Particulate organic matter as a functional soil component for persistent soil organic carbon

Study site and soil sampling

The soil was collected at 5 − 20 cm (Ap horizon) from an agricultural field in Southern Germany (Freising, Bavaria, 48°23’53.8“N, 11°38’39.7“E) in December 2017. The sampling area is situated within the lower Bavarian upland, and characterized by a mean annual temperature of 7.8 °C and mean annual precipitation of 786 mm. The soil type is a Cambisol (silty clay loam; 32% clay, 53% silt, and 14% sand) with a considerable amount of loess mixed with underlying Neogene sandy sediments. The soil was selected to represent a widely distributed soil type and land use. The collected soil was oven-dried (2 days, 40 °C), sieved (<2 mm), and visible plant remains were manually removed using tweezers.

Experimental setup

The experimental design involved four treatments; soils of two textures, either with or without ¹³C-labeled maize stalks. In order to obtain a coarse-textured soil (sandy clay loam; 24% clay, 15% silt, and 60% sand), half of the initial soil was mixed with cleaned quartz sand (Quarzwerke, Frechen, Germany) to increase the sand content from 14% to 60% (Supplementary Table 3). To achieve consistent bulk densities between treatments (0.9–1.3 g cm⁻³) 120 g (for coarser texture) and 90 g (for finer texture) soil was filled homogeneously and gently packed into microcosms (height: 5 cm, internal diameter: 5 cm, total volume: 98.2 cm³; polyoxymethylene, 1.4 g cm⁻³; Sahlberg, Munich, Germany). While the control microcosms were filled entirely with soil, 330 mg of air-dried and grounded ¹³C-labeled maize stalks (2–3 mm, δ¹³C = 2129 ± 82‰ V-PDB; Supplementary Table 4; Agroscope, Zurich, Switzerland) were mixed into the upper 1.67 cm of the soil within the other microcosms to create a quasi-natural gradient, with aboveground litter addition from the top (Fig. 9). Maize was chosen as litter substrate as it is a crop grown worldwide in agricultural systems. Each of the four treatments was replicated five times. The microcosms were sealed from below with polyester gauzes (37-µm mesh) and placed into Ball Mason Jars (475 ml) on top of metal grids to ensure downward gas diffusion.

Heterotrophic respiration

After making all containers gas-tight and rinsing them with synthetic air (Westfalen AG, Münster, Germany), 12 ml of gas samples (IVA Analysentechnik, Meerbusch, Germany) were collected from the headspace of the Mason Jars on day 2, 3, 4, 8, 10, 15, 23, 31, 44, 65, 80, and 95. For each measurement of CO₂ respiration, two samplings of the container atmosphere were carried out, and the time in between the two samplings was adapted to the current respiration rates. During the incubation period of 95 d, the CO₂ concentration, as well as the ¹³C abundance in the respired CO₂, was measured via gas chromatography isotope ratio mass spectrometry (GC/IRMS; Delta Plus, Thermo Fisher, Dreieich, Germany). The CO₂ levels were calibrated against three calibration gases (890, 1500, and 3000 ppm CO₂; Linde AG, Pullach, Germany). Then, CO₂ with known isotopic composition, diluted in helium, was used as a lab standard. This standard was in turn calibrated against three international standards (RM 8562, RM 8563, and RM 8564; International Atomic Energy Agency, Vienna, Austria) with a dual inlet system. The temperature and water holding capacity were kept constant at 21 °C and 60%, respectively, along with the incubation period.

Sampling

Following 95 days of incubation, each microcosm was cut into three horizontal sections with a razor blade, separating the top, center, and bottom layer (each 1.67-cm high). The microcosms were designed to be opened from the side, allowing for precise separation of the depth increments (Supplementary Fig. 4). Subsamples for subsequent microbial analyses were freeze-dried and stored at 4 °C, and dried aliquots for fractionation were stored in sealed plastic containers at 20 °C. Furthermore, a few POM particles were selected manually for NanoSIMS measurements.

Physical fractionation and subsequent analyses

The soil was separated into five distinct OM fractions using a combined density and particle size fractionation scheme¹¹. Air-dried soil (18–20 g) was gently capillary-saturated with sodium polytungstate solution (Na₆[H₂W₁₂O₄₀]; 1.8 g cm⁻³) and after 12 h, the free-floating particulate organic matter (fPOM) was collected using a vacuum pump. oPOM was released from aggregated soil structures via ultrasonic dispersion (Bandelin, Sonoplus HD 2200; energy input of 440 J ml⁻¹)⁴² allowing its separation from heavier minerals. The excess salt was removed from the oPOM by washing it with deionized water over a sieve (20-µm mesh size), which yielded an oPOM fraction of <20 µm (oPOM_small). Both fPOM and oPOM fractions were washed several times using deionized water and pressure filtration (20-µm mesh) until the solution dropped below an electric conductivity of <5 µS cm⁻¹ via pressure filtration. The oPOM_small fraction was cleaned via saturation with deionized water for 24 h. While sand and coarse silt fractions were separated by wet sieving, mineral fractions <20 µm were separated via sedimentation, and later combined as one MAOM fraction. The C, N, and ¹³C contents were determined for freeze-dried and milled OM fractions, as well as milled bulk soil, via dry combustion with an isotope ratio mass spectrometer (Delta V Advantage, Thermo Fisher, Dreieich, Germany) coupled with an elemental analyzer (Euro EA, Eurovector, Milano, Italy). Acetanilide was used as a lab standard for calibration and to determine the isotope linearity of the system, and was in turn calibrated against several suitable isotope standards (International Atomic Energy Agency, Vienna, Austria). International and lab isotope standards were included in every sequence to create a final ¹³C correction. Since the samples did not contain carbonates, the C contents were assumed to be equal to organic C contents.

¹³C nuclear magnetic resonance spectroscopy

The chemical compositions of the POM fractions were determined via ¹³C CP-MAS NMR in solid-state (Bruker DSX 200, Bruker BioSpin GmbH, Karlsruhe, Germany), where samples were filled into 7-mm zirconium dioxide rotors and spun in a magic angle spinning probe at a rotation speed of 6.8 kHz and 0.01024 s acquisition time. The recorded ¹³C spectra were quantified in the following chemical shift regions: alkyl C (−10–45 ppm), O/N alkyl C (45–110 ppm), aromatic C (110–160 ppm), and carbonyl/carboxyl C (160–220 ppm)¹¹. The regions were integrated and an alkyl C:O alkyl C ratio (−10–45/45–110 ppm) was computed to describe the degree of aliphaticity of the different fractions⁴³. Lastly, the obtained spectra were transformed into OM compound classes via the molecular mixing model^18,44 with the following chemical shift regions: 0–45, 45–60, 60–95, 95–110, 100–145, 145–165, and 165–215 ppm.

Calculations of litter-derived C in CO₂, soil, and OM fractions

Along with the incubation period, the amount of C respired per hour was computed (Eq. 1).

where ΔCO₂/Δt is CO₂ increase over time, V_HSP is the volume of the headspace of Mason Jars, the volume of an ideal gas at 21 °C is set at 24.1, and 12 represents the atomic mass of C.

Subsequently, the percentage of respired CO₂ originating from the litter was calculated (Eq. 2).

where δ¹³C_resp emission gives the δ¹³C for the current CO₂ emission between the two samplings (‰ V-PDB), δ¹³C_control is the average δ¹³C of the control soils at the time of measurement, and δ¹³C_litter is the δ¹³C signature of the labeled litter. Finally, the respired C originating from the soil was computed (Eq. 3).

The proportion of litter-derived C (%) in the OM fractions was calculated (Eq. 4)⁴⁵.

where δ¹³C_labeled is the ¹³C enrichment in labeled samples, δ¹³C_control is the ¹³C enrichment in controls (natural abundance level, i.e., 28‰ V-PDB), and δ¹³C_litter is the ¹³C enrichment in the added litter (i.e., 2129‰ V-PDB) from which the amount of litter-derived C within each OM fraction could then be determined (Eq. 5).

where C_fraction is the amount of C in mg g⁻¹, and m is the recovered mass (g) of each fraction after the fractionation.

PLFA analyses

The PLFA patterns were analyzed⁴⁶ and adjusted according to the ISO/TS 29843-2:2011F standard. In summary, the soil lipids from 3 g of soil (freeze-dried aliquots) were extracted with a Bligh and Dyer solution [methanol, chloroform, and citrate buffer (pH = 4 ± 0.1), 2:1:0.8, v/v/v]. A biphasic system was achieved by adding chloroform and citrate buffer from which the lipid phase was evaporated at 30 °C under a nitrogen stream. The phospholipids were separated from neutral lipids and glycolipids by solid-phase extraction on silica tubes (SPE DSC-Si, 500 mg, Discovery^®) and evaporated. The PLFA were turned into fatty acid methyl esters (FAMEs) via alkaline methanolysis⁴⁷ and later quantified via gas chromatic retention time comparison with a gas chromatograph (GC Agilent HP6890, G1530A, Chemstation, Santa Clara, USA) connected to a flame ionization detector equipped with a capillary column (SGE, BPX5, 60 m × 0,25 mm × 0,25 mm). The FAME concentrations were quantified relative to methyl nonadecanoate (19:0), enabling methylated lipids to be identified. A standard soil was used and extracted in parallel to detect potential deviations between the extraction rounds, expressed in nmol C-FA per g of soil. Mono-unsaturated and cyclopropylated PLFA (C16:1w7c, C18:1w9c, and C18:1w9t) were assigned to gram-negative bacteria, iso-branched and anteiso-branched PLFA (iC15:0, aC15:0, iC16:0, i-C17:0, C:17, and C18:0) were assigned to gram-positive bacteria and C18:2w6c, C18:3w3c, respectively C20:5w3c were assigned to fungi⁴⁸. The total content of bacteria was expressed by adding gram-positive, gram-negative together with the markers C14:0, C16:0, C20:0, and C15:1. Lastly, the ¹³C-labeling of FAME was concluded by correcting for the added methyl moieties during methanolysis and relating it to the chain length of fatty acids (Eq. 6).

where δ¹³CF_A represents the δ¹³C of the fatty acid, C_n the number of C atoms in the fatty acid, δ¹³C_FAME is the δ¹³C of the fatty acid methyl ester, and δ¹³C_MeOH is the δ¹³C of the methanol used for the methylation (−63%) to calculate the isotope ratios of the fatty acids. The relative incorporation of ¹³C into four microbial groups was calculated by relating the proportions of each fatty acid to the total ¹³C incorporation, and the absolute incorporation of ¹³C in each microbial group was calculated by dividing the amount of ¹³C enriched fatty acid by the total amount of extracted fatty acid for that particular group.

SEM and NanoSIMS microspectroscopy

In order to gain insights into the microscale distribution of the assemblages of litter with microbes and minerals, we used SEM and NanoSIMS. Free POM from non-fractionated soil was hand-picked and fixed onto graphene sample substrates on metal stubs (10 mm in diameter). To avoid the charging phenomena, samples were gold-coated prior to SEM analyses by physical vapor deposition under an argon atmosphere (Emitech Sputtercoater SC7620, Gala Instrumente, Bad Schwalbach, Germany). To analyze the microscale structures of the assemblages of POM, microorganisms and soil minerals of the samples were first analyzed using SEM (Jeol JSM 5900LV, Freising, Germany), and subsequently, the spots that best exemplified the microbial transformation on the decaying litter (POM) surface were analyzed using a Cameca NanoSIMS 50 L (Cameca, Gennevilliers, France)⁴⁹. For the NanoSIMS measurements, a 270-pA high primary beam was used to locally sputter away impurities and gold coating, and to implant primary ions (Cs⁺) into the sample’s surface (impact energy of 16 keV) to enhance the yields of secondary ions. Subsequently, secondary ions were measured using electron multipliers; ¹²C⁻, ¹³C⁻, ¹²C¹⁴N⁻ to display OM fragments and ¹⁶O⁻, ²⁸Si⁻, ²⁷Al¹⁶O⁻, and ⁵⁶Fe¹⁶O⁻ secondary ions to record the mineral phase. The instrument was tuned to a high mass resolution in order to accurately separate mass isobars at mass 13 (¹³C⁻, ¹²C¹H⁻). The ion images were acquired with a 25 × 25 µm field of view, 40 planes and 1 ms pixel⁻¹ dwell time for all measurements. Charging effects were compensated for with an electron flood gun if necessary. The acquired measurements were dead time (44 ns) and drift corrected using the OpenMIMS plugin of the ImageJ software. The ¹³C⁻🙁¹²C⁻ + ¹³C⁻) and ¹²C¹⁴N⁻🙁¹²C⁻ + ¹²C¹⁴N⁻) ratios were computed for distinct regions of interests which were chosen manually with respect to the major compartments: continuous fragments of fungal hyphae, individual bacteria, EPS patches, as well as exposed POM surfaces. To account for instrumental mass fractionation, the electron multipliers were carefully checked, and the control measurements of non-labeled POM samples were conducted regularly along with the sessions. Here, the mean ¹³C⁻🙁¹²C⁻ + ¹³C⁻) ratios were in line with the level of natural abundance, which meant that a correction of ratios for labeled POM samples was not necessary.

Statistical analyses

All parameters were separately tested for normality with Shapiro–Wilk test and for homoscedasticity with Bartlett’s test. In addition, the distribution of the datasets was checked with Q–Q plots. In cases where the assumptions of normality or homoscedasticity were not met, a log-transformation was applied on the raw data, and analyses were carried out on the log-transformed data. The differences caused by texture and litter addition were tested using unpaired t-tests, and depth differences were tested using one-way analysis of variance with Tukey’s honestly significant difference as the post-hoc test. In cases where the log-transformed data did not meet the requirements for parametrical testing, the unpaired two-samples Wilcoxon test or Kruskal-Wallis test was applied. The statistical findings were considered significant if the confidence limits were in excess of 95% (P < 0.05). All statistical testing was carried out in the R statistical environment⁵⁰ using agricolae⁵¹ and ggpubr⁵² packages.

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Source: Ecology - nature.com