1. Introduction
Straw return to the field is a sustainable alternative to chemical
fertilisers for soil enrichment and improving crop growth because of its
renewability and carbon sequestration benefits (Li et al., 2022a).
Excessive use of chemical fertilisers leads to nitrate leaching and
residual nutrients in the soil (Patle et al., 2019). Additionally, straw
burning generates substantial amounts of fine particulate matter,
affecting human health and safety (Huang et al., 2021). Returning straw
to the field instead of using partial fertilisers can reduce the cost of
agricultural production and air pollution caused by straw burning,
improving soil quality. Chen et al. (2022) reported that straw
application enhances soil carbon, nitrogen, and phosphorus cycling,
improving soil physicochemical and biological properties. Straw
application significantly increases the levels of stable and active
carbon and nitrogen pools in the soil, increasing wheat yield (Hu et
al., 2021; Cui et al., 2022). Furthermore, straw improves the metabolic
function and diversity of microbial communities (Liu et al., 2020; Jin
et al., 2020). However, the effect of straw application on soil
nutrients and microorganisms depend on the amount of straw incorporated.
Using a meta-analysis, Wang et al. (2021a) found that soil organic
carbon considerably increased with increases in the quantity of straw
incorporated. Moreover, Zhao et al. (2016) reported that incorporating a
low amount of straw had marginal effects on soil microorganisms;
however, incorporating a high amount increased total phosphorus fatty
acids, resulting in changes in the microbial community structure,
suggesting that incorporating optimal amounts of straw in the field can
increase soil nutrients and alter the soil microbial diversity.
Soil carbon and nitrogen influence terrestrial ecosystems and global
biogeochemical cycles (Han et al., 2020; Lewis et al., 2021). Studies on
soil carbon and nitrogen contents have received attention from
researchers owing to the rapid shifts in global climate and ecology. The
cycling and accumulation of soil carbon and nitrogen are crucial for
assessing soil nutrient content (Xiao et al., 2023). Soil carbon and
nitrogen fractions are classified into stable and active pools. Changes
in these indicators reflect soil quality and health, particularly in the
context of active carbon and nitrogen pools. Active carbon and nitrogen
pools are formed owing to the accumulation of carbon and nitrogen in the
soil because organic matter is involved in biological and chemical
processes. Although they constitute a relatively small proportion of
soil organic carbon and nitrogen, active carbon and nitrogen pools are
key indicators of soil quality (Ren et al., 2023a; Wang et al., 2022a;
Benbi et al., 2012). Specifically, soil active carbon is a marker of the
soil carbon cycle, carbon sequestration potential, and soil response to
climate change (Wu et al., 2020a; Benbi et al., 2012). Furthermore, soil
active nitrogen content is used to determine nitrogen transformation,
nitrification, reduction, and fixation processes in the soil (Koković et
al., 2021). Soil carbon and nitrogen is derived primarily from
underground root exudation and decomposition of aboveground plant
residues (Zhu et al., 2021). Soil microorganisms play a crucial role in
element cycling by decomposing plant residues through metabolic
activities, converting plant organic matter into soil organic matter.
Microorganisms increase oil-stable carbon and nitrogen pools by
decomposing organic matter in the rhizosphere soil (Adamczyk et al.,
2019). The abundance and activity of microbial species influence the
changes in the carbon and nitrogen fractions, regulating the stability
and efficacy of soil carbon and nitrogen cycles (Wang et al., 2023a;
Coonan et al., 2020). Straw application alters the soil microbial
community by providing cellulose and lignin (Yan et al., 2019). Guan et
al. (2023) reported that straw application stimulated the growth of soil
microorganisms, affecting the structure and function of the microbial
community. Wu et al. (2020b) found that straw application combined with
the application of inorganic fertilisers considerably altered soil
bacterial communities, hypothesised to be correlated with soil carbon
and nitrogen contents, suggesting that straw application stimulates soil
microorganisms. However, whether the straw application changes and
optimises the soil nutrient fraction is unclear.
Therefore, the present study carried out a 5-year straw application
experiment to determine its effect on the contents of soil carbon and
nitrogen pool components, soil bacterial and fungal community diversity,
and the capacity of soil microorganisms to metabolise carbon.
Furthermore, the study aimed to determine the effects of incorporating
different amounts of straw on soil carbon and nitrogen fractions and
microbial structure and function. The study analyses the effects of soil
microorganisms on carbon and nitrogen fractions under straw application.
The results of this study provide a theoretical basis for the
improvement of soil nutrient content and the soil ecological environment
in dryland wheat production areas.
2. Materials and methods
2.1. Study
area
The experiment was conducted from 2018 to 2023 in Hongtong County,
Linfen City, Shanxi Province, China (36°15’34” N, 111°40’31” E). The
study site is in a typical Loess Plateau dryland farming area and has a
temperate continental monsoon climate with an average annual temperature
of 12.6 ℃ and mean annual precipitation of 500 mm concentrated from
June–September. The study area has calcareous brown soils with a
physical clayiness of approximately 40%. The results of the
physicochemical properties of the soil at the study site are presented
in Table S1.
2.2. Experimental design and sample
collection
The field experiment was laid in a randomised complete block design with
four treatments: S0 (0% straw returned to the field), S1/2 (50% straw
returned to the field), S1 (100% straw returned to the field), and S2
(200% straw returned to the field). The experiment had four blocks.
Each plot measured 126 m2 (30 m ×4.2 m [length ×
width]). The experiment was set up in 2018. Wheat was harvested, and
straw was returned to the field in June each year for the 5-year study
period. Wheat was sown in mid-October, and fertiliser was spread in late
October each year for the 5-year study period. We crushed the wheat
straw into 2 cm pieces and incorporated it into the field by tilling
using a tiller at 0–20 cm soil layer depth. The amount of straw
returned to the field was calculated considering the straw yield of the
previous harvest; therefore, the amount of straw returned to the field
differed in each treatment in each recurrent year for the 5-year study
period. Wheat was sown in furrows on a ridge and film. After fertiliser
application and land preparation, the ridge was covered with a film, and
the seeds were sown on the side of the film in the furrow. The wheat
variety used was ”Jinmai 47”, planted at 150 kg/hm2sowing rate without irrigation during the experiment.
The fertiliser application rate for each treatment was determined before
the sowing of winter wheat by assessing the nitrate nitrogen in the 0–2
m soil layer and the available phosphorous and potassium in the 0–40 cm
soil layer. The calculated fertiliser application rate was adjusted by
subtracting the amount of nutrients contributed by the straw application
obtain the final fertiliser application rate. Briefly, the nutrient
content of the straw was measured after harvesting the wheat in each
season. The nutrient release rates for straw nitrogen, phosphorus, and
potassium were 50 %, 65 %, and 90 %, respectively, and the amount of
nutrients provided by straw application was determined. Soil samples
were collected in June 2023 from the 0–20 cm soil layer. The amount of
fertilisers applied in each year of the 5-year experimental period are
presented in Table S2. The soil at the experimental site had sufficient
potassium. Therefore, only nitrogen and phosphorus fertilisers were
applied. Urea (containing 46 % N) was applied as the nitrogen source,
and calcium superphosphate (containing 16 %
P2O5) was applied as the phosphorus
source. Same fertilisers were applied throughout the experimental
period. After harvesting the winter wheat in June 2023, four soil
samples were randomly collected from each plot from a depth of 0—20
cm. Furthermore, soil samples were collected for subsequent
microbiological analyses as described above. Samples for DNA
high-throughput sequencing and Biolog-ECO analysis were stored in a
refrigerator at -80 ℃, and the remaining samples were stored in a
refrigerator at -4 ℃.
2.3. Sample determination and
methods
(1) Analysis of soil carbon and nitrogen indicators
Soil organic carbon content was determined using the potassium
dichromate external heating method (Bao, 2000). Soil total nitrogen (TN)
content was determined using the Kjeldahl method. Soil microbial biomass
carbon (MBC) and nitrogen (MBN) contents were determined using the
K2SO4-chloroform fumigation method (Wu
et al., 2006) and a total organic carbon (TOC) metre (Thermo Fisher
Scientific, Waltham, MA, USA). Soil dissolved organic carbon (DOC) and
nitrogen (DON) were extracted using 1 mol/L KCl and filtered through a
0.45-μm membrane filter (Thermo Fisher Scientific). DOC and DON were
determined using a TOC metre (Thermo Fisher Scientific). For the
determination of soil light fraction organic carbon (LFOC), heavy
fraction organic carbon (HFOC), light fraction organic nitrogen (LFON),
and heavy fraction organic nitrogen (HFON), recombinant soil samples
were obtained by shaking and centrifuging 1.8 g/cm3 of
zinc bromide solution with the soil samples in a centrifuge tube thrice,
shaking and centrifuging the soil samples with 95% ethanol thrice to
wash out the excess zinc bromide, and finally shaking and centrifuging
the soil samples with distilled water twice to wash out the excess
ethanol. Subsequently, HFOC, HFON, and LFOC and LFON were determined by
the potassium dichromate heating, Kjeldahl, and the difference
subtraction methods.
(2) DNA extraction and sequencing
DNA from the soil samples was extracted using the E.Z.N.A.® soil DNA kit
(Omega Bio-Tek, Norcross, GA, USA). DNA concentration and purity were
determined using a NanoDrop2000 spectrophotometer (Thermo Fisher
Scientific). Polymerase chain reaction (PCR) of the V3–V4 variable
region of the 16 S rRNA gene was performed using the primers 338F
(5’-ACTCCTACGGGGAGGCAGCAG-3’) and 806R (5’-GGACTACHVGGGTWTCTAAT-3’). The
V4 variable region of the 16S rRNA gene was amplified using PCR. The PCR
products were recovered on a 2% agarose gel (Agarose. Konya, Turkey)
and purified using a DNA Gel Recovery and Purification Kit (PCR Clean-Up
Kit, Omega Bio-tek, Shengzhen, China). The recovered products were
quantified using Qubit 4.0 (Thermo Fisher Scientific) at the Shanghai
Majorbio Bio-Pharm Technology Co., Ltd., Shanghai, China.
(3) Determination of soil microbial metabolism indicators
Soil microbial carbon metabolism capacity was assessed using a
Biolog-ECO microplate (Thermo Fisher Scientific). Briefly, 10.00 g of
the soil sample was added into a conical flask containing 90 mL of
0.85% sodium chloride solution and shaken for 1 min followed by a 1-min
ice bath, and the procedure was repeated thrice. Next, the mixture was
allowed to settle, and a 1000-fold dilution of the soil suspension was
prepared, and 150 µL of the soil suspension was extracted and inoculated
onto the Biolog-ECO microplate (Thermo Fisher Scientific). Finally, the
microplate was placed in an incubator for continuous cultivation at 30 ℃
for 144 h.
Optical density was measured at 0, 6, 12, 24, 48, 72, 96, 120, and 144 h
at the wavelength of 595 nm using an ELX 808 microplate reader system
(Thermo Fisher Scientific). Subsequently, Average well color
development (AWCD) was calculated, and the Shannon (H ), Simpson
(D ), and Pielou indices (E ) were calculated to
characterise the functional diversity of the soil microbial community
using the following formulas:
AWCD = Σ (Ci-R) / 31
(1)