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.