Keywords: Soil organic matter decomposition, carbon and nitrogen cycling, microbial activation, maize, plant intraspecific competition, root trait, arable soil
1. Introduction
The coexistence of organisms in unfertilized soils is characterized by strong competition for nutrients between plant communities and between plant roots and microorganisms (Kuzyakov & Xu 2013; Moreau et al. 2015; Adler et al. 2018). Soil organic matter (SOM) is a major reservoir of essential nutrients required for plant growth, but the availability of these nutrients, notably nitrogen (N), largely relies on microbial mediated transformation. Hence, microbial N mineralization from SOM, and subsequent immobilization/nitrification are crucial processes determining the intrinsic N supply of soil for plant productivity in natural and agroecosystems (Tiessen, Cuevas & Chacon 1994). The microbially driven processes can be significantly regulated by plant roots, as root-derived inputs of labile organic compounds (i.e., rhizodeposits) fuel the activity and growth of heterotrophic microorganisms (Paterson 2003; Cheng & Kuzyakov 2005). This, in turn, stimulates the synthesis of extracellular enzymes to mine for nutrients necessary for microbial growth, causing the acceleration of SOM mineralization (microbial N mining hypothesis; Craine et al., 2007). As a result, microbial N mineralization and immobilization tend to increase (Zhu et al. 2014; Murphy, Baggs, Morley, Wall & Paterson 2015; Ehtesham & Bengtson 2017), facilitating plant N uptake (Dijkstra, Bader, Johnson & Cheng 2009; Frank & Groffman 2009). Such root-mediated changes in SOM mineralization have increasingly been recognized as a nutrient acquisition strategy of plants to exchange carbon (C) to soil microorganisms for N and other nutrients (Kuzyakov & Xu 2013). Hence, to consider the processes by which plant roots govern SOM turnover has far-reaching implications for understanding plant-microbial-soil interactions in terrestrial ecosystems.
Previous studies have shown that root-mediated changes in SOM mineralization vary with plant growth stages and plant-plant interactions (Cheng et al. 2014; Huo, Luo & Cheng 2017). Root physiological and morphological traits are major factors regulating SOM turnover, such as root biomass (Dijkstra & Cheng 2007), the quality and quantity of root exudates (Zhu & Cheng 2012), root architecture and morphology (Pauschet al. 2016). Root traits are spatially and temporally dynamic, changing in response to its controlling factors such as light, soil nutrients and water content (Kuzyakov et al. 2002; Craine, Wedin, Chapin & Reich 2003; Sanaullah, Chabbi, Rumpel & Kuzyakov 2012), and associated plant performance (i.e., photosynthesis) throughout the growth stages (Bardgett, Bowman, Kaufmann & Schmidt 2005). The alterations of root traits may shape microbial function and consequently modify SOM mineralization, through the changes in the abundance of available C and nutrients and in soil physical and chemical properties, i.e., water, pH values and soil aggregation (Shields, Paul, Lowe & Parkinson 1973; Jenkinson & Rayner 1977).
Plant-plant interactions often impose interspecific and intraspecific competition for above and belowground resources, i.e., light and nutrients. However, the effect of plant-plant interactions on soil C and N turnover through rhizosphere processes remains uncertain, since contradictory results have been shown previously (Fan, Zhang & Lu 2011; Pausch, Zhu, Kuzyakov & Cheng 2013; Yin, Dijkstra, Wang, Zhu & Cheng 2018). With increased plant competition for nutrients, the plant may increase C allocation to roots and adjust root physiological and morphological traits that enhance their competitive capacity relative to their neighbors, thereby potentially enhancing the C-N exchanges and SOM mineralization (Tilman 1990; Schenk 2006; Kunstler et al. 2016). In contrast, it has previously been reported that SOM mineralization can also be suppressed under the interspecific (Dijkstra, Morgan, Blumenthal & Follett 2010; Pausch et al. 2013) and intraspecific competition (Yin et al., 2018). These negative effects are explained by the nutrient competition hypothesis (Dormaar, 1990; Kuzyakov, 2002), which suggests that strong competition for nutrients between roots and microorganisms may inhibit microbial activity for decomposing SOM due to nutrients limitation. In addition to nutrients, plants compete for light with denser canopies, leading to the uncertainty outcome of photo-assimilate supply to belowground and hence SOM mineralization (Aerts 1999; Wang et al. 2020). Despite several studies that have examined the effects of plant competition on SOM turnover, they were mostly conducted under controlled conditions with a restricted soil volume for nutrient uptake by plants (Dijkstra et al. 2010; Pausch et al. 2013; Yin et al. 2018; Schofield et al. 2019; Wang et al. 2020). Direct estimation of plant competition on SOM mineralization and the relevance of microbial mechanisms under field conditions still remain elusive.
In natural ecosystems, the microbial activation by roots and subsequent mining for N from SOM has been identified to be an essential driver for the coupling of C and N turnover of soils (Phillips, Finzi & Bernhardt 2011; Finzi et al. 2015). However, conventional agriculture often assumes that the N supply of N from SOM decomposition is inadequate to meet the plant demands of plants, especially for crops with a high N uptake rate, e.g. maize (Zea mays) (Loecke, Cambardella & Liebman 2012; Osterholz, Rinot, Liebman & Castellano 2017). Thus, maize has received much more N-fertilizers than other crops worldwide (FAO 2006). Intensive fertilization may hamper the reliance of plant N-uptake on N mineralization, causing the potential decoupling of soil C and N cycling (Drinkwater & Snapp 2007). Very few experiments have been conducted to examine the processes by which roots regulate SOM decomposition and N mineralization in the agriculture field, though it is essential in the context of fertilization management in the agroecosystem (Franciset al. 2003; Spiertz 2010).
Therefore, this study aims to investigate how plants control the coupling between C-input and SOM turnover for nutrient uptake in an arable soil under field condition, with a focus on assessing the temporal dynamics at different plant growth stages and the effects of plant intraspecific competition. Experimental trials with varying plant densities were established. We grew maize (C4 plants) with three planting densities for 132 days on a C3 soil (C3-to-C4 vegetation change; Kumar, Kuzyakov & Pausch 2016). The 13C natural abundance approach was used to partition total CO2 efflux for SOM-derived CO2 and root-derived CO2, and in situ15N pool dilution was applied to quantify gross N transformation. Concurrently, soil and microbial properties, and root morphology were measured at three plant growth stages (heading, flowering and maize ripening). We hypothesize that i) root mediated changes in SOM decomposition are associated with N mineralization during plant growth because of microbial activation by roots for N mining, and ii) the modulation of SOM decomposition and N mineralization by intraspecific competition depends on the root traits for nutrient uptake and the soil mineral nutrient status.
2. Materials and methods
2.1 Study site
The experiment was conducted on an agricultural field at the “Reinshof” research station of the Georg-August University of Göttingen (51°29′37.2″N and 9°55′36.9″E). The study area has a temperate oceanic climate with an annual mean temperature of 8.5 °C and a mean precipitation of 850 mm. The meteorological parameters during the experimental monitoring period are shown in Fig. S1. The soil is classified as a Haplic Luisol with a silty loam texture (74.9% silt, 3.2% sand, 21.9% clay) (Berger 1999). It contains 1.41 ± 0.04% total C and 0.16 ± 0.002% total N and has a bulk density of about 1.3 g cm−3 from 0-35 cm depth and a pH of 6.2 (Kumar, Dorodnikov, Splettstößer, Kuzyakov & Pausch 2017; Mason-Jones, Schmücker & Kuzyakov 2018). The organic C at the site originates from permanent C3 vegetation (δ13C of soil = -25.1 ‰). To separate root-derived C from SOM-derived C, a vegetation change from C3 to C4 (maize; Zea mays L. cv. Colisee ; δ13C of maize = -13.3 ‰) crops was applied. This allowed introducing a distinct 13C signal into the soil with a difference in δ13C values between soil and plant is > 11.8 ‰.
2.2 Experimental setup
Sixteen experimental plots (each with an area of 5 × 5 m) were aligned in 4 rows in the field, each with 2-m wide buffer stripes to exclude the neighboring effects. A gradient of plant densities was established following a random design, with 4 replicates each: 1) a plant density of 5 plants m-2 as the control (P); 2) a double plant density of 10 plants m-2 (DP), which is equivalent to the common maize plant density for conventional farming in Germany; 3) a triple plant density of 15 plants m-2 (TP). Additionally, four plots were kept free from vegetation as bare fallow.
Maize seeds were firstly sown in plant-treated plots with a density of 15 plants m-2. They were manually thinned to the respective low and double plant density 30 days after planting (DAP). Before maize sowing, conventional tillage practices were operated up to 30 cm of soil depth and all plots received phosphorus (P) and potassium (K) fertilizers. Temperature sensors (32 PT-100 sensors) were installed at 10 cm depth to monitor soil temperature. Soil moisture (0-10 cm depth) was measured by 6 ECH2O EC-5 moisture sensors (decagon devices). Hourly air temperature and atmospheric pressure were obtained from the weather station of the German Weather Service in Göttingen.
2.3 Sampling and analyses
2.3.1 Plant and soil sampling
Plants and soil were sampled at 72, 102 and 30 days after planting. The shoots of the plants were randomly sampled from each plot and weighed after oven-drying at 60 °C for 48 hours. Since the roots of maize plants are mainly concentrated in the upper 30 cm (Amos & Walters 2006), soils and roots were collected together by soil cores (~7 cm diameter) from four soil depths at 0–5 cm, 5–15 cm, 15–25 cm and 25–35 cm in the middle of the diagonal between two plants. In the lab, root samples were separated from soils by sieving and were washed. Roots were then scanned on a flatbed scanner and analyzed for length and diameter (WinRhizo, Regent Instruments Inc., Quebec City, Canada). After scanning, the roots were dried at 60 °C for 48 hours and weighed.
Microbial biomass C (MBC) and microbial biomass N (MBN) were determined for all depth at each of the three sampling times by the chloroform fumigation-extraction method with modifications (Vance, Brookes & Jenkinson 1987). Briefly, 8 g of fresh soil was extracted with 40 mL of 0.05 M K2SO4 after shaking for 60 min on a reciprocating shaker (Laboratory shaker, GFL 3016) and the filtrates were measured for total extractable C and N with a multi C/N analyzer (multi C/N analyzer 2100S, Analytik, Jena). The same extraction procedure was used for fumigated soil, which was fumigated with ethanol-free CHCl3 at room temperature for 24 h. Extractable organic C and N of non-fumigated soil were used as a measure of dissolved organic C (DOC) and dissolved N (DN). MBC and MBN were calculated as a difference of total extractable organic C and N between fumigated and non-fumigated samples using the extraction efficiency of 0.45 and 0.54 for C and N, respectively (Joergensen & Mueller 1996). Besides, 10 g soil of each core was oven-dried at 105°C for 24h to determine the gravimetric water content.
2.3.2 Respiration measurements
From July to late October 2015, soil CO2 efflux was measured in situ using pre-installed soil chambers at 54, 74, 90, 102, 122, 132 days after the planting. The vented static chambers made of polyvinyl chloride (area 0.05 m2 and approx. 14.5 L total volume) were inserted ~2 cm into the soil in the center of each plot for the entire measurement period. Concurrently, 20 mL gas samples were collected at 20-min intervals using a syringe and stored in pre-evacuated exetainer vials with rubber septa (Exetainer; Labco Limited, Lampeter, UK) and analyzed for the δ13C of CO2 using an isotope ratio mass spectrometer (IRMS) (Finnigan Delta plus XP, Thermo Electron Corporation, Germany).
2.3.3 Gross N mineralization and nitrification
Gross rates of N mineralization and nitrification were measured in situ three times at 74, 102, and 132 days’ after planting, which corresponds to the heading, flowering and ripening stages of maize plants (Meier 2001). The 15N pool dilution approach by intact soil cores was used to estimate gross N mineralization (GNM) and gross nitrification rates (GNN) (Davidson, Hart, Shanks & Firestone 1991; Hart, Nason, David D & A 1994). In each plot, five intact soil cores (5 cm high with a volume of 251.2 cm3) were taken: 15NH4Cl solution consisting of a mixture of 0.6 µg 15N g−1 soil (99 atom% 13C, Sigma Aldrich) and 2.4 µg14N g−1 soil, was added to two of the five cores for determining gross N mineralization; K15NO3 solution, a mixture of 0.6 µg15N g−1 soil (99 atom%13C, Sigma Aldrich) and 2.4 µg 14N g−1 soil, were added to another two cores with for determining gross nitrification. Water was added to the remaining cores for measuring the initial level of NH4+ and NO3-. The 15N enrichments for 15NH4Cl (or K15NO3) solutions were 20 atom%. One of each set of cores was well mixed and extracted 10 minutes after15N labeling (T0 soil cores), while the others from each pair were extracted after 24 of incubation (T1 soil cores). Briefly, 80 g soil from each core was extracted with 210 ml of 0.05 mol L-1 K2SO4 after shaking for 60 min on a reciprocating shaker (Laboratory shaker, GFL 3016). The concentrations of NO3- and exchangeable NH4+ in extracts were measured with a continuous flow analyzer (Skalar Analytical, Breda, Netherlands). Then, the 15N enrichment of NH4+ and NO3- were determined by IRMS (Finnigan Delta plus XP, Thermo Electron Corporation, Germany) following the diffusion procedures (Murphy et al. 2003).
2.4 Calculations and statistics
The soil CO2 efflux rates are calculated as the slope of linear regressions describing the change in CO2concentration in the chamber headspace over time and are adjusted to field-measured air temperature and pressure during measurement. We used Keeling-Plots (Miller & Tans 2003) to calculate the δ13C values of pure soil CO2 without the admixture of atmospheric CO2. Afterward, a linear two-source isotopic mixing model (Phillips & Gregg 2001) was applied to partition total CO2 efflux into its sources, SOM- and root-derived CO2. Gross N mineralization and gross nitrification were calculated following the equations in (Davidsonet al. 1991; Sun, Schleuss, Pausch, Xu & Kuzyakov 2018).
To assess the potential intensity of the shoot and root competition, we use the modified version of relative competition intensity (RCI) according to (Callaway et al. 2002):
RCI = (X c - X t) / x,
where X c and X t is the shoot biomass (or root biomass, g m-2) in the control (P) and density treatments (DP and TP), respectively, and x is the highest value of (X c:X t). Positive RCI value denotes competition.
Statistical analysis
The experiment was conducted with 4 field replicates. Normality (Shapiro-Wilk test, p > 0.05) and homogeneity of variance (Levene test, p > 0.05) were examined and data were log-transformed prior to analysis if necessary. We implemented two-way ANOVA with plant growth stages as the first factor and plant density as the second factor to identify single and their interacting effects on the total CO2 efflux, root-derived CO2 efflux, SOM-derived CO2 efflux, GNM, specific root length and root-derived CO2 efflux. However, two-way ANOVA revealed hardly any interactions between plant growth stages and plant density, and plant growth stages to be the predominant factor controlling the rate of C and N cycling. Therefore, we test additionly the independent effects of plant growth stages and plant density with a post hoc unequal N Tukey-Kramer significant difference (HSD). For plant and soil parameters (Table 1 and 2), we first used two way ANOVA to assess the effects of planting density, growth stages and their interactions. In addition, significant differences of those parameters among the density treatment at each growth stage were obtained by a one-way ANOVA analysis. We used a one-tailed t -test to assess the significances between RCI and zero. All statistical analyses were performed with SPSS 22, with the significance level at p < 0.05. Simple regressions were used to identify relationships between response variables with significances at p < 0.05.
3. Results
3.1 Plant biomass and root morphology
The shoot biomass per m-2 increased from maize heading to ripening stages. Root biomass per m-2 remained unchanged with growth stages for the single and double densities, but it increased for the triple density (Table 1). For each growth stage, maize produced a similar shoot and root biomass in total at the double and triple planting densities, which were higher than that at the single density (Table 1). However, both shoot and root biomass decreased after normalizing for planting density at both double and triple planting densities, and this was consistent with the positive relative competition intensity (RCI) of shoots and roots (Table 1), indicating the strong above- and belowground intraspecific competition with increased planting density. For either double or triple planting density, a higher belowground intraspecific competition (as indicated by RCI of roots) was reached at flowering. The RCI of root at the triple planting density was higher than the double planting density (Table 1), and was negatively correlated to mineral N concentrations in soil (R2 = 0.6, p = 0.03; Fig. S2a).
For each planting density, maize had a higher root length density (RLD; in the upper 35 cm soil) at flowering stages. The RLD at the double and triple planting densities were averagely 1.7 and 2.2 times higher than the single planting density, respectively (across all growth stages; Table 1). At the heading and ripening stages, root length per unit of root biomass (specific root length; SRL) was similar among the planting densities, but the SRL at maize flowering stage increased by 46% and 49% at the double and triple densities, respectively (as compared with single density; Fig. 2a). Moreover, SRL increased with the RCI of root for maize at both double (R2 = 0.7, p = 0.001) and triple densities (R2 = 0.3, p = 0.05) (Fig. S2b).
3.2 Fluxes and sources of CO2
Total CO2 efflux ranged from 75 to 251 mg C m-2 day-1. It remained similar at the heading and flowering stages and then decreased by 64% at maize ripening, irrespective of planting density (Fig. 1a). The contributions of root- and SOM-derived sources to total CO2 efflux was similar between the planting densities but was dependent on maize growth stages. The root-derived CO2 averagely contributed 56%, 28% and 46% to the total CO2 at heading, flowering, and ripening stages, respectively (across planting density; Fig. 1a). Root-derived CO2 efflux significantly decreased from the heading to ripening (ranged from 39.8 to 135 mg C m-2day-1) independently of the planting density (Fig. 1b). Root-derived CO2 per unit of root biomass (specific root-derived CO2) declined with maize growth stages, and the double and triple planting densities led to lower specific root-derived CO2 effluxes than single density at the heading and flowering stages (Fig. 2b). SOM-derived CO2efflux was higher at maize flowering stage, followed by the heading and ripening stages for all levels of planting densities (ranged from 35.5 to 195.1 mg C m-2 day-1; Fig. 2c), but the planting density had a minor effect on SOM-derived CO2 efflux for each growth stage (Fig. 2c).
3.3 Soil N mineralization and nitrification
Maize growth stages, but not the planting density exerted significant control over gross N mineralization (GNM), which varied between 41.9 and 88.2 mg N m-2 day-1 (Fig. 2d). A higher rate of GNM occurred at maize flowering and it remained similar at the heading and ripening stages across all planting densities. Likewise, there was no difference between the planting density in gross nitrification (GNN) and the higher GNN at the flowering stage compared to other stages (Table 2). The GNN rates at heading and flowering stages were higher than the rates of GNM. Mineral N concentrations (exchangeable NH4+ plus NO3-) averagely decreased with maize growth by 40% at the flowering and 30% at ripening as compared with heading stages (across overall planting densities; Table 2).
3.4 Soil C and N content and microbial biomass
Dissolved N (DN) concentration in soil decreased with maize growth. For each growth stage, the double and triple densities led to similar amounts of DN contents, which were approximately 1.7 times lower than that at the single density (Table 2). Dissolved organic C (DOC) concentration remained similar at the heading and ripening, but decreased at maize flowering (Table 2). Soil microbial biomass C (MBC) gradually increased from the heading to ripening stages and the increase was up to 57%, 12% and 62% at the single, double and triple densities, respectively, while microbial biomass N (MBN) was independent on maize growth (Table 2).
3.5 Relationships between soil C and N mineralization
There was a positive linear relation between SOM-derived CO2 and GNM across all planting densities (R2 = 0.4, p < 0.04; Fig. 3a), indicating that the SOM-C mineralization is highly correlated with N mineralization. The C: N ratio of SOM mineralization (the ratio between SOM-derived CO2 and GNM) declined with growth stages, and it was reduced with increased planting densities compared to the single density at the earlier two growth stages (Table 2). Moreover, the C:N ratio of SOM mineralization increased with soil DN concentration (r2 = 0.3, p = 0.02; Fig. 3b). SOM-derived CO2 and GNM were independent on root biomass (Fig. S3a, b) and specific root-derived CO2 efflux (Fig. S3c, d). However, both SOM-derived CO2 and gross N mineralization increased with specific root length for the double and triple planting densities (Fig. 3a, b), indicating the root morphology modulated soil C and N turnover at higher planting densities.