2.1. Characterization of the Study Area
This is a case study of two commercial cotton fields (Gossypium hirsutum L.) cultivated in Brazil: cultivar TMG 47B2RF was tested in Sapezal (SPZ, 12°59′22″ S; 58°45′52″ W 560 m asl) in Mato Grosso State (MT) during the 2021 season, and cultivar FM 974GLT was tested in Riachão das Neves (RN, 11°31′49″ S; 45°43′57″ W 795 m asl) in Bahia State (BA) in the 2021/2022 season. Sowing was carried out on 01/23/2021 in SPZ (rows spaced at 0.9 m) and on 12/13/2021 in RN (rows spaced at 0.76 m), using 10 seeds per meter, and emergence occurred 5 days later.
The field in SPZ was 20 hectares and was cultivated with soybean (1st crop) and cotton (2nd crop) over the last 10 seasons and represents a growing area of 266,000 ha (14% of Brazilian cotton area). In RN, the field size was 90 hectares, including soybean (2020/2021) and a mixture of cover crops (pearl millet, turnip, and
Urochloa ruziziensis) after soybean harvest preceded cotton cultivation. Riachão das Neves had a cotton cultivation area of 33,500 hectares, which represents 9% of Bahia State cotton area and 2% of Brazilian cotton area. The soil is characterized as oxisol in both areas (Soil Survey Staff) with a clayey texture in SPZ and sandy loam texture in RN. The regional climate, according to Köppen and Geiger, is Aw (tropical savanna climate with dry winter). Meteorological parameters such as rainfall, maximum and minimum temperatures, and solar radiation (
Figure 2) were recorded in a weather station 1 km from the experiment. Degree day ((DD) = ((T + t)/2) − 15) was used to calculate the effective daily temperature, and vapor pressure deficit (VPD) was calculated using average air temperature and air humidity (
Figure 3).
2.3. Data Collection, Evaluation and Analysis
At harvest, soil samples were collected for chemical (0–60 cm), physical (0–40 cm in SPZ and 0–60 cm in RN), and soil enzyme activity (0–20 cm). The chemical analysis was carried out using the methodologies described by Raij et al. [
7]. The particle size was analyzed [
8]. Soil pH was measured in a 0.01 mol/L
−1 CaCl
2 suspension, and soil organic carbon (OC) was determined using potassium dichromate (K
2Cr
2O
7). Available phosphorus (P) was determined colorimetrically using a spectrophotometer and exchangeable basic cations calcium (Ca), magnesium (Mg), and potassium (K) were extracted using an ion-exchange resin by atomic absorption spectrometry (PerkinElmer analyst 200, Shelton, CT, USA). Exchangeable aluminum (Al
3+) was extracted using 1 mol/L
−1 KCl in a 1:10 soil-to-solution ratio. The potential acidity (H + Al) was measured immediately after measuring soil pH by adding 5 mL of SMP buffer solution (pH = 7.0) to the suspensions. Sulfur (S) determination was performed at a 0.01 mol/L
−1 Ca(H
2PO
4)
2 solution, and available micronutrients iron (Fe), copper (Cu), manganese (Mn), zinc (Zn), and boron (B) were extracted with DTPA and determined by inductively coupled plasma optical emission spectroscopy (ICP-OES).
For soil physical properties, undisturbed soil samples were collected from the midpoint of 0–20 and 20–40 cm depth intervals in SPZ and 0–20, 20–40, and 40–60 cm depth intervals in RN. Small trenches were dug, and undisturbed samples were collected using stainless steel cylinders. The soil samples were wrapped with aluminum foil, carefully transported to the laboratory, and maintained at 5 °C to reduce biological activity [
9]. Subsequently, the samples were placed in trays filled with distilled water and left until they reached saturation. The samples were then weighed and placed on a tension table [
10] at matric potential (ψ) of −10 kPa. Upon reaching hydraulic equilibrium, the samples were weighed again and dried in an oven at 105 °C for 48 h. Soil bulk density (ρ
b) was then calculated as the ratio between oven-dried soil mass and sample volume [
11]. Total porosity (TP) of soil was obtained as the saturated soil water volume [
12]. Field capacity (FC) was equated with the soil water content retained at ψ = −10 kPa, and the air-filled porosity (ε
a) was computed by the difference between TP and FC [
9]. Other soil physical indicators were obtained as per Reynolds et al. [
13], including (1) the storage capacity of soil water (dimensionless), that is, the ratio between the soil water content at ψ = −10 kPa (FC) and TP, and (2) the storage capacity of soil air (dimensionless), that is, the ratio between air-filled porosity at ψ = −10 kPa and TP.
After equilibration at ψ = −10 kPa, air permeability (
Kair) was measured using a constant head permeameter [
14]. A mass flow controller was used to set the airflow values that flowed into each sample, in addition to a differential pressure manometer to identify the pressure differences, modified according to the structural soil conditions. Kair (µm
2) was calculated using Equation (1).
where Q is the mass flow (m
3/s); η is the air viscosity at 20 °C (1.84 × 10
−5 N s/m
2); As is the area perpendicular to the air movement (m
2); z is the soil column height (m); and P is the differential air pressure (Pa).
The soil pore organization index,
k1 (μm
2 m
3 m
−3), was calculated according to Groenevelt et al. [
15]:
The degree of compactness (DC) in both fields was estimated using Equation (3).
The soil bulk density reference (ρ
ref) was calculated using the pedotransfer function [
16].
Soil enzyme activity was determined by sampling soil at 0–10 cm and 10–20 cm soil depths. Acid phosphatase was determined by the method described by Tabatabai and Bremner [
17]. We added 4 mL of MUB pH 6.5, and 1 mL of PNF was diluted in MUB pH 6.5 in 1 g of soil. All samples were placed in the bath for 1 h at 37 °C; after this, 1 mL of CaCl
2 0.5 mol/L
−1 and 4 mL of NaOH 0.5 M were added. The samples were then shaken and filtered. The samples were recorded at 400 nm absorbance using a spectrophotometer. Arylsulfatase enzyme activity was measured [
17] after weighing 1 g of soil and adding 4 mL of acetate–acetic acid buffer 0.5 mol/L
−1 (pH 5.8) and 1 mL of solution p-nitrophenyl sulfate (potassium sulfate 0.025 M) PNS. The samples were shaken and incubated for one hour under a constant temperature of 37 °C. After incubation, 1 mL of CaCl
2 0.5 M and 4 mL of THAM (pH = 12) were added to the samples. The samples were filtered and recorded at 400 nm absorbance. β-glucosidase enzyme activity was measured [
18]. A total of 1 gram of soil was homogenized in 4 mL of MUB buffer (pH = 6.0) and 1 mL of PNG (p-Nitrophenol 1000 ppm). After incubating for one hour, 1 mL of 0.5 mol/L
−1 CaCl
2 and 4 mL of 0.1 M THAM (pH = 12) were added to the samples, which were then shaken and filtered for readings at 400 nm absorbance.
Plant mapping was performed in five plants per replication by counting and weighing each boll and indicating node and fruit position (P1, P2, and P3+) right before mechanical harvest. Boll retention in each fruit position (P1, P2, and P3+) and the total boll setting (considering all positions) were calculated by dividing the number of bolls retained by the total fruiting sites issued by the plant. Accumulated yield was calculated to determine the contribution of each node to yield using the data from plant mapping and consisted of the boll weight multiplied by the number of bolls in each node multiplied by plant density. The production of vegetative branches (monopodial) was summed to the following fruiting branch and so on until the last node at the top of the plant.
At harvest, 190 DAE in SPZ and 180 DAE in RN, yield was determined by a mechanical machine (John Deere CP 690, Moline, IL, USA) in all fields. Yield components (boll weight and boll number) were determined in 2 m of a row (1.8 m
−2 in SPZ and 1.52 m
−2 in RN) with 20 replications by counting, handpicking, and dividing the total boll weight by the boll number to obtain mean boll weight (
Figure 4). A sample of 150 g of seed cotton was used for fiber quality determination (High Volume Instruments, Uster Technologies, Uster, Switzerland). The gin turnout used to calculate lint yield was from the commercial field.