Conservation tillage promotes crop productivity by changing soil aggregation in oasis farming ecosystems

doi:10.21203/rs.3.rs-3089715/v1

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Research Article

Conservation tillage promotes crop productivity by changing soil aggregation in oasis farming ecosystems

https://doi.org/10.21203/rs.3.rs-3089715/v1

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published 03 Jan, 2024

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Background Aims

Conservation tillage benefited soil aggregate function—essential for soil nutrient cycling and plant growth. Nevertheless, little is known about tillage practices' impact on soil particle aggregation, particularly soil nutrients and distribution of enzymes and microbes among different-sized aggregates, and their possible later influence upon other soil functions and processes.

Methods

We conducted a long-term experiment with maize (Zea mays L.) in a region of oasis farming in Northwest China, in which tillage was applied in four treatments: no-tillage, minimal tillage, fold-tillage, and sub-tillage. Soil aggregates were categorized as follow: <0.25 mm (‘micro’), 0.25–2 mm (‘small’), and > 2 mm (‘macro’), whose corresponding nutrient contents, enzymatic activity and stoichiometry were measured, as well as the structure of soil microbial community of those fraction, along with crop productivity.

Results

The no-tillage treatment increased the amount of N, P, and C in soil, the biomass of microbiota, and the availability P and N within micro-aggregates and bulk soil. It also augmented enzymatic activity for acquiring C- and P as well the enzymatic ratio of C: N but decreased that of N: P in micro-aggregates, and promoted crop productivity compared with conventional tillage. Additionally, microbial community structure differed under the four tillage regimes and between the aggregate fractions, especially under conventional tillage, but the tillage system did not affect alpha diversity.

Conclusions

Our results showed that no-tillage restructured the soil microbial community composition of aggregate fractions. Thus, long-term conservation tillage promotes soil functioning and crop productivity by changing soil aggregates in oasis farmland ecosystems.

No-tillage

Soil aggregates

Nutrient distribution

Microbial community structure

Soil functioning

Sustainable agriculture

Sustainable agriculture based on conservation tillage relies on healthy soil microbial properties (Saleem et al. 2019). Microbial community diversity benefits soil health, being responsible for soil nutrient cycling and maintaining soil function, structure, and productivity (Bérard et al. 2011; Griffiths and Philippot 2013). Moreover, microbial community composition is important for soil function, regulating substrate cycling and energy flow between plants and soil, and is an indicator of soil health (Kumar et al. 2018; Schmidt et al. 2018). However, for a given microbial community in soil, not only its composition and taxonomical diversity but also its activity may be sensitive to local changes in the environment. For example, in agriculture, certain tillage practices such as conventional and conservation tillage can alter soil microbial properties (Bünemann et al. 2006; Fierer et al. 2021).

Deep tillage (conventional practice) is used widely to prepare seedbeds, prevent weed and disease proliferation, and increase soil organic matter decomposition (Kumar et al. 2012). Using deep tillage also bolsters bulk density, but reduces its mineralizable nitrogen (N) content and weakens the soil’s capacity to hold water by disrupting soil structure and exposing soil aggregates to microbial activity (Karlen et al. 2013). In contrast, foregoing tillage entirely (i.e., no-tillage) or minimizing its use (i.e., reduced-tillage practices) have emerged as promising conservation practices to enhance various properties (physical, chemical, and biological) of farmed soil. No-tillage management is an important strategy for increasing soil nutrient storage and availability (Garland et al. 2018; Sá et al. 2017), aggregate stability (Bronick and Lal 2005), and the diversity and metabolic activities of the microbial communities associated with soil aggregates (Hsiao et al. 2018; Tian et al. 2022), in addition to preventing soil erosion. Ridge tillage, disk tillage, and strip tillage (among others)are examples of conservation tillage characterized by non-inversion, meaning that the proportion crop residue remaining on the ground is 30% or greater (Morris et al. 2010). Adopting no-tillage practices reduces soil disturbance and maintains crop residues on the soil surface or underground. No-tillage practices benefit soil geobiochemistry properties (e.g., aggregates, carbon storage, and microbes) more than conventional tillage (Endale et al. 2017; Morris et al. 2010). However, it is unknown whether no-tillage can mediate nutrient cycles, soil aggregate-associated microbial community properties, and their functions by changing soil aggregates in oasis farming regions.

Soil aggregates are an important component of soil structure, with their spatial configurations (shape, size, and pore network) produced through plant root growth and faunal activity (da Luz et al. 2022). Aggregates differing in size can harbor specific properties related to pore space size, nutrient content, aeration, water-holding capacity, and the availability of pores. For instance, organic matter occurring in a stable form (e.g., recalcitrant plant body) tends to be less prone to biological degradation in soil, being also usually more stable in size with a ˂0.25 mm (i.e., micro-aggregates) (Six and Paustian 2014). However, labile organic carbon, namely that occurring in either roots or freshly fallen litter, but also fungal hyphae, tends to be more commonly found in higher amounts in macro-aggregates (soil particle diameter ˃0.25 mm) (Plante et al. 2002). While crop residues add organic matter to the soil, high-intensity tillage breaks down macro-aggregates, accelerating soil organic C losses (Gami et al. 2009). Therefore, it is imperative to strengthen the structure of soil as well as the stability of its aggregates in order to improve soil C storage (Meurer et al. 2020), microbial activity and composition (Tian et al. 2022), and crop growth (Balesdent et al. 2000). Soil aggregate structure is determined by soil substrates inputted from plant root and shoot litter (Wander and Yang 2000) and organic fertilization (Tian et al. 2022).

Improved soil aggregate structure can foster greater spatial heterogeneity for microbes and microzoon dwelling in soil, and soil aggregate-associated communities of those microbiota are capable of influencing elemental cycling (Gupta and Germida 2015). Soil microbial properties in different-sized aggregates are often linked to key soil processes and nutrient cycling that are pivotal for sustaining various ecological functions (Trivedi et al. 2017). Further, the activity of soil enzyme is quite sensitive to variation in soil C as well as the nutrient availability of aggregates (Wang et al. 2015); hence, enzymatic stoichiometry has been deemed a reliable indicator of soil nutrient limitations or the availability of suitable substrates (Sinsabaugh et al. 2009). Consequently, the nutrient content and availability which characterizes soil aggregates differing in size can jointly impact the growth of plants. Yet surprisingly litter field research has investigated how tillage practices may affect arising among crops and belowground microbial communities and nutrients on the scale of aggregates, especially in arid regions.

Therefore, here we investigated the effects of two types of tillage regimes (conventional vs. conservation tillage treatments) upon the composition of microbial communities in soil and the latter’s functioning (i.e., ability to retain nutrients and their availability) at the aggregates in an oasis farming ecosystem. In a 7-year long field manipulation adjacent to the Tenggeli and Badanjaran Deserts, the site’s was subjected to conservation and/or conventional tillage management, to experimentally test three hypotheses: (1) Both the properties of tilled soil and the structure of its microbial community differ according to its –aggregate size fraction, whose variable specific soil functions are reflected in enzymatic activity and stoichiometry; (2) tillage practices mediate soil functioning by modulating the pattern of soil aggregates' distribution and the nutrient pool; (3) conservation tillage practices change the bulk soil C and its availability of nutrient, along with microbial community structure, thereby increasing plant productivity.

Study area

We carried out this research work in Gansu Province (China) in Minqin County, (103°07′ 16″E, 38^°37′10″N) situated on the Shiyang River Basin’s lower reaches in the Hexi Corridor, in an area bound by the Tenggeli and Badanjaran Deserts (Feng et al. 2011). In this area a typical arid continental climate prevails, whose mean annual air temperature and precipitation is respectively 7.8℃ and 113.2 mm (most of the latter falling between July to September), and an average annual evaporation totaling 2646 mm. The uneven rainfall distribution causes intense seasonal droughts in spring and/or winter. According to the, The research site has an irrigated desert soil (Gansu Provincial Soil Survey Office. 1992), this not unlike similar to Anthropic Camborthids according to Soil Taxonomy (Jiang et al. 2017). Feng et al. (2011) describe other relevant environmental conditions and agriculture production in this study area.

Design of the experimental and soil sampling

In 2015 the experiment was installed, by flat planting the field with spring maize (Zea mays L.) early on in April. The crop was allowed to grow until October’s end, when it was harvested. During each winter, all plots in the field were left fallow. When the experiment was begun, the soil pH was 8.63, and it 0.29 ms cm^–1 specific conductance, 9.80 g kg^–1 organic matter, 4.84 g kg^–1 of total N, 0.40 mg kg^–1 of available P, and 144.14 mg kg^–1 of available K. An RBD (randomized block design) was used for this maize experiment, in which tillage was applied in four treatments, each having three replicates (i.e., plots). The tillage systems mainly differed in the cultivation method practiced each autumn after harvesting the main crop: 1) conventional sub-tillage (ST): moldboard plow down to 30 cm; 2) fold-tillage with sub-soiling (FT): chisel plowing to 20 cm; 3) conservation minimal tillage (MT): no autumn cultivation and rotary tillage to 10 cm; 4) conservation no-tillage (NT) (Fig. 1). Supplemental irrigation of all maize plots was applied using basin irrigation at the seedling, jointing, pre-heading, silking, flowering, and grain-filling stages, respectively. Plots were 30 m × 20 m, separated by concrete borders.

The plow layer (0–20 cm depth) was sampled for soil, by using10 cm diameter auger inserted at five position (these chosen randomly) within given crop row, about 20 cm from maize stems during their harvest in October 2021; these soil sampled were then mixed to into a composite sample at the plot level. These freshly collected soil samples were stored in containers made of hard plastic; to safeguard their soil structure from destruction while in transit to laboratory, each container was filled with buffer foam pieces.

Soil aggregate fractionation

To isolate the aggregates, a modified dry-sieving method was used (Dorodnikov et al. 2009) to minimize the disturbance of soil microbes. In a well-ventilated place, sample was dried to ca. 10% gravimetric soil water content (‘optimal’ moisture), to limit the mechanical stress “to induce maximum, brittle failure along natural planes of weakness” (Dorodnikov et al. 2009). That is, along their natural breakpoints, individual soil clods were gently separated, then sieved through 8-mm mesh to remove any visible stones (and other debris). Then, each soil sample was divided into two parts: the first was for bulk soil analysis and aggregate fractionation, by using the following sieving procedure. Around 1 kg soil per plot was positioned over two sieves (2.0 and 0.25 mm mesh) and gently shaken thrice, for 2 min each time. Soil material that was left on the sieves was collected and respectively designated as large macro-aggregates (i.e., > 2 mm) and small macro-aggregates (i.e., 0.25–2 mm). Any remaining soil which passed through the 0.25-mm sieve was then collected, this being the micro-aggregate particles (hereon ‘micro-aggregates’) (i.e., < 0.25 mm). Next, we weighed each soil aggregate fraction and divided each into two portions (1) for analysis of available nutrients, microbial community biomass, diversity and composition, and soil enzymatic activity assays; and (2) for air drying prior to its analysis of soil properties (chemical).

To obtain the soil aggregate distribution, the respective proportions corresponding to each soil aggregate fraction were calculated, with the mean weight diameter (MWD) used an index of aggregate stability: MWD = \(\sum _{i}^{3}xi\times wi,\) where i denotes the number count of aggregate size fractions; xi denotes the mean diameter (mm) of the aggregates in any size fraction; and wi denotes the weight of the aggregates in that particular size fraction, expressed as a proportion of the total soil dry weight (Somasundaram et al. 2017).

Chemical and biological properties of soil

Soil chemical analysis

To determined soil moisture, the difference in mass before and after drying at 105℃ to constant weight was used. To quantify soil organic C (SOC), the Walkley and Black dichromate oxidation method was used with 1.08 as the factor. For total N (TN), it was measured by the K₂SO₄–CuSO₄–Se (100:10:1) distillation method; for total P (TP), the molybdate colorimetric method after perchloric acid digestion and ascorbic acid reduction was used. After first soaking them in 2 mol L^–1 KCl, soil samples were shaken (200 rpm, 1 h) and analyzed for their NH₄⁺-N and NO₃^–-N by an auto-flow injection system (Auto-Analyzer AA3, Germany). To quantify the available P (AP) in soil, we applied the Olsen-P method (Yuan et al. 2016). To determine the microbial biomass C (MBC) or N (MBN) in soil, the difference in values between the C or N extracted via chloroform fumigated vis-à-vis non-fumigated soil samples was obtained, to which the K_EC factor (0.45) or K_EN (0.57) was respectively applied. Briefly, 20 g fresh soil was fumigated for 24 h at 25°C with CHCl₃ and then added to 80 mL of 0.5 mol L^–1 K₂SO₄, shaken (200 rpm, 1 h) and filtered after removing the fumigant. A 20 g sample of non-fumigated soil was extracted in tandem. To determine the OC and TN contents of each extract a total C analyzer was used (TOC-L CPH Basic Analyzer System, Japan). In the non-fumigated samples, their organic C concentration corresponded to the dissolved organic C (DOC) (Wallenstein et al. 2006).

Soil enzyme assays

Four hydrolytic soil enzymes were quantified for their respectively activity. These consisted of two enzymes participating in C-acquisition (β-1,4-glucosidase and cellobiohyrolase), one enzyme participating in N-acquisition (β-1,4-N-acetylglucosaminidase), and one enzyme participating in P-acquisition (alkaline phosphatase) (Sinsabaugh et al. 2009). We also quantified the activities of two oxidases, phenoloxidase and peroxidase, which enable the degradation of recalcitrant organic C (Sinsabaugh 2010). To measure hydrolytic enzymes, a fluorescence microplate assay was employed, in which the substrates were labeled with 4-methylumbelliferone (MUB) (Bell et al. 2013). To do this, 1.25 g of fresh soil and 125 mL of 50 mmol L^–1 Tris buffer solution (pH 8.30) were mixed at 25°C for 20 min by an oscillator, this then serving as a soil buffer suspension. Next, 400 µL of that suspension and 100 µL of substrate solution (200 µmol L^–1) were pipetted into a 96-well microplate. Meanwhile, 400 µL of each sample mixture, a negative control, the reference standard, a blank control, and a quench control were pipetted into a black 96-well plate and incubated at 25°C for 3 h in the dark; to end the reaction, 20 µL of NaOH was added. Activity levels were measured fluorometrically, by a fluorescence plate reader (Varioskan LUX, ThermoFisher Scientific, USA) at excitation and emission of 365 and 450 nm, respectively, and expressed in units of substrate converted (µmol) per mL of sample (µmol g^− 1 h^− 1). Oxidases were measured spectrophotometrically in a clear 96-well microplate using the substrate L-3, 4-dihydroxyphenylalanine (DOPA). To represent the potential capacity of these C-, N-, and P-acquisition activities, their corresponding enzymes were grouped accordingly, and their data transformed (normalized) as follows: respectively, ln(β-glucosidase + cellobiohyrolase + phenoloxidase + peroxidase), ln(β-N-acetylglucosaminidase), and ln(phosphatase) (Sinsabaugh et al. 2009).

Soil microbial community composition

Soil DNA extraction and sequencing

From each soil sample (0.5 g) its microbial DNA was extracted by using the HiPure Soil DNA Kits (Magen, Guangzhou, China). The amplicons were assessed using 2% agarose gel electrophoresis and purified with an AxyPrep DNA Gel Extraction Kit (Axygen Bioscience, Union City, USA). From the bacterial 16S rDNA target region of the rRNA gene, the V3–V4 hypervariable region was PCR-amplified (conditions: 95°C for 2 min, then 35 cycles of 95°C for 30 s, 60°C for 45 s, and 72°C for 90 s; ending with a final extension 72°C for 10 min), using the primers 515F: GTGYCAGGMGCCGCGGTAA; 806 R: GGACTACNVGGGTWTCTAAT. To amplify the fungal ITS2 region, the primers ITS3-KYO2: GATGAAGAACGYAGYRAA and ITS4: TCCTCCGCTTATTGATATGC were used under the same condition. Each PCR reaction was performed in a 50-µL reaction volume that included TransGen High-Fidelity PCR SuperMix (TransGen Biotech, Beijing, China), 0.2 µM each F/R primer, and 5 ng of template DNA. To detect amplicons, 2% agarose gels were used, followed by purification with the AxyPrep DNA Gel Extraction Kit (Axygen Biosciences, Union City, CA, USA). To generate the sequencing libraries, SMRTbell TM Template Prep Kit (PacBio, Menlo Park, CA, USA) was used; to assess library quality, we used the Qubit 3.0 Fluorometer (ThermoFisher Scientific, USA) and FEMTO Pulse system (Agilent Technologies, Santa Clara, CA, USA). All the obtained sequence data has been deposited into the CNGB Sequence Archive (CNSA) of China National GeneBank DataBase (CNGBdb), where it can be found under accession number CNP0003819.

Bioinformatics analysis

To merge the paired-end clean reads (raw) FLASH (v 1.2.11) tool (Magoč and Salzberg 2011) was used with 10-bp minimal overlap and 2% mismatch error rate at most. Noisy reads sequences were screened by prescribed filtering conditions to obtain a subset of clean reads high in quality, as described by Bokulich et al. (2013). The retained clean reads were clustered into operational taxonomic units (OTUs) have ≥ 97% similarity in identity, by implementing the UPARSE pipeline (v 9.2.64) (Edgar 2013). Any chimeric sequences present were found and excised by running the UCHIME algorithm (Edgar et al. 2011). Bacteria were identified taxonomically by the RDP classifier (Wang et al. 2007) according to the SILVA database (Prüesse et al. 2007), and the fungi were classified using the UNITE database (Nilsson et al. 2018) with the BLAST tool. A suite of alpha diversity metrics, namely the observed number of species (richness), Pielou’s evenness, in addition to the well-known Shannon–Wiener index, as well as the Chao1 estimator, were derived for the communities of bacteria or fungi, using QIIME. Bioinformatics analyses were carried out in R software (v 2.2.1).

Statistical analyses

To assess the tillage intensity (regime) effect on aggregate stability (MWD), properties of soil, biomass of microbial, enzymatic activity and stoichiometry, aggregate size fraction, and microbial diversity, we used ANOVA (one-way model) followed by pairwise comparisons using Tukey’s HSD test. In addition, the single effects of each size fraction of soil aggregate on enzymatic activity and stoichiometry as well as soil properties under each tillage treatment were assessed by univariate ANOVAs (one-way). Statistical significance was designated using an alpha = 0.05 (i.e., P ˂ 0.05). All above ANOVAs and multiple mean comparisons were implemented in Genstat 18 software (VSN International Ltd., Rothamsted, UK); to draw the figures, we used Origin 9.2 software (OriginLab 2016, USA). Principal component analysis (PCA) was applied to based on OUT levels data, to examine in what way the structures of bacterial and fungal communities were related or differed between aggregate size fractions and tillage management regimes.

Aggregate distribution

The tillage regime significantly affected soil aggregate distribution (Fig. 2A). Conservation management (NT and MT) significantly increased macro-aggregate and micro-aggregate fractions relative conventional managements (FT and ST). Conventional management significantly decreased MWD relative to conservation management (Fig. 2B).

Conservation management significantly augmented the SOC, TN, and TP of bulk soil; values for SOC were greatest in the NT treatment, with the highest soil TN and TP in the MT treatment (Fig. 3A–C). NT had the highest bulk soil mineral N and available P contents (Fig. 3E, F), while ST had higher DOC contents than the other three treatments (Fig. 3D). Evidently, with reductions in aggregate size, other soil nutrients showed a trend of increasing that was similar (Fig. 3). The DOC content diminished in tandem with declining aggregate size in NT, MT, and FT. In contrast, micro-aggregates in the ST treatment had higher DOC contents than the other three treatments (Fig. 3D). More specifically, micro-aggregates mainly contributed bulk soil nutrients in the conventional and conservation treatments.

Enzyme activities and enzymatic stoichiometry

Tillage regime significantly affected enzyme activities and enzymatic stoichiometry among large, small and micro-aggregates (Fig. 4). The aggregate size fraction affected C-acquiring and P-acquiring enzyme activities, increasing with decreasing aggregate size (Fig. 4A, C); however, concerning the N-acquiring enzyme activity, it was similar across the aggregate size fractions under the FT and ST treatments (Fig. 4B). For enzymatic stoichiometry, aggregate size fraction affected the C: N enzyme ratio, increasing with reduction in aggregate size (Fig. 4D). By contrast, the enzyme ratio of N: P significantly decreased with declining aggregate size, being lower overall in FT and ST than either NT or MT (Fig. 4F). However, the C: P enzyme ratio followed no apparent trend among aggregate size fractions across tillage treatments (Fig. 4E).

Microbial community biomass, diversity, and composition

Tillage regime significantly affected soil microbial biomass within aggregate fractions (Table 1). Generally, as aggregate size decreased, both soil microbial C and N

Table 1

Soil microbial biomass C, N, and diversity under different tillage regimes. Values are means ± SD (n = 3). Different lowercase letters indicate significant differences among aggregate size fractions in the same tillage treatment (P ˂ 0.05), while different uppercase letters indicate significant differences among tillage treatments for each aggregate size class (P ˂ 0.05).
				Chao1 estimator		Pielou’s envenness		Shannon index
		MBC	MBN	Bacteria	Fungi	Bacteria	Fungi	Bacteria	Fungi
	LMA	5.20 ± 0.01cC	11.6 ± 1.22bC	4269 ± 217b	457 ± 23.7bA	0.81 ± 0.008	0.71 ± 0.033	9.7 ± 0.20	6.2 ± 0.38
NT	SMA	76.0 ± 4.68bB	13.5 ± 0.12ab	4266 ± 57.6bB	504 ± 18.0aA	0.81 ± 0.017	0.68 ± 0.049	9.7 ± 0.29	6.0 ± 0.43
	MIA	93.7 ± 8.59aA	15.4 ± 1.37aA	4942 ± 137aA	436 ± 10.7b	0.81 ± 0.017	0.68 ± 0.040	9.9 ± 0.20	6.1 ± 0.38
	LMA	18.9 ± 0.19cB	17.1 ± 0.83aA	4181 ± 61.5bA	375 ± 23.7bB	0.82 ± 0.005	0.64 ± 0.027	9.7 ± 0.08	5.4 ± 0.27
MT	SMA	70.3 ± 3.47bB	12.3 ± 2.25b	4268 ± 42.4abB	428 ± 17.4aB	0.82 ± 0.005	0.64 ± 0.028	9.9 ± 0.11	5.5 ± 0.21
	MIA	62.8 ± 2.95aC	11.8 ± 1.14bBC	4389 ± 107aB	379 ± 25.1b	0.82 ± 0.005	0.65 ± 0.014	9.9 ± 0.07	5.5 ± 0.06
	LMA	22.4 ± 1.91bB	14.7 ± 0.81aB	4477 ± 66.3bB	345 ± 31.9bB	0.82 ± 0.009	0.66 ± 0.060	9.8 ± 0.02	5.6 ± 0.54
FT	SMA	73.5 ± 1.37aB	14.8 ± 1.00a	4626 ± 118abA	310 ± 27.7bC	0.82 ± 0.001	0.59 ± 0.031	9.9 ± 0.09	4.9 ± 0.30
	MIA	73.9 ± 2.49aB	13.7 ± 1.00aAB	4795 ± 29.7aA	483 ± 67.5a	0.82 ± 0.005	0.59 ± 0.049	9.9 ± 0.12	5.1 ± 0.63
	LMA	60.0 ± 3.55bA	17.5 ± 0.37aA	4400 ± 84.7b	333 ± 13.4cB	0.82 ± 0.003	0.69 ± 0.072	9.8 ± 0.09	5.8 ± 0.44
ST	SMA	85.1 ± 5.72aA	13.5 ± 1.81b	4696 ± 85.8aA	376 ± 26.9bD	0.82 ± 0.003	0.66 ± 0.037	9.8 ± 0.10	5.6 ± 0.40
	MIA	30.7 ± 5.49cD	10.1 ± 0.83cC	4565 ± 82.9abB	485 ± 13.6a	0.83 ± 0.006	0.68 ± 0.038	9.9 ± 0.10	5.8 ± 0.39

LMA: large macro-aggregate; SMA: small macro-aggregates; MIA: micro-aggregates; MBC: microbial biomass C; MBN: microbial biomass N.

rose in the NT treatment, with the highest MBC and MBN values in micro-aggregates. In contrast, microbial C and N in the ST treatment decreased with decreasing aggregate size, with the highest MBC and MBN values in large macro-aggregates. Soil MBC gradually increased in large macro-aggregates with increasing tillage intensity (Table 1).

The Chao1 estimator for evaluating richness revealed that, in general, the bacterial community got diverse (richer) as aggregate size was reduced, irrespective of tillage treatments (Table 1). Compared to conventional tillage (ST), NT had significantly higher Chao1 values for micro-aggregate. Similarly, NT had significantly higher Chao1 values than ST for macro-aggregates. As estimated by the Shannon index and Pielou’s evenness, the diversity of bacterial and fungal communities did not significantly differ among tillage regimes or aggregate fractions.

The relative abundance of microbial communities for OTUs was assigned to > 10 classes. In every tillage treatment, the bacterial phylum most abundant was evidently Actinobacteria, followed by Proteobacteria, Chloroflexi, Acidobacteria, and Planctomycetes (Fig. 5). The identified bacterial OTUs had similar abundances regardless of the four tillage treatments or aggregate fractions. However, the PCA based on OTUs uncovered a significant modification to the structure of the bacterial community between conservation (NT and MT) and conventional (ST and FT) tillage (Fig. 6). Similarly, while the abundance of fungal community member were similar among all aggregate fraction or tillage treatment, the PCA revealed stark differences in fungal community composition between certain tillage treatments, namely for LMA, SMA, and MIA between FT and ST yet not between NT and MT (Fig. 6).

Crop productivity under different tillage regime

Long-term tillage regimes significantly affected the crop productivity of maize in three ways: yield of grain, biomass of straw, and root biomass (Fig. 7), as well as the root/shoot ratio. All these four aspects were increased significantly by applying conservation tillage (NT) relative to conventional tillage (ST and FT) (Fig. 7).

Tillage influences the pool of soil nutrients harbored by aggregate fractions

After seven years of tillage treatments, most soil nutrients had higher values under no-tillage than sub-tillage conditions. Similar findings from other studies (Endale et al. 2017; Leskovar et al. 2016), soil nutrient pools increased under conservation tillage (NT) relative to conventional tillage (ST), indicating that no-tillage management carried out over the long term enable organic C and available nutrients (especially mineral N and available P)to accumulate in soil. This is likely best explained by the fate of crop residues, which remained on the ground under no-tillage management and slowly decomposed, this generating a sustained input to both C and N sources for microbial communities in the topsoil (Li et al. 2018). Under no-tillage, the continuous input of organic residues increases soil microbial activity (Hungria et al. 2009), generating various exudations from microbes and plant roots (Six et al. 2002) that likely increase small organic particles in the soil, thereby forming a system that is functional porous system (Gonçalves et al. 2014). Soil microbial activity is generally driven by C and N resources, with the previous crop’s residues and roots being an important input source of C in agroecosystems (Li et al. 2018; Nunes et al. 2020). Soil no-tillage management resulted in better aggregation in tropical and sub-tropical soils by forming macro-aggregates (Six et al. 2002), consistent with the current study; however, as the mean aggregate size was reduced, the NT treatment increased the amount of C, P, and N, in soil. Our findings contradict those of Tian et al. (2022), who reported contents of soil C, P, N that instead increased with greater aggregate size on average, mainly due to macro-aggregates either small or large combining with organic materials and plant debris with high nutrient levels to support more microbial biomass. Indeed, the status of nutrients present in individual soil aggregates fraction can convey their respective contribution to the overall nutrient status of soil (Zhang et al. 2016). Our first hypothesis is supported in part by the results, while suggests no-till management is able to bolster the availability of nutrient in bulk soil by augmenting their concentrations in micro-aggregates given the latter’s substantial root biomass and microbial exudation. During the development of root, soil particles’ spatial distribution undergoes reorganization, this changing the size and structure of pores in soil (Lucas et al. 2019).

Enzyme distribution in aggregate fractions is dependent on the tillage system

We found that, in micro-aggregates, those enzymes involved acquiring C- and P were more active than in macro-aggregates, which is consistent with a pronounced microbial demand for both C and P resources (Luo et al. 2017). Whereas irrespective of size of aggregate, for N-acquiring enzymes their activity was similar, thus indicating that microbes incurred more or less equal N limitation among aggregates (Rosinger et al. 2019). That C- and P-acquiring enzyme activities increased with increasing tillage intensity in micro-aggregates is in accord with the relatively lower concentration of DOC and available P in micro-aggregates among treatments without ST. This also indicates that high-intensity tillage can alter the manner in which aggregate size modulates C- and P-acquiring enzymes. Moreover, that micro-aggregates had greater C and phosphatase activity is perhaps indicative of litter debris and organic matter less-humified in them as well as for the bulk soil overall (Rojo et al. 1990; Tian et al. 2022).

Conventional tillage increased the C: N enzyme ratio with decreasing aggregate size, indicating a relatively pronounced demand by microbes for soil organic C that micro-aggregates contain. This suggests that long-term conventional tillage influences the nutrient demands of microbes via alteration to the stoichiometry of soil resources. To be specific, soil C exposed in relatively greater amounts through high-intensity tillage would spur the production of more N-acquiring enzymes by microorganisms to satisfy their N demand and ensure homeostasis of microbial stoichiometry (Cleveland and Liptzin 2007). In contrast, we found more labile organic C in macro-aggregates under the no-till treatment in addition to their greater N supply capacity, whereas in micro-aggregates there was more organic C that was stabilized, imply that some microbes could be C-limited. However, conventional tillage decreased the enzyme ratio of C: P as aggregate size was reduced, indicating that within macro-aggregates their P mineralization rate was relatively low, perhaps because the strongly adsorbed P in macro-aggregates are not so easily accessible to either microbes in soil or plants, or both (Garland et al. 2018; Hunt et al. 2007). We interpret these finding as support for our second hypothesis, namely, that tillage management can modify the patterns of microbial enzyme activities’ distribution and their stoichiometry depending in the soil aggregates’ size and change the way in which aggregates contribute to soil functioning.

Tillage altered the microbial community structure under aggregate fractions

An increase in species diversity and abundances usually characterizes soil microbial community change due to soil organic matter’s accumulation and the quality of those components from plants (Chen et al. 2017; Jing et al. 2014; Yang et al. 2017; Zhang et al. 2016). Further, the composition of the soil microbial community was not only shaped by tillage but also affected by the aggregate size and the interaction of both factors. Our results agree with work by Zhang et al. (2016); they found that smaller aggregates had higher microbial biomass than larger aggregates. Differences in microbial biomass with declining aggregate size could be explained by the pore size of micro-aggregates being small (Chenu et al. 2001) and the microbial cell adherence being promoted by larger specific surface area of clay and silt particles (van Gestel et al. 1996). We find that soil microbial community structure varies as aggregate size changes, a result consistent with Jiang et al. (2018). Accordingly, how soil microbial communities are changed may be shaped by the differential chemical composition of aggregate fractions (Fanin et al. 2014).

The dynamics of microbial communities’ interaction with the soil substrate is a contentious topic. Evidently, the composition of bacterial community showed a pronounced pattern in response to tillage management (Fig. 6), because the soil substrate differed in both qualitatively and quantitatively, with high-intensity tillage practices increasing soil organic matter decomposition by disrupting the soil and aerating it (Chen et al. 2017; Zhang et al. 2016). Bacterial communities underwent a shifted in dominance, going from oligotrophic groups that are fast growing to opiotrophic groups that are slow-growing (Zhou et al. 2017), such as some Proteobacteria and Actinobacteria (r strategists), which those usually dominate communities at the oligotrophic conditions with low resource availability (Fierer et al. 2007). Such a shift could be driven by soils’ differential C and N substrates, this in line SOC and TN contents that were greater with conservation tillage than conventional tillage. In contrast, for communities harboring k strategists, namely Acidobacteria, those members become usually dominant as the soil which has greater substrates (Banning et al. 2011). A small number of Acidobacteria in all treatments suggests that restricting the input of plant debris to soil resulted to microbial community development limited by soil substrate. Interestingly, our study revealed that the abundance of bacterial community groups did not significantly differ between conventional (ST and FT) and conservation (NT and MT) tillage, suggesting that seven years of conservation tillage does not change bacterial community abundance and diversity (Table 1) in oasis farming systems. Other studies have reported that conservation tillage increased (Dong et al. 2017), decreased (Tyler 2019), or had no effect (Mackay et al. 2023) on microbial alpha diversities compared to conventional tillage, probably due to the different diversity method used (such as analyses of phospholipid fatty acids, according to which many taxa get grouped) (Mackay et al. 2023). However, we found distinct bacterial community compositions dominating different-sized aggregates in two tillage regimes, with MIA in the high-intensity tillage (ST and FT) differing from LMA and SMA. However, the bacterial community composition of LMA differed from SMA and MIA under conservation tillage. The difference in bacterial community composition between the two tillage systems (conservation and conventional) could be due to the different substrates available to microbes (Zhang et al. 2018), as affected by long-term disruptions of high-intensity tillage.

Our results found differences in fungal community composition with increasing tillage intensity (Fig. 6). In our study it was saprophytic fungi that were mostly collected, which are closely associated with organic matter decomposition and nutrients cycling through the secretion of hydrolase and oxidase, increasing soil nutrient availability (Duchicela et al. 2013). Structure changes to the microbial community of soil may correspond to the differential chemical composition of aggregate fractions driven by differing preferences for soil substrates among microbes (Fanin et al. 2014), consistent with SOC, TN, and TP contents being greater under conservation tillage than conventional tillage as aggregate size decreased. The fungal community may prefer to colonize those soil macro-aggregates characterized by pronounced porosity (Harris et al. 2003), and fungal hyphae are physically capable of binding smaller soil particles with plant residue or root exudates into aggregates of larger size (Duchicela et al. 2013; Rillig and Mummey 2006). However, in our study, the fungal community composition of aggregate fractions in the conventional tillage (FT and ST) differed from conservation tillage (MT and NT). Conventional tillage adversely affects the soil’s structure because the soil disruption lessens its overall porosity (Cavalcanti et al. 2020); breaks up soil macro-aggregates and the hyphae net; modifies microbial community composition, biomass and activity; and via soil substrate oxidation, leads to more soil organic C loss (Oliveira et al. 2017; Tenelli et al. 2019). Overall then, bacterial and fungal communities’ compositional changes in in response to soil aggregation points to distinctive successional patterns in microbial communities under long-term tillage management in this oasis farming area.

Crop growth is promoted by micro-aggregates

The dynamics of four key aspect of maize productively—grain yield, biomass of straw, root biomass, and the root: shoot ratio differed under different long-term tillage practices. Our experiment demonstrates that micro-aggregates hold more of soil nutrients in all four tillage treatments; hence, that aggregate type contributes to bulk soil the most available nutrients which in turn is consequential for crop properties. It is known that macro-aggregates can contain labile organic matter in greater proportions from plant litter or residue (Plante et al. 2002) and immobilized soil organic C and/or total N may enhance nutrient availability to crops following subsequent soil microbial turnover(Zhang et al. 2015). Our results disagree with those of Tian et al. (2022), who reported that macro-aggregates positively affected maize properties, such as the concentrations of C, P, and N in plants along with their ratios under organic fertilization. This discrepancy may be associated with the quantity and quality of soil substrates providing the microorganisms with nutrients. Likewise, the greater microbial biomass in micro-aggregates that enhance soil nutrient availability could also promote greater uptake of nutrients by plants. Alternatively, fungal hyphae residing in root in symbiosis could carry out functionally similar roles by absorbing nutrients and transferring them to host plants (Richardson et al. 2011).

All in all, applying conventional tillage regimes over the long term may impair the reduce stability of soil aggregates with adverse consequence for the formation of large macro-aggregates in addition to the nutrient pools contained in aggregates, as well as affecting the microbial communities of aggregates of all sizes (Fig. 8). Conversely, conservation tillage is able to promote soil macro-aggregate formation, nutrient distribution, and microbial community structure variation in different-sized aggregates, positively affecting plant growth. Our results suggest that conservation tillage management and micro-aggregates mostly affected the performance of above and below-ground maize biomass (Fig. 8). Studies have suggested the crop residues provided by no-till treatments in turn provide a sustained input of C into microbial communities that dwell in topsoil (Nunes et al. 2020), and that nutrients occurring in organic forms are crucial for ensuring soil functioning and productivity in the long term (Egan et al. 2018). Hence, third hypothesis of our study is supported, in that conservation tillage enhances sustainable soil functioning by changing the proportion of aggregate-associated nutrient pools and microbial community structure.

Conservation tillage regimes carried out over the long term can change the biomass of microbes, shift their community composition, modulate enzymatic activity in soil, and alter the soil aggregates’ chemical properties. No-tillage or reduced-tillage could benefit soil nutrient restoration, resource availability, microbial community restructure, and soil aggregate stability. Notably, for soil organic matter, not only its quantity but also quality important factors for restricting aggregate formation and changing microbial community composition. Therefore, we recommend the reasonable use of conservation tillage, such as no tillage, in oasis farmland ecosystems in semiarid regions where long-term, high-intensity tillage decreases soil fertility. The way in which the community composition of bacteria vis-à-vis fungi respond to different tillage management differs in the three aggregate-size fractions, likely due to soil substrate content and availability. Our results offer insights into the mechanisms of soil microorganism succession and the management of semiarid oasis farming areas.

Acknowledgments

This work was financially supported by the National Natural Sciences Foundation of China (NO. 41867013 and NO. 31960625), Gansu Province Sciences Foundation (NO. 20JR5RA074), and the Doctoral Scientiﬁc Research Funds (No. 19E056). We also thank the anonymous reviewers for providing critical comments and suggestions that improved the manuscript.

Author contributions

Chenglong Han: Methodology, investigation, writing—original draft. Weidi Zhou: Experimentation, investigation and data curation. Yanjie Gu: Methodology, Review. Junqiang Wang: Conceptualization, review and project administration. Yanfang Zhou: Investigation, Review. Yunyin Xue: Experimentation. Zhiguo Shi: Review, Supervision. Kadambot H.M. Siddique: Review and editing.

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Conservation tillage promotes crop productivity by changing soil aggregation in oasis farming ecosystems

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Abstract

Background Aims

Methods

Results

Conclusions

Figures

Introduction

Materials and methods

Results

Aggregate distribution

Enzyme activities and enzymatic stoichiometry

Microbial community biomass, diversity, and composition

Crop productivity under different tillage regime

Discussion

Crop growth is promoted by micro-aggregates

Conclusions

Declarations

References

Status:

Journal Publication

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