Experimental set-up
The current study included three cultivation cycles carried out in a glasshouse at China Agricultural University (40°N, 116.3°E), Beijing, China. The 1st cultivation was from December 2017 to March 2018, and the 3rd one from December 2018 to February 2019. The glasshouse temperature in those periods was maintained at 21–25°C during the day and 15–18°C at night, with a photoperiod of 10–12 h throughout the growth period. The 2nd cultivation was from April to June 2018, and the temperature was maintained at 24–28°C during the day and 18–20°C at night, with a photoperiod of 12–14 h over the growing season.
Growth and harvest of the first cultivation
We used the soil collected from 0–20 cm top soil at the Quzhou Experimental Station (36.9°N, 115.2°E; 39.6 m a.s.l.), Hebei Province, China. It is calcareous alluvial silt loam with the following properties: Olsen-P 5.6 mg kg-1, total C 18.2 g kg-1, total N 1.08 g kg-1, total P 0.7 g kg-1, NH4OAc-potassium (K) 32.3 mg kg-1 and pH 8.3 (2.5:1 water/soil), all of which were determined before growing plants. The cultivation included four treatments (no-plants, sole maize, sole faba bean and maize/faba bean intercropping), and each treatment had 10 replicates. Each pot contained 2.0 kg of the air-dried soil that was beforehand passed through a 2-mm sieve. To ensure sufficient nutrient supply, plants were also fertilized with basal nutrients at the following rates (unit: mg kg-1 soil): 200 N as Ca(NO3)2·4H2O, 100 K as K2SO4, 50 Mg as MgSO4·7H2O, 2.2 Mn as MnSO4·H2O, 2.3 Zn as ZnSO4·7H2O, 0.51 Cu as CuSO4·5H2O, 0.12 B as H3BO3, 0.02 Mo as (NH4)6Mo7O24, and 0.88 Fe as EDTAFe-Na. No P was applied.
The seeds of maize (Zea mays L. cv ZD958) and faba bean (Vicia faba L. cv Lincan5) were first surface-sterilized in 30% v/v H2O2 for 10 min, and then rinsed with deionized water. They were soaked in a saturated CaSO4 solution for 12 h and then grown for 48 h in Petri dishes covered with wet filter papers. After emergence of the radicle, two maize seedlings or four faba bean seedlings were kept in each pot for the corresponding sole cropping, which was consistent with the density in a field study (Liao et al. 2020). To keep plant density consistent between cropping systems, one maize seedling and two faba bean seedlings were grown in each pot for intercropping. In addition, the study included a no-plant treatment, in which no plant was grown but the soil was supplemented with the same amounts of water and nutrients as other treatments.
All pots were arranged in a completely randomized design, and re-randomized weekly during the growing period. The plants were watered every day to maintain a soil field capacity of 80%. The cultivation lasted for 91 days, and then we harvested shoots and roots separately. Shoots and roots were used for measuring biomass and P concentration. The soils of 10 replications of each treatment were mixed, homogenized and stored at 4°C until the start of the cultivation 2.
Growth and harvest of the second and third cultivations
In the 2nd cultivation, we investigated the influence of root residues from cultivation 1 on plant growth. There were in total four soil treatments in cultivation 1, and each soil treatment was randomly divided into two even parts. One part was used for the treatments of no return of root residues, and another for the treatments of return of root residues, where the soil was a mixture with corresponding pre-crop root residues. Regarding the treatment of return of root residues, root samples of pre-crops were first oven-dried at 45°C for four days to minimize alteration of plant chemistry, and then cut with scissors into pieces of less than 2 mm before incorporating them into the soil. The cropping systems were the same as in cultivation 1, and thus there were seven treatments in total, including one no-plant treatment and six treatments comprising three cropping systems (sole maize, sole faba bean and maize/faba bean intercropping) without or with corresponding pre-crop root residues. Each treatment had five replicates. The same cropping system was grown in conspecific soil collected from the 1st cultivation. The root residues were added at a rate of 5.1 g pot-1, containing a P amount ranging from 3.6 to 4.8 mg pot-1, and incorporated into the topsoil (0–10 cm). Detailed information of the root residues is presented in Table 1. In addition to root residues, each treatment received the same amount of nutrients as those in the 1st cultivation. Plants were grown for 56 days and shoots and roots were harvested separately. Biomass and P content of the shoot samples were determined, and root samples were treated as in the 1st cultivation, stored at 4°C until used for the 3rd cultivation. Soils of all treatments were collected separately and then stored at 4°C to be used for the 3rd cultivation.
The 3rd cultivation had the same experimental setup as the 2nd, but the amount and the nature of the root residues were different from those in the previous run. The root residues used in this cultivation had a total P content ranging from 2.0 to 3.8 mg pot-1, and a C/P ratio from 257 to 533 (Table 1). Plants were grown for 75 days, and then we sampled shoots, roots and soil, separately, for different measurements as mentioned below.
The soil samples for chemical and Hedley P-fraction measurement
At the harvest of the 3rd cultivation, plants were separated into shoots and roots. Roots were removed carefully from the soil, and shaken gently to remove loosely adhering soil. The soil adhering to roots was defined as rhizosheath soil (Kamh et al. 1999). Then roots were transferred to a tube containing 50 mL 0.2 mΜ CaCl2 and gently shaken to dislodge the rhizosheath soil, followed by shaking for 5–10 s to create a homogeneous suspension (Veneklaas et al. 2003). We also sampled bulk soil from the no-plant control and placed it in 50 mL 0.2 mΜ CaCl2. A subsample of the extract was filtered into a 1 mL high performance liquid chromatography (HPLC) vial through a 0.22 μm syringe filters for carboxylate analysis in the rhizosphere and bulk soil (Shen et al. 2003). In order to determine the activity of acid phosphatase in the rhizosheath and bulk soil, a 0.5 mL suspension was placed in a 2 mL centrifuge tube with 0.4 mL sodium acetate buffer and 0.1 mL p-nitrophenyl phosphate (pNPP) substrate added, and then incubated at 30°C for 60 min; the reaction was terminated by adding 0.5 mL 0.5 M NaOH (Alvey et al. 2001). The P concentration was measured using a spectrophotometer (UV757T, Shanghai Instrument Co. Ltd., Shanghai, China) at a wavelength of 405 nm. The amount of rhizosheath soil collected differed among treatments. In order to eliminate effects of a different water/soil ratio on pH determination of rhizosheath extracts, a modified pH (water/soil ratio was adjusted to 2.5:1) was calculated from the measured pH by an equation according to Li et al. (2010). The delta pH was defined as the pH difference between rhizosheath and bulk soil, and this calculation was also applied to delta organic acid and delta acid phosphatase activity.
At the harvest of the 3rd cultivation, the bulk soil of each treatment was collected, and then Microbial-P, sequential Hedley P fractions, total N and organic C were measured. The air-dried soil samples were used to analyze sequential P fractions as described by Hedley et al. (1982) and modified by Tiessen and Moir (1993). Different extractors were added to 0.5 g of soil in the following sequential order: anion-exchange resin (referred to Resin-P) and 0.5 M NaHCO3 (referred to NaHCO3-Pi and NaHCO3-Po), denoting labile inorganic and organic P; 0.1 M NaOH (referred to NaOH-Pi and NaOH-Po) and 1.0 M HCl (referred to 1 M HCl-Pi), denoting moderately labile inorganic and organic P; and concentrated HCl (referred to conc. HCl-Pi and conc. HCl-Po) and concentrated H2SO4–H2O2 (referred to Residual-P), denoting non-labile inorganic and organic P. After adding each extractor, we repeated the following steps. The suspension was first stirred for 16 h in a shaker (200 rpm), then centrifuged for 10 min at 25,000×g at 0°C; after passing through a 0.45-μm membrane filter, the supernatant was stored prior to colorimetric analysis. Inorganic P was determined according to the method of Murphy and Riley (1962). The total P concentration in the different extracts (NaHCO3-P, NaOH-P and conc. HCl-P) was determined by digestion of each extract with ammonium persulfate. Organic P concentration was the difference between total P and inorganic P concentration. The accuracy of the sequential P extraction was assessed by comparing the sum of all P fractions with the concentration of the total P concentration determined with the reported approach (Ai et al. 2017). The sum of the P fractions was, on average, 110% of the measured total P concentration. Phytate (phytic acid sodium salt hydrate, Sigma, America) was used as standard organic P to carry out the recovery test, which was 56% in this case. The detailed information about the procedures of soil P extraction and P pool grouping were provided in the previous study (Liao et al. 2020).
Soil microbial biomass P (referred to as Microbial-P) was estimated by using the fumigation extraction method (Brookes et al. 1982). The associated processes are as follows: fresh soil was first adjusted to a soil moisture content ranging from 10–15%, and then three individual portions of 2.88 g fresh soil (about 2.5 g dry soil) were weighed and added to jars separately. One portion was fumigated with chloroform, one portion was left unfumigated, and another was spiked with 0.25 mL 250 mg P as KH2PO4 L-1 to assess P recovery. Three soil portions were incubated in a vacuum desiccator for 24 h, followed by extracting with 50 mL 0.5 M NaHCO3, respectively. The P concentration was measured using the molybdovanadophosphate method (Olsen et al. 1954). Microbial biomass P was calculated as chloroform-released Pi concentration by dividing by 0.4, i.e. assuming that 40% of the P in the biomass is rendered extractable as Pi by chloroform. Chloroform-released Pi concentration was the concentration difference of Pi between fumigated and non-fumigated soils. Microbial biomass P was corrected by using recovery of added phosphate.
Plant samples for chemical analyses
All shoot and root samples collected across three cultivations were first oven-dried for 72 h at 70°C and then weighed. After grinding, the samples were used to determine P concentrations as detailed above (Johnson and Ulrich 1959). Carbon (C) and nitrogen (N) concentration of plant and soil samples were determined with an Elemental Analyzer (vario MACRO CUBE, Hanau, Germany).
Calculations and statistical analyses
We compared the observed and expected values of shoot biomass, root biomass and total P content to assess the intercropping advantage (Loreau and Hector 2001). The observed value was the real sum of intercropped maize and faba bean, while the expected value was a weighted sum of sole maize and sole faba bean based on each crop’s relative density in intercropping (the relative density of each crop was 0.5 in this study).
Differences between treatments in biomass and P concentration of shoot and root, total P uptake, delta soil pH, delta soil carboxylates and delta soil phosphatase activity, were statistically tested using three-way analysis of variance (ANOVA), with crops (maize vs faba bean), cropping systems (sole cropping vs intercropping) and root residue treatments (removed vs added) as the treatment effects. Soil P fractions were analyzed for significant differences using two-way ANOVA, with cropping systems (no-plant, sole maize, sole faba bean and maize/faba bean intercropping) and root residue treatments (removed vs added) as the treatment effects. One-way ANOVA was used to test significant differences between observed and expected values. All ANOVAs were conducted using the SAS software package (SAS v.8.0).