen after grain harvest in October 2018. Maize straw was shock frozen in liquid N 2 and stored at −20 • C. Prior to setting up the experiment, maize straw was unfrozen and all maize litter was cut to a size of 2 cm. A subsample of soil and maize litter was analyzed for total N, total C, and δ 13 C using an elemental analyzer (NA1110, CE-Instruments, Rodano, Milano, Italy) linked to a gas-isotope ratio mass spectrometer (Delta Plus, Finnigan MAT, Bremen, Germany) via a Conflo III Interface (Finnigan MAT, Bremen, Germany). Further, plant litter was analyzed for water-extractable C and N content. Briefly, 0.2 g of finely ground plant litter were extracted in 100 mL H 2 O bidest , shaken for 1 h, filtered with 0.45 µm polyether sulfone filters (Labsolute, Renningen, Germany), and stored at −20 • C. The extracts were analyzed for organic C and total N content using a multi N/C ® Analyzer (Analytik Jena, Jena, Germany). Another subsample of finely ground plant litter was analyzed by 13 C solidstate cross polarization magic angle spinning nuclear magnetic resonance spectroscopy ( 13 C-CPMAS NMR) using a Bruker Avance III 200 spectrometer (Bruker BioSpin GmbH, Karlsruhe, Germany). Samples were weighed into zircon oxide rotors and spun around a magic angle at a speed of 6.8 kHz. Contact time was 1 ms and the recycle delay time was set to 2 s, line broadening was set at 0. Peak integration areas were separated into −10-45 ppm (alkyl C), 45-110 ppm (O/N-alkyl C), 110-160 ppm (aryl C), and 160-220 ppm (carboxylic C).

Automatized Laboratory Incubation Experiment and Gas Analysis
The incubation experiment was carried out under fully controlled conditions using an automated soil incubation system with artificial atmosphere similar to systems described earlier [25][26][27][28]. For the incubation experiment, soil moisture was adjusted to 70% waterfilled pore space (WFPS, equivalent to 67.7% WHC or 31.9% gravimetric water content) and 50 mg N kg −1 was added by spraying a KNO 3 solution onto the soil and thoroughly stirring it with a spoon. For treatments with litter, litter was homogenously mixed with soil (Maize leaves: 40.5 g FM kg −1 , maize roots: 42 g FM kg −1 , maize straw: 12.8 g FM kg −1 ). The soil for each pot was prepared separately to ensure the same amount of litter was added. Then, the equivalent of 2.5 kg dry soil was filled into acrylic glass pots (inner diameter 172 mm, height 210 mm) with a porous ceramic plate at the bottom and compacted in a stepwise mode by filling a 2 cm-layer of soil in pots and compacting it with a plunger. To ensure continuity between soil layers, the surface of the compacted layer was gently scratched before adding the next soil layer. Soil height in the pots was 10 cm, and bulk density was 1.1 g cm −3 . Each litter treatment was replicated five times, a control treatment without litter was replicated four times, and one empty pot was included as reference to determine background gas concentrations.
Pots were tightly closed with transparent acrylic glass lids with rubber seals, and the outside of the pots was covered with dark plastic sheets to prevent algae growth. Pots were alternately evacuated using a rotary vacuum pump (Pfeiffer Vacuum GmbH, Asslar, Germany) and flushed with a gas mixture (80% He, 20% O 2 ) for 24 h. The gas mixture was prepared by using stainless steel capillaries of different length and inner diameter. For the first cycles, pots were evacuated from the top and the bottom and, subsequently, flushed with the He/O 2 gas mixture. Then, pots were evacuated from the bottom and simultaneously flushed from the top to replace the atmosphere inside the soil column. For measurements, the outlet of all pots was connected to flow-through multiposition valves (16 ports, Vici Valco Instruments, Houston, TX, USA) with multi-position actuator control modules (Vici Valco Instruments, Houston, TX, USA), and controlled by Trilution Software (Gilson Inc., Middleton, WI, USA) via an interface module (506C System Interface, Gilson Inc., Middleton, WI, USA). The selected stream outlet tube of the multi-position valve was connected to a gas chromatograph (GC-450, Bruker, Billerica, USA) equipped with a thermal conductivity detector (TCD) for measurement of CO 2 and a pulsed discharge detector (PDD, Vici AG International, Schenkon, Switzerland) for N 2 O and N 2 . The sample gas outlet of the GC was connected to a flow-through massflowmeter (Alicat Scientific, Tucson, AZ, USA), and a trace-level gas analyzer (CLD 88Yp, Eco Physics AG, Dürnten, Switzerland) equipped with a chemoluminescence detector (CLD) to analyze NO concentrations. To add up to the required 300-mL-flow of the NO analyzer, samples were diluted with synthetic air. Processing of GC data was done using CompassCDS software (SCION Instruments, Livingston, UK). Data from the NO analyzer and flowmeter were read out every 10 s via a serial port.
The analytical precision of the GC was determined by repeated measurements of standard gases (CO 2 , N 2 O, N 2 ) and was consistently < 2%. Detection limits were 0.08 µg N 2 O-N kg −1 h −1 and 5.5 µg N 2 -N kg −1 h −1 . The non-selected outlet streams of the multiposition valves were used to sample headspace gas for analysis of isotopic compositions (δ 13 C of CO 2 , isotopocules of N 2 O). After 47 days, the pots were flushed with pure helium to establish anoxic conditions to determine potential denitrification. After 8 days of anoxic incubation (55 days in total), the pots were opened for final sampling.

13 CO 2 Sampling, Analysis, and Calculations
For determination of δ 13 C of soil-emitted CO 2 , samples were flushed into 12 mL Exetainer ® septum-capped vials (Labco, High Wycombe, UK). Samples were taken twice a day for the first 5 days, daily for the next 12 days, every second day for the next 14 days, and every 3 days until day 43. Samples were introduced by a Combi-Pal autosampler (CTC-Analytics, Zwingen, Switzerland) to a GC (GC-Box, Thermo Fisher Scientific, Bremen, Germany) coupled to an isotope ratio mass spectrometer (Delta plus XP, Thermo Fisher Scientific, Bremen, Germany) via a Conflo III Interface (Thermo Fisher Scientific, Bremen, Germany). The fractions of CO 2 derived from litter (f litter ) and SOM (f SOM ) were calculated using Equations (1) and (2): f litter + f SOM = 1 (2) where δ 13 C treatment is the measured δ 13 C (‰) of CO 2 from litter treatment, δ 13 C Control is the measured δ 13 C (‰) of CO 2 from control treatment without litter addition, and δ 13 C litter is the mean measured δ 13 C (‰) of CO 2 lost from maize litter (Leaf: −7.91 δ‰, Root: −7.50 δ‰, Straw: −9.33 δ‰, see supplement for details, Figure S2). For each treatment, the priming effect (PE) was calculated as the difference between SOM-derived CO 2 -C (C SOM ) and CO 2 -C from control treatment without litter (C Control ) (3): Site preference (δ 15 N SP N2O ) was calculated as the difference between δ 15 N α N2O and δ 15 N β N2O . We used the scrambling factor of 0.096 determined by Buchen et al. (2018) [31] to correct measured data [32]. δ 18 O of soil water was −6.7 δ‰.
We applied the isotopocule mapping approach by Lewicka-Szczebak et al.  where δ r is the isotopic signature of residual N 2 O after partial reduction, δ 0 is the isotopic signature of initial N 2 O before reduction, and η red is the isotopic fractionation factor associated with N 2 O reduction to N 2 . Two main scenarios were considered: (1) N 2 O emitted from bacterial denitrification is first reduced to N 2 and residual N 2 O is then mixed with N 2 O originating from nitrification or fungal denitrification (Scenario 1, reduction-mixing). Alternatively, (2) N 2 O from bacterial denitrification and nitrification or fungal denitrification is first mixed and then partially reduced to N 2 (Scenario 2, mixing-reduction).
Recently, non-overlapping signatures for N 2 O produced by nitrification or fungal denitrification were proposed [23], and we calculated both scenarios for mixing of bacterial denitrification with either nitrification or fungal denitrification. A detailed description of the calculations can be found in [33]. We used the isotopic fractionation factors proposed by [23] (Supplementary Table S1), which were corrected for δ 18 O of soil water (−6.7 δ‰) for mapping and calculations.
In addition, calculations can be based on minimum or maximum end-member values, fractionation factors, and reduction factors, leading to a total of 14 different scenarios [31]. In our study, we used mean values for mixing, fractionation, and reduction whenever possible. However, as during anoxic incubation, samples were distributed outside the mean mixing-reduction area, we used minimum reduction values (mean mixing, mean fractionation) for 51 and 54 DAO. When calculations yielded values < 0 or > 1 for fbD or rN 2 O, these values were removed from the dataset before calculating means and plotting.

Soil Analyses
Samples of pre-incubated soil were taken prior to experimental setup. After opening pots at the end of the experiment, soil from each pot was homogenized and a sample was taken for analyses. Subsamples were analyzed for soil mineral N, water-extractable organic C (WEOC), and soil water content. For analysis of mineral N (NO 3 − and NH 4 + ), 50 g fresh soil were weighed into plastic bottles and frozen at −20 • C until analysis. Frozen samples were extracted with 2 M KCl solution (1:5 w:v) and shaken on an overhead shaker for 60 min. Samples were filtered with 615 1 4 filter paper (Macherey-Nagel GmbH & Co. KG, Düren, Germany) and extracts were analyzed colorimetrically using the San ++ Continuous-Flow Analyzer (Skalar Analytical B.V., Breda, The Netherlands). To determine isotopic signatures of nitrate, the bacterial denitrification method with Pseudomonas aureofaciens was applied [34,35].
WEOC was extracted by homogenizing 2 g of fresh soil with 10 mL H 2 O bidest . Samples were centrifuged, filtered with 0.45 µm polyether sulfone filters (Labsolute, Renningen, Germany), and stored at −20 • C. The extracts were analyzed for organic C and total N content using a multi N/C ® Analyzer (Analytik Jena, Jena, Germany). Soil water content was determined by oven drying at 105 • C.

Calculations and Statistics
For all calculations and statistical analyses, the statistical software R version 3.6.0 [36] was used. Fluxes of CO 2 , N 2 O, N 2 , and NO (F, µg kg −1 h −1 ) were calculated using the dynamic chamber approach (7): where C o is the concentration at the outflow and C i is the concentration at the inflow of each vessel (mg N m −3 , or mg C m −3 ), Q is the flow rate through the headspace (m 3 h −1 ), and m is the dry mass of soil per vessel (kg). Net N mineralization was calculated according to Equation (8): Mean values and standard deviations were calculated using the SlidingWindow function from the package evobiR v.1.1 [37] or the rollapply function from the package zoo [38].
Cumulative emissions were calculated by interpolation between measured fluxes. To test for differences between treatments, a one-way ANOVA was calculated when data were normally distributed or the Kruskal-Wallis rank sum test for non-normally distributed data followed by the LSD post hoc test. A t-test at p < 0.05 was used to compare soil NO 3 − , NH 4 + , and WEOC content before and after the incubation. To analyze the effect of litter input and litter quality on CO 2 and N emissions, simple linear regression models were tested. In all plots, color schemes from the R package viridisLite v0.3.0 [39] were used.

Characterization of Maize Litter
Maize litter types differed in their chemical composition (Table 1). Total C content ranged between 40% in maize roots and 47% in maize leaves. Total N content ranged between 3.8% in maize leaves and 0.85% in maize straw. C:N ratio was highest in maize straw (51.4) and similar in maize leaves and roots (12.3 and 13.8, respectively). Watersoluble C contents were similar in all maize litter types (8-8.5%). Water-soluble N content was highest in maize roots (1.22%) and lowest in maize straw (0.39%). Thus, water-soluble C:N was highest in maize straw and lowest in maize roots. 13 C-CPMAS NMR spectroscopy of maize litter revealed that maize straw and maize leaves were closer in their chemical composition than maize roots (Table 1, spectra in Supplementary Figure S1). Maize roots were characterized by the lowest shares of alkyl C and carboxyl C and the highest share of O/N-alkyl C, while maize leaves had highest shares of carboxyl C and alkyl C.

Soil N and C Content
Soil NO 3 − content increased in Control, Leaf, and Root treatments during the incubation experiment due to a net mineralization of N ( Table 2). In contrast, addition of maize straw significantly decreased soil NO 3 − content and immobilized N. Soil NH 4 + content strongly decreased in all treatments during the incubation period and was significantly higher in all maize litter treatments than in Control, but differences between treatments were small. WEOC content increased in all maize litter treatments, but did not change in Control. No differences were found in soil WEOC content between different litter treatments at the end of the experiment. δ 15 N NO3 and δ 18 O NO3 of added KNO 3 were higher compared to initial soil NO 3 − at onset of incubation (Table 2). At the end of the incubation experiment, δ 15 N NO3 and δ 18 O NO3 of soil NO 3 − differed between litter treatments. The lowest δ 15 N NO3 was measured in Root, and the lowest δ 18 O NO3 in Control and Leaf. The highest δ 15 N NO3 and δ 18 O NO3 were measured in Straw. δ 15 N NO3 increased with decreasing net N mineralization (adj. R 2 = 0.73, p < 0.001, Table 3). Table 1. Characteristics of maize litter used in the incubation experiment: total and water-soluble C and N content, C:N ratios, δ 13 C, and distribution of C species in different plant litter types (values represent the percentage contribution of the different integrated chemical shift regions determined by 13   Values represent means ± standard deviation (n = 5, Control and Initial n = 4). Different letters in the same column indicate a significant difference according to the LSD post hoc test at p ≤ 0.05. *** indicates a significant difference to Initial content according to t-test at p ≤ 0.05. 1 initial soil NO 3 − after pre-incubation/added KNO 3 (means ± standard deviation, n = 4). 3.3. CO 2 and 13 CO 2 Fluxes and Cumulative Emissions CO 2 fluxes from all litter treatments increased after onset of incubation compared to Control (Figure 1a,b, Supplementary Figure S3). Total CO 2 fluxes were highest in Leaf reaching 5.1 mg C kg −1 h −1 on 2 DAO. In Root, CO 2 flux peaked on 2 DAO (2.57 mg C kg −1 h −1 ) and then decreased throughout the incubation period. In Straw, the highest CO 2 fluxes were measured directly after onset of incubation (2.8 mg C kg −1 h −1 ), and continuously decreased afterwards. Litter-derived and SOM-derived CO 2 followed a similar pattern as total CO 2 fluxes and were highly correlated (adj. R 2 = 0.88, p < 0.001, Table 3). Highest litter-derived CO 2 fluxes were measured in Leaf on 3 DAO (3.0 mg C kg −1 h −1 ), in Straw on 1 DAO (1.2 mg C kg −1 h −1 ), and in Root on 2 DAO (1.7 mg C kg −1 h −1 ) (Figure 1a). SOM-derived CO 2 was highest in Leaf and higher in all litter treatments compared to Control for the first week after onset of incubation ( Figure 1b). Accordingly, the cumulative priming effect increased most strongly in all litter treatments during the first days of incubation ( Figure 1c) with highest values in Leaf.
Cumulative CO 2 emissions from all litter treatments were significantly higher than from Control without litter (p < 0.05, Table 4). The highest cumulative and litter-derived CO 2 emissions were measured after addition of maize leaves, followed by maize straw and maize roots; however, cumulative SOM-derived CO 2 emissions were higher than litterderived CO 2 emissions in all treatments (Table 4). Total cumulative CO 2 emissions were significantly positively correlated with total C and water-soluble C input from maize litter (adj. R 2 = 0.80 and adj. R 2 = 0.90, respectively, p < 0.001, Table 3). When total cumulative CO 2 emissions were standardized against C input from litter, no differences were found (Supplementary Table S2).
Highest litter-derived CO2 fluxes were measured in Leaf on 3 DAO (3.0 mg C kg −1 h −1 ), in Straw on 1 DAO (1.2 mg C kg −1 h −1 ), and in Root on 2 DAO (1.7 mg C kg −1 h −1 ) (Figure 1a). SOM-derived CO2 was highest in Leaf and higher in all litter treatments compared to Control for the first week after onset of incubation (Figure 1b). Accordingly, the cumulative priming effect increased most strongly in all litter treatments during the first days of incubation (Figure 1c) with highest values in Leaf.

N Fluxes and Cumulative Emissions
During the oxic incubation phase, only N 2 O and NO fluxes were measured as N 2 fluxes were below the detection limit (Figure 2a,b). N 2 O fluxes from litter treatments were higher than 3.7 µg N kg  During the oxic phase, cumulative N2O emissions from litter-amended soil were higher than from Control (p < 0.05, Table 5). Cumulative emissions in Straw were higher than in Root and similar to Leaf. Cumulative NO emissions were highest in Control and lowest in Straw, and NO/N2O ratio was significantly higher in Control than in litter-amended treatments. NO emissions strongly decreased with decreasing litter C:N ratio (adj. R 2 = 0.86, p < 0.001) and increased with increasing N mineralization (adj. R 2 = 0.57, p < 0.001) confirming that litter quality affected nitrification-derived NO emissions during the oxic incubation phase.
After 47 days, anoxic incubation conditions were induced by flushing the headspace with pure helium gas. N2O, NO, and N2 fluxes strongly increased with onset of anoxic incubation conditions (Figure 2c-e). N2O and NO fluxes peaked on 48 DAO and then decreased until the end of the experiment. N2 fluxes increased after onset of anoxic conditions until the end of the experiment. During the anoxic phase, cumulative N2O, NO, and N2 emissions were higher in litter treatments than in Control, although the effect was not always statistically significant for maize roots ( Table 5). The highest cumulative emissions were measured for NO (0.64 mg N kg −1 ) and N2O in Leaf (29.8 mg N kg −1 ), and for N2 in Straw (8.4 mg N kg −1 ). The ratio of the gaseous end products N2O/(N2O+N2) was highest in Control (0.95) and lowest in Straw (0.75). During the oxic phase, cumulative N 2 O emissions from litter-amended soil were higher than from Control (p < 0.05, Table 5). Cumulative emissions in Straw were higher than in Root and similar to Leaf. Cumulative NO emissions were highest in Control and lowest in Straw, and NO/N 2 O ratio was significantly higher in Control than in litteramended treatments. NO emissions strongly decreased with decreasing litter C:N ratio (adj. R 2 = 0.86, p < 0.001) and increased with increasing N mineralization (adj. R 2 = 0.57, p < 0.001) confirming that litter quality affected nitrification-derived NO emissions during the oxic incubation phase. After 47 days, anoxic incubation conditions were induced by flushing the headspace with pure helium gas. N 2 O, NO, and N 2 fluxes strongly increased with onset of anoxic incubation conditions (Figure 2c-e). N 2 O and NO fluxes peaked on 48 DAO and then decreased until the end of the experiment. N 2 fluxes increased after onset of anoxic conditions until the end of the experiment. During the anoxic phase, cumulative N 2 O, NO, and N 2 emissions were higher in litter treatments than in Control, although the effect was not always statistically significant for maize roots ( Table 5). The highest cumulative emissions were measured for NO (0.64 mg N kg −1 ) and N 2 O in Leaf (29.8 mg N kg −1 ), and for N 2 in Straw (8.4 mg N kg −1 ). The ratio of the gaseous end products N 2 O/(N 2 O+N 2 ) was highest in Control (0.95) and lowest in Straw (0.75).

N 2 O Isotopocule Mapping Approach, fbD and rN 2 O Values
The δ 15 N SP /δ 18

N2O Isotopocule Mapping Approach, fbD and rN2O Values
The δ 15 N SP /δ 18 ON2O isotopocule map showed a strong influence of the incubation day on the isotopic signature of soil-emitted N2O (Figure 3). Most data points were distributed between the mixing line of bacterial and fungal denitrification and the N2O reduction line during the oxic incubation phase (0-47 DAO). With onset of anoxic incubation conditions, bacterial denitrification became the dominant process as samples measured on 48 DAO cluster tightly above the reported ranges for heterotrophic bacterial denitrification. With ongoing anoxic incubation, the samples cluster along the reduction line indicating increasing N2O reduction with ongoing anoxic incubation conditions (51 and 54 DAO).   Table S1 for details). The mixing line connects the mean values of bD and fD (mix_bD-fD) or bD and ni (mix_bD-ni), respectively. The slope of the reduction lines (red) is based on the isotopic fractionation factor associated with N 2 O reduction to N 2 . Dashed line represents the minimum reduction line (n = 178, oxic incubation conditions from 0 DAO to 46 DAO, anoxic incubation conditions from 47 DAO to 55 DAO). δ 15 N bulk and δ 18 O N2O of soil-emitted N 2 O followed a similar pattern. Both values increased slightly during anoxic incubation (Figures 4 and S6). With onset of anoxic conditions, both δ 15 N bulk and δ 18 O N2O decreased strongly and increased again until the end of the experiment.
(corrected for δ 18 OH2O) for heterotrophic bacterial denitrification (bD), nitrifier denitrification (nD), nitrification (ni), and fungal denitrification (fD) (view Table S1 for details). The mixing line connects the mean values of bD and fD (mix_bD-fD) or bD and ni (mix_bD-ni), respectively. The slope of the reduction lines (red) is based on the isotopic fractionation factor associated with N2O reduction to N2. Dashed line represents the minimum reduction line (n = 178, oxic incubation conditions from 0 DAO to 46 DAO, anoxic incubation conditions from 47 DAO to 55 DAO). δ 15 Nbulk and δ 18 ON2O of soil-emitted N2O followed a similar pattern. Both values increased slightly during anoxic incubation (Figure 4, Figure S6). With onset of anoxic conditions, both δ 15 Nbulk and δ 18 ON2O decreased strongly and increased again until the end of the experiment. fbD and rN2O values exhibited similar patterns for mixing between heterotrophic bacterial denitrification/nitrifier denitrification and nitrification or fungal denitrification (Figure 5a,b, supplementary Figure S6). After onset of incubation, the fraction of soil-emitted N2O from heterotrophic bacterial denitrification/nitrifier denitrification (fbD, Figure 5a) was in tendency higher in maize litter treatments compared to Control. While fbD decreased in maize litter treatments during the oxic incubation period, it increased in Control without litter addition. With onset of anoxic incubation conditions, fbD increased strongly in all treatments, reaching values > 0.9, indicating that bacterial denitrification became the dominant process under anoxic incubation conditions. The residual, unreduced N2O fraction (rN2O, Figure 5b) was mostly < 0.5 and decreased during the oxic incubation phase, highlighting the significance of N2O reduction. On 51 DAO, rN2O was highest and decreased until 54 DAO.

Interactions between C and N Availability and N Fluxes
To test the effect of C availability and SOM turnover on N fluxes and cumulative emissions, simple linear regression models were tested ( Table 3).
The relationship between hourly NO+N2O and CO2 fluxes was very weak during the oxic incubation phase (adj. R 2 = 0.08, p < 0.001, Table 3). In contrast, NO+N2O+N2 fluxes were highly positively correlated with CO2 fluxes during the anoxic incubation period fbD and rN 2 O values exhibited similar patterns for mixing between heterotrophic bacterial denitrification/nitrifier denitrification and nitrification or fungal denitrification (Figure 5a,b, Supplementary Figure S6). After onset of incubation, the fraction of soilemitted N 2 O from heterotrophic bacterial denitrification/nitrifier denitrification (fbD, Figure 5a) was in tendency higher in maize litter treatments compared to Control. While fbD decreased in maize litter treatments during the oxic incubation period, it increased in Control without litter addition. With onset of anoxic incubation conditions, fbD increased strongly in all treatments, reaching values > 0.9, indicating that bacterial denitrification became the dominant process under anoxic incubation conditions. The residual, unreduced N 2 O fraction (rN 2 O, Figure 5b) was mostly < 0.5 and decreased during the oxic incubation phase, highlighting the significance of N 2 O reduction. On 51 DAO, rN 2 O was highest and decreased until 54 DAO. lated with total litter C input (adj. R 2 = 0.51, p < 0.001, Table 3). Furthermore, we found a significant negative correlation between the ratio of cumulative N2O/(N2O+N2) emissions during anoxic incubation and the ratio of water-soluble C:N of maize litter types (adj. R 2 = 0.51, p < 0.001, Table 3) and also with the soil NO3 -:WEOC ratio at the end of the experiment (adj. R 2 = 0.41, p < 0.01, Table 3). When standardized against C input from litter, total cumulative N emissions did not differ between litter treatments (Supplementary Table  S2).  [22] and represent results for scenario 1 (reduction-mixing) of bacterial denitrification with fungal denitrification (mean and standard deviation for n = 5, n = 4 for Control, data points missing for samples with an isotopic signature outside the reduction-mixing area, no N2O emitted from Control on 39 DAO).  [22] and represent results for scenario 1 (reduction-mixing) of bacterial denitrification with fungal denitrification (mean and standard deviation for n = 5, n = 4 for Control, data points missing for samples with an isotopic signature outside the reduction-mixing area, no N 2 O emitted from Control on 39 DAO).

Interactions between C and N Availability and N Fluxes
To test the effect of C availability and SOM turnover on N fluxes and cumulative emissions, simple linear regression models were tested ( Table 3).
The relationship between hourly NO+N 2 O and CO 2 fluxes was very weak during the oxic incubation phase (adj. R 2 = 0.08, p < 0.001, Table 3). In contrast, NO+N 2 O+N 2 fluxes were highly positively correlated with CO 2 fluxes during the anoxic incubation period (adj. R 2 = 0.86, p < 0.001, Table 3). Similarly, fbD and rN 2 O were positively correlated with total CO 2 flux from soil, but the relationship was weak (adj. R 2 = 0.23 and 0.31, respectively, p < 0.001, Table 3).
Cumulative anoxic N emissions (NO+N 2 O+N 2 ) were significantly positively correlated with total litter C input (adj. R 2 = 0.51, p < 0.001, Table 3). Furthermore, we found a significant negative correlation between the ratio of cumulative N 2 O/(N 2 O+N 2 ) emissions during anoxic incubation and the ratio of water-soluble C:N of maize litter types (adj. R 2 = 0.51, p < 0.001, Table 3) and also with the soil NO 3 − :WEOC ratio at the end of the experiment (adj. R 2 = 0.41, p < 0.01, Table 3). When standardized against C input from litter, total cumulative N emissions did not differ between litter treatments (Supplementary Table S2).

Maize Litter Quality Controls N Mineralization
Soil mineral N content, mineralization, and immobilization strongly depended on maize litter quality. In Control without litter addition, soil NH 4 + content decreased, but soil NO 3 − content strongly increased due to mineralization and nitrification of soil organic N. Tillage often leads to increased soil mineral N content [40,41], and net N mineralization from control soils without litter addition has been reported [8,13,42]. In Leaf and Root, mineralization was higher than in Control as additional organic N from litter was mineralized.
In contrast, addition of maize straw immobilized N, which coincides with the higher C:N ratio in Straw compared to Leaf and Root. In general, immobilization of N follows the addition of litter with C:N ratios > 25:1 [7]. Net N mineralization after addition of maize roots is in contrast to most other studies reporting net N immobilization after addition of maize roots [8,13,42,43]. However, we used maize roots grown in a nutrient solution, which had higher total and water-soluble N content than those reported in other studies [13,44].

Effect of Maize Litter Quality on CO 2 Emissions and Priming Effect
After onset of incubation, both litter and SOM-derived CO 2 fluxes strongly increased in litter treatments, while CO 2 efflux in Control was stable. Total cumulative CO 2 emissions increased with increasing input of total and water-soluble C, indicating that decomposition dynamics were controlled by the amount and quality of added plant material. The chemical composition of plant litter is known to be a primary controller of decomposition rates of both roots [11,12,15] and aboveground plant litter [4,10].
Especially during the first week after litter addition, SOM turnover was increased in all litter treatments, leading to a positive priming effect. The highest increase in SOM turnover was observed after addition of maize leaves which were characterized by high amounts of easily degradable compounds as indicated by high shares of carboxyl C (i.e., organic acids, amino acids) and alkyl C (i.e., fatty acids, amino acids, paraffines) [45]. Litter and SOM-derived CO 2 fluxes followed the same pattern and were highly positively correlated confirming that litter and SOM turnover are interrelated. Thus, our results are in accordance with the concept that litter with high degradability increases SOM turnover leading to a positive priming effect [17][18][19].

Effect of Litter Quality and N Mineralization on N Emissions and Production Pathways under Oxic Atmosphere
Directly after onset of oxic incubation, NO fluxes were highest in Control and Root while N 2 O fluxes were higher in litter amended treatments than in Control. The ratio of NO/N 2 O can be used as an indicator whether NO is produced from nitrification or denitrification [46,47]. While the NO/N 2 O emission ratio of bacterial denitrification is mostly around 0.01, the emission ratios of NO/N 2 O from nitrification are often higher than 1 [47]. In our study, the emission ratio of NO/N 2 O was highest during the first 10 days after onset of incubation, with maximum values of 0.47 in Control indicating a high contribution of nitrification to NO formation. Analysis of soil samples taken prior to the onset of incubation revealed high NH 4 + content of soil, which further supports that nitrification contributed to NO emissions during the initial incubation phase. In Straw, where N was immobilized to decompose C compounds, NO fluxes decreased faster and were lower than in all other treatments. Furthermore, oxic cumulative NO emissions strongly decreased with increasing litter C:N ratio (adj. R 2 = 0.86, p < 0.001) and increased with increasing N mineralization (adj. R 2 = 0.57, p < 0.001) confirming that litter quality affected nitrification-derived NO emissions in the beginning of the oxic incubation phase.
Addition of maize litter increased N 2 O fluxes compared to non-amended Control. As first, the headspace atmosphere had to be replaced by the mixture of He/O 2 , measurements started approximately one day after onset of incubation conditions. At this time, decreasing N 2 O fluxes indicated that N 2 O fluxes had peaked within 24 h after water and NO 3 − addition. After this initial increase, N 2 O fluxes decreased rapidly and then stayed on a similar level throughout the oxic phase. Immediately after onset of the incubation, bacterial denitrification (i.e., heterotrophic bacterial denitrification and nitrifier denitrification) was the dominant N 2 O emitting process in litter-amended treatments as indicated by the fraction of bacterial denitrification (fbD) > 0.6. Gradually decreasing fbD values then indicate a shift towards nitrification or fungal denitrification. Litter addition [27,48] and soil moisture of 70% WFPS may have promoted fungi, which often contribute to denitrification under weakly anoxic conditions [49,50]. Several studies have described a shift from bacterial to fungal dominance with ongoing incubations [27,[51][52][53]. However, nitrification may have contributed to N 2 O formation in Root and Leaf as indicated by high mineralization and NO/N 2 O ratio. In Control, nitrification presumably contributed to N 2 O formation, especially during the first days of the experiment, when fbD was < 0.3 and the NO/N 2 O ratio was high. rN 2 O values were mostly < 0.5, highlighting the significance of N 2 O reduction, also during the oxic incubation period. Thus, although N 2 fluxes were lower than the detection limit of our incubation system (5.5 µg N 2 -N kg −1 h −1 ), they significantly contributed to gaseous N losses. N 2 O reduction to N 2 is the last step of denitrification [54] and it usually takes place when availability of NO x is limited [55]. Furthermore, pore size and distribution, and soil moisture may affect N 2 O reduction to N 2 , as they control diffusion of O 2 and N 2 O in soil [20,56]. Accordingly, for interpretation of the isotopocule mapping approach in our experiment, we think that the reduction-mixing scenario is more plausible: N 2 O was produced by denitrifying bacteria and partly reduced to N 2 in anoxic microsites, and then N 2 O diffusing out of these hotspots was mixed with N 2 O from nitrification and fungal denitrification [57]. We anticipate that nitrification contributed to N 2 O formation when mineralization was high, while fungal denitrification became more important in litter-amended treatments with ongoing incubation.

Effect of Maize Litter Quality and Mineralization on Potential Denitrification
With onset of anoxic incubation conditions on 47 DAO, total NO and N 2 O fluxes increased rapidly, while N 2 fluxes increased more slowly. δ 18 O N2O of N 2 O emitted on 48 DAO falls in the range of heterotrophic bacterial denitrification reported in earlier studies [23], indicating that heterotrophic bacterial denitrification was the main N 2 O-emitting process with low reduction to N 2 . Interestingly, the δ 15 N bulk values on 48 DAO strongly deviated from measured δ 15 N bulk values on all other sampling days and were slightly outside the reported endmember values of heterotrophic bacterial denitrification. Under oxic conditions, denitrification mostly took place in anoxic hotspots where ongoing reduction led to a fractionation in the NO 3 − pool undergoing denitrification, which is reflected in gradually increasing δ 15 N bulk values. With onset of anoxic conditions, previously unreduced NO 3 − contributed to N 2 O formation leading to a shift towards more negative δ 15 N bulk values [29,58]. When the contribution of pools with different N dynamics changes, shifts in the isotopic signature have been reported [59][60][61] [23]. δ 15 N NO3 was higher at the end of the experiment compared to initial soil NO 3 − and added KNO 3 , confirming the ongoing fractionation during the reduction of the soil NO 3 − pool. Furthermore, δ 15 N NO3 increased with decreasing mineralization, indicating a higher share of added KNO 3 to residual NO 3 − at the end of the incubation experiment. Higher δ 18 O in Straw and Root may point towards a higher contribution of fungal denitrification, which is consistent with the higher contribution of fungi to decomposition of plant materials rich in celluloses and lignin [16,27]. Overall, it needs to be taken into account, that estimating N 2 O formation processes based on N 2 O isotopomers is subject to large uncertainties. Endmember values, reduction and fractionation factors have been obtained under differing incubation and environmental conditions, and may thus lead to over or underestimation of contributing processes [57,61,62].

Interaction between C Turnover and Denitrification
In agricultural soils, denitrification is often controlled by the availability of readily decomposable organic matter with increasing C availability leading to increased N losses [6,13,14,63,64]. In contrast, we measured low denitrification derived N fluxes under oxic conditions, and the correlation between oxic N and CO 2 fluxes was very weak indicating that denitrification was not directly affected by litter decomposition in our study. Although soil NO 3 − content was high and high litter and SOM turnover confirmed high C org availability, N 2 O fluxes were very low, indicating that conditions for denitrifying microorganisms were not optimal. Rohe et al. (2021) [65] reported very low N 2 O + N 2 fluxes from an incubation study with the same soil at 60% WFPS and higher fluxes compared to our study for 75 and 85% WFPS. Thus, our incubation conditions with a soil moisture of 70% WFPS may have been too low to promote denitrifying soil microorganisms. With onset of anoxic conditions, N fluxes increased immediately, confirming that high pO 2 was restricting denitrification during the oxic incubation phase. In contrast to our expectations, the microbial respiration of litter and SOM did not promote the formation of litter associated anoxic hotspots for denitrification as high O 2 diffusivity limited denitrification [65,66].
In contrast, N and CO 2 fluxes were highly positively correlated (adj. R 2 = 0.86, p < 0.001) under anoxic conditions, and cumulative N emissions increased with increasing litter C input (adj. R 2 = 0.51, p < 0.001) confirming our hypothesis that higher C availability leads to increased gaseous N losses. However, this effect was based on the role of C as energy source for denitrifiers, as the potential O 2 consumption by C decomposition was not relevant under anoxic conditions. The ratio of denitrification end products N 2 O/(N 2 O+N 2 ) decreased with increasing water-extractable C:N ratio of litter (adj. R 2 = 0.73, p < 0.001) and increasing soil WEOC:NO 3 − ratio at the end of the experiment (adj. R 2 = 0.41, p < 0.01) confirming that the ratio of available C to oxidized N is a major control of denitrification product stoichiometry [67]. Immobilization of N after addition of maize straw with high C:N ratio restricted N availability leading to higher N 2 O reduction to N 2 . However, as soil NO 3 − content was still very high (> 70 mg NO 3 − -N kg −1 in all treatments at the end of the incubation experiment), N 2 O was the dominant end product, as NO 3 − is preferentially used as electron acceptor and high soil NO 3 − content can inhibit the reduction of N 2 O to N 2 [27,67].

Conclusions
We investigated the effect of different maize litter types (young leaves and roots, straw) on CO 2 , NO, N 2 O, and N 2 emissions under oxic and anoxic conditions in a laboratory incubation study. Addition of maize litter increased litter and SOM derived CO 2 emissions, leading to a positive priming effect. SOM priming was highest after addition of maize leaves with a high share of easily degradable C compounds during the first week after onset of incubation. Although litter and SOM turnover were high, NO and N 2 O fluxes were low under oxic conditions as high O 2 diffusivity limited denitrification.
The NO/N 2 O ratio indicated that nitrification contributed to NO and N 2 O formation during the first two weeks of incubation, especially in Control without litter addition. In the litter-amended treatments, isotopocule mapping revealed that bacterial denitrification dominated N 2 O formation in the beginning of the incubation experiment with a subsequent shift towards fungal denitrification. With onset of anoxic incubation conditions after 47 days, N fluxes strongly increased and heterotrophic bacterial denitrification became the dominating source of N 2 O. The N 2 O/(N 2 O + N 2 ) ratio decreased with increasing litter C:N ratio and C org :NO 3 − ratio in soil confirming that the ratio of available C:N is a major control of denitrification product stoichiometry.
Supplementary Materials: The following are available online at https://www.mdpi.com/article/10.3 390/app11115309/s1, Figure S1a-c: Solid state 13 C-CPMAS NMR spectra of maize litter used in the incubation experiment, Figure S2: δ 13 C of CO 2 derived from maize litter, Figure S3: Total CO 2 efflux from soil during oxic and anoxic incubation, Figure S4a: δ 13 C of CO 2 evolving from soil and b: fraction of litter-derived CO 2 , Figure S5: NO/N 2 O ratio during oxic and anoxic incubation, Figure S6: δ 18 O of N 2 O, added KNO 3 , and soil NO 3 − at first and last day of incubation, Figure S7a: Fraction of N 2 O originating from heterotrophic bacterial denitrification/nitrifier denitrification and b+c: fraction of residual unreduced N 2 O,  Table S2: Cumulative CO 2 , NO, N 2 O, and N 2 emissions and denitrification product ratio standardized against litter C input.