The isolate P-23 of P. nicotianae, isolated from a symptomatic pepper plant and previously evaluated for pathogenicity (Rodríguez-Molina et al., 2010), was used for the experiments.

Chlamydospores production followed the methodology described by Rodríguez-Molina et al. (2016), based on the culture of P. cinnamomi isolate in 150 ml of V8 juice broth in 20-cm Petri plates and incubation at 25°C in the dark for 10 days. Subsequently, the V8 juice broth was removed, replaced with distilled water and plates were incubated again in the dark at 18°C for 7 days. Then, the mycelium clumps were collected, washed with sterile distilled water, homogenized in 50 ml of sterile distilled water (homogenizer MICCRA D-1, ART-moderne Labortechnik, Germany) and sonicated (HD 2070, Sonoplus, Bandelin, Germany) on a 60-s cycle (active interval: 0.9 s, passive interval: 0.1 s). After centrifugation (2 min; 1,760 rpm), the resulting pellet was resuspended in distilled water. The concentration of chlamydospores was assessed with a Neubauer counting chamber and their viability was estimated by staining with Rose Bengal solution (Tsao, 1971).

For field and laboratory experiments, inoculum bags were prepared using field disinfected soil (autoclaved twice 1 h at 120°C) which subsequently was infested with chlamydospores of P. nicotianae, according to the procedure described by Rodríguez-Molina et al. (2016). The soil was placed inside the Agryl cloth bags which were closed with string. Bags with 5 g of soil and 500 chlamydospores g⁻¹ were prepared for the laboratory trials, and bags with 100 g of soil and 50 chlamydospores g⁻¹ for the field trial.

The trial was performed in a field with sandy loam soil (pH = 6.5; organic matter = 0.55%) at the Agricultural Research Institute Finca La Orden-Valdesequera (Extremadura, central-western Spain). The experimental design was a randomized complete block design with 4 replicates. The experimental plot size was 1.5 ×1.5 m, and each plot was artificially infested with 4 inoculum bags, with 100 g of soil and 50 chlamydospores per gram of dry soil each, which were buried at 20-cm depth, as used by Serrano-Pérez et al. (2017b).

The trial was conducted using commercial pellets of defatted B. carinata seed meal (BioFence®, organic N 6%, P 2.2%, K 2%, organic C 52%; Triumph Italia SPA). The concentration of sinigrin, the predominant glucosinolate, was 84.31 micromol g⁻¹ dry weight (Rodríguez-Molina et al., 2021) and the pellets also contained active myrosinase (Galletti et al., 2008). The pellets were added to the soil at 3 tons ha⁻¹, according to the manufacturer's recommendations, and incorporated 20-cm depth using a motorized tiller. The plots were covered with transparent polyethylene film (0.05 mm thick). Two control treatments were included: control non-amended (CP) and control non-amended and without plastic cover (C). The soil was irrigated with 50 mm of water at the beginning of the experiment and again after 14 days with the same dose to maintain it near saturation [volumetric water content (VWC) at 0.21 m³ m⁻³] in the first 24-cm depth. The treatment was carried out for 4 weeks. Plastic covers were removed after that time and inoculum bags were dug up for determining inoculum survival and infectivity. A composite soil sample was collected at 0 to 25-cm depth from each plot to analyze soil physicochemical properties and microbial activity.

As was performed in Serrano-Pérez et al. (2017b), soil temperature was continuously monitored at 20-cm depth, using soil probes and an automatic data logging system (HOBO Weather Logger, Pocasset, MA, USA) as well as moisture (volumetric water content, VWC) using 10HS sensors (ECH₂O, Decagon Devices Inc., Pullman, WA, USA), during the 4 weeks. Soil samples were collected before the beginning of the treatments and every 7 days during the trial to determine the evolution of soil pH. A pH electrode (HI12963, Hanna Instruments Inc., Woonsocket, RI, USA), in a 1:1 v/v slurry of soil and deionized water, was used for pH determination.

To evaluate the possible herbicidal effect of the biofumigation treatment, all vegetation grown during the treatments within the quadrat of each experimental plot was cut, weighed and dried in the oven (102°C) and the dry weight m⁻² was calculated. Total Fusarium density (CFU g⁻¹) in each plot was estimated on two occasions: before and after application of the treatments. Soil samples were air-dried, crushed and sieved (0.2 mm mesh size), and Fusarium density was evaluated by the soil-plate method (Warcup, 1950) using Komada's medium modified (Tello et al., 1991). Four replicates were prepared per soil sample.

Soil samples were randomly collected from each field plot at two times: (i) before treatments application (0 weeks) and (ii) at the end of the experiment (4 weeks). Before analysis, soils were air-dried and ground to pass through a 2-mm sieve. Organic matter (OM) was determined by dichromate oxidation (Walkley and Black, 1934). The electrical conductivity (EC) was measured in water at a soil:extractant ratio of 1:5. Ammonium (NH4+) content was determined after extraction with 2 M KCl (Mulvaney, 1996). Available phosphorus (P) was measured using the molybdate reactive method (Murphy and Riley, 1962) after bicarbonate extraction (Olsen et al., 1954).

Enzyme activities were determined on the same soil samples collected for physicochemical characterization. The methodology used for the determination of each of the enzymes analyzed can be found in Serrano-Pérez et al. (2017b). Briefly, the β-glucosidase activity was determined as the amount of p–nitrophenol (PNP) formed from p–nitrophenyl-β-D-Glucoside (PGN) (Eivazi and Tabatabai, 1988). Acid phosphatase activity was determined at pH 6.3, using 16mM p-nitrophenyl phosphate (EC 206.353.9) as substrate (Tabatabai and Bremner, 1969). For both enzyme activities, the concentration of PNP was determined photometrically at 400 nm. Dehydrogenase activity was determined by measuring the amount of triphenylformazan (TPF) released after incubating the soil with 2,3,5-triphenyl-tetrazolium chloride. TPF was extracted with methanol (Trevors et al., 1982) and determined by reading at 490 nm. Urease activity was assayed by the method modified by Nannipieri et al. (1980). The N-NH4+ was measured colorimetrically at 667 nm (Mulvaney, 1996).

The soil used in the laboratory experiment was taken from the field experiment plot at the Agricultural Research Institute Finca La Orden-Valdesequera. Before use, the soil was sieved (2 mm sieve).

The experimental set up to evaluate the effectiveness of different rates of pellets is practically the same as the one proposed by Serrano-Pérez et al. (2017b), in which closed controlled-temperature system was established to emulate the physical, chemical and microbial changes that take place in the field during the treatment. In this system, chlamydospores were in contact with all compounds released into the soil solution during the biofumigation process or only exposed to the volatile compounds generated during the treatments. The controlled system consisted of 1-liter airtight glass containers (14-cm height and 10-cm diam) filled with 600 g of soil mixed with the pellets and inoculated with a suspension of chlamydospores. The inoculum concentration was 50 chlamydospores g⁻¹ dry soil. In addition, small bags of soil (5 g) inoculated with 100 chlamydospores g-1 were prepared and hung in the headspace of the container, avoiding contact with the soil placed at the bottom of the container. Ninety ml of tap water was added to saturate the bottom soil and the containers were hermetically sealed for 4 weeks. They were placed in a programmable incubator with a complete randomized design with 4 replicates per treatment. The temperature regime in the incubator was: 17.5°C for 5 h/day; 22.5°C for 5 h/day; 27.5°C for 4 h/day; 32.5°C for 2 h/day; 27.5°C for 3 h/day; 22.5°C for 5 h/day. These temperatures were selected as they had been recorded during a spring solarization field trial in Extremadura (Rodríguez-Molina et al., 2016).

Three different pellets rates were assayed: 3 tons ha⁻¹ (BF3 = biofumigant commercial rate), 6 tons ha⁻¹ (BF6), and 20 tons ha⁻¹ (BF20). For the conversion between weight and surface area units, the bulk density of the soil (1.163 g cm⁻³) and a product incorporation depth of 20-cm were considered. Control non-amended and inoculated (CPhy+) and non-amended and non-inoculated (CPhy–) were included in the experiment. Besides, to verify the importance of soil microbiota in the process, a treatment with autoclaved soil (1 h at 120°C twice in 2 consecutive days) mixed with the pellets at the commercial field rate (3 tons ha⁻¹) was included (BF3-AS).

When finished the incubation period, the containers were opened, and the soil from the inoculum bags was analyzed to estimate the number of chlamydospores of P. nicotianae surviving after the volatile exposure as described by Serrano-Pérez et al. (2017b) as follows: “the 5 g of soil from the bags were added to 45 ml of 0.25% water-agar (1:10, v:v) and stirred for 2 min; five 1-ml aliquots from this slurry were spread evenly over each of five Petri plates containing 12 ml of NARPH medium (Romero et al., 2007) giving the detection threshold of 2 colony-forming units (CFU) g⁻¹ of soil. After 48 h of incubation in the dark at 25°C, the soil overlay was removed by gently washing the agar surface with tap water. Macroscopically visible colonies of P. nicotianae were counted and reported as CFU g⁻¹ dry soil.”

In the treated soil at the bottom of the containers, inoculum survival and infectivity were analyzed.

For the assessment of inoculum survival, 2 g soil samples (10 samples per container in laboratory experiments and 3 samples per inoculum bag in the field experiment) were analyzed for the presence of P. nicotianae. The 2 g soil samples were placed in 9-cm Petri plates and flooded with distilled water. Immature carnation petals floating on the water were used as baits for the detection of P. nicotianae (Tello et al., 1991). Results of survival are expressed as the percentage of soil samples in which P. nicotianae was detected.

The soil remaining in the containers (580 g per container) and in the bags (94 g per bag) after sampling for survival assessment was used for chlamydospores infectivity bioassays on pepper plants. The soil was put into 250-cm³ plastic pots (5 pots per container, with 116 g soil each, and 1 pot per inoculum bag). The soil in each pot was mixed with a previously disinfected (autoclaved for 1 h at 120°C) substrate of peat and vermiculite (1:3, v:v). The ratio soil:substrate (v:v) was 1:2 in pots from container and 1:2.5 in pots from bags. One pepper seedling (Capsicum annuum L. cv. Jaranda) at the 2 to 4-true-leaf stage was transplanted into each pot and the pots were placed in a growth chamber with a 16 h light at 28°C/8 h dark at 24°C cycle. Disease symptoms were assessed weekly for 2 months, when the bioassay was concluded. To confirm plant death by P. nicotianae, as the plants died, fragments of the crown and tap water-washed roots were plated on potato dextrose agar and NARPH medium. At the end of the bioassay, roots of all plants were inspected for the presence of disease symptoms and analyzed. Infectivity results are expressed as the percentage of diseased plants. As the blocks were kept separate, there were 4 replicates per treatment.

Radish (Raphanus sativus L.) and white mustard (Sinapis alba L.) seeds were used in these experiments as sensitive species to phytotoxic metabolites. The germination tests were conducted according to methods described previously (Zucconi et al., 1981; Carballo et al., 2009) with slight modifications. A pellet solution was prepared using sterile distilled water at 1:5 ratio (w:v), stirred vigorously for 30 min, and then centrifuged 1 min at 2,500 rpm. Dilutions from the solution were prepared using sterile distilled water to give the following concentrations: 100, 50, 25, 10, 5, and 0%. A sterile filter paper was placed inside each 55-mm diameter Petri plate, and 10 disinfected seeds were placed on the filter paper. One milliliter of each dilution was used to moisten the paper, and the plates were closed. Ten replicates per dilution and species were prepared. Seeds were incubated at 25°C in darkness, and germination and root length were recorded after 3 days. If the primary root was higher than 1 mm, seed germination was considered positive. The germination index (GI) was calculated for each dilution and species with the mean from 10 replicates according to the following formula: GI = G/G_(control) × L/L_(control), where G and G_(control) are the mean percentage of germination with B. carinata pellets and with the control, respectively, and L and L_(control) are the mean root lengths with B. carinata pellets and with the control, respectively.

Data of survival and infectivity of inoculum were analyzed by a Generalized Linear Model (GLM) with a binomial error distribution and logit link (binary dependent variable: detection/non-detection of P. nicotianae for Survival, and disease/non-disease for Infectivity). The treatment was included as fixed independent factor with three levels in the field experiment and five levels in the laboratory experiment and post-hoc comparisons of means were performed by Tukey's tests. The program R Core Team (2020) was used for these analyses.

Data of P. nicotianae chlamydospores survival in laboratory experiments, expressed as CFU g⁻¹ dry soil, were subjected to one-way ANOVA on log-transformed data [log(x+1)] followed by Tukey's multiple range test. Field pH was analyzed with two-way ANOVA for repeated measures (rmANOVA), including statistical significance for the effects of treatment (between-subjects factor) and sampling time (within-subjects factor), as well as the interactions between them. The degrees of freedom were corrected using Greenhouse-Geisser estimates of sphericity. Data on soil enzyme activity and soil properties were also subjected to two-way rmANOVA, studying the effects of treatments (between-subjects factor) and sampling time (within-subjects factor) and their interactions. Post-rmANOVA means comparisons were carried out with Bonferroni's correction. Data of Fusarium spp. population density was analyzed using Kruskal-Wallis non-parametric test. Phytotoxicity data were subjected to two-way ANOVA with dose and species as factors followed by Tukey's multiple range tests. Effects and differences were considered significant at P < 0.05. All analyses were performed with the software package SPSS version 20.0 (SPSS Inc., Chicago, Illinois, USA).

Soil temperatures at 20-cm in plastic-covered treatments (CP and BF3) fluctuated between 15 and 35°C, and a similar pattern in soil temperature was observed in these treatments. In covered plots, the greatest number of cumulative hours registered (~200 h) were between 20 and 25°C and only around 150 h above 25°C (Figure 1). Soil temperatures in the non-covered treatment fluctuated between 12 and 27.5°C, with the highest number of accumulated hours (217 h) around 17.5°C and only 15 h above 25°C.

Soil moisture in covered plots was maintained around field capacity (20 to 22 m³ m⁻³ of VWC) throughout the experiment, while in control without plastic cover soil moisture decreased, inconstantly due to occasional rains, to values of 12 m³ m⁻³ of VWC.

Soil pH did not significantly change in time, either in the control treatments (C and CP) or in BF3 treatment (Figure 2). Soil pH was kept between 6.5 and 7.2, and no differences were found in pH values among treatments throughout the experiment.

There was no noticeable effect of the biofumigation with B. carinata pellets on the survival of P. nicotianae in soil from recovered field inoculum bags. The estimated survival was very high for all treatments, and no statistical differences were found in any case (Chi² = 1.363; P = 0.506). The percentages of positive baits detected were 88 ± 14% (mean ± sd; n = 4) in CP and 94 ± 13% in BF3, achieving 100 ± 0% of positive baits when the plastic cover was not used (C). The use of a plant disease bioassay confirmed the pathogenicity of the surviving population of P. nicotianae. There was no significant effect of treatment on infectivity (Chi² = 8.6 e⁻⁶; P = 1.00), with trends very similar to that observed related to survival. Both covered treatments (CP and BF3) showed the same infectivity results (88 ± 14% of diseased pepper plants), while all plants showed root and crown rot disease in control without plastic cover (C).

Weed biomass was not affected by treatment [F₍₂₎ = 1.869, P = 0.210]. Biofumigated plots (BF3) had the highest level of weeds (31.5 ± 39 g of dry weight m⁻¹; n = 4), with Cyperus rotundus and Setellaria media being the predominant species, although these results were inconsistent due to differences between blocks and did not show significant differences compared with CP (23.8 ± 5.5 g of dry weight m⁻¹). Weed populations were lower in the non-covered control (C) (1.43 ± 1.9 g of dry weight m⁻¹), probably due to less favorable temperature and moisture conditions for seed germination in these plots.

Four Fusarium species were isolated from the soil before and after the treatments: F. oxysporum, F. solani, F. roseum and F. moniliforme. The total Fusarium spp. population density was over 3,982 ± 1,795 CFU g⁻¹ of soil (n = 48) in all plots at the end of the experiment. The Kruskal-Wallis test conducted did not show differences among treatments [H(2) = 1.69, P = 0.428].

Physicochemical soil properties before and after field experiment are presented in Table 1. Before pellet incorporation (0 weeks), no significant differences were found between plots for any of the soil properties studied, indicating the homogeneity of the plots before the setting up of the trial. At the end of the field experiment, there were significant increases in EC. No significant increase was observed in OM in BF3 treatment. The increases in NH4+ were significant in all cases, but no differences were found between treatments at the end of the experiment. The increases in available P were significant only in BF3, although there were no differences among treatments at the end of the experiment.

Soil enzymatic activities before and after field experiment are presented in Table 2. Just as with the physicochemical properties, there were no differences among plots at the beginning of the experiments. The treatment with pellets (BF3) did not enhance the soil enzymatic activities, and the results showed no significant differences compared with the controls (C and CP) for any enzyme studied.

Volatile compounds released into the headspace of containers from the biofumigant rate BF3 did not suppress the chlamydospore germination of P. nicotianae (Table 3). Similarly, doubling the recommended commercial rate (BF6) was also ineffective on chlamydospore inactivation. However, when the pellets rate was increasing (BF20) or the rate BF3 was added into autoclaved soil (BF3-AS) no germination of chlamydospores was observed in any plate.

All rates of pellets assayed decreased the infectivity of the pathogen with respect to the control without pellets and the effect of treatment was significant on infectivity (Chi² = 10.17; P = 0.037). No significant differences were detected between BF3 and BF6 biofumigant rates and the control treatment (Figure 3). This lack of significance may be due to low infectivity reported in the control treatment (25% of diseased plants). Marginally significant differences (P = 0.056) were detected between the BF20 and BF3-AS treatments and the control treatment. The effect of the B. carinata pellets was more noticeable on the survival of P. nicotianae inoculum than on infectivity on pepper (Chi² = 66.23; P < 0.001). The doubled commercial rate (BF6) reduced the inoculum survival significantly, but no differences were found between the commercial rate BF3 and the control without pellets. On the contrary, when the rate BF3 was added into the autoclaved soil (BF3-AS) or the rate was increased (BF20), the infectivity on pepper and the inoculum survival were drastically reduced (Figure 3).

The germination indices (GI) of radish and mustard with increasing B. carinata pellet dilutions found in the plate experiment are shown in Figure 4. The ANOVA indicated that the dilution had a significant effect [F₍₄₎ = 69.46; P < 0.001], while the interaction between the two factors (dilution × species) was not significant [F₍₄₎ = 1.697; P = 0.158). The GI was gradually decreasing for rising concentrations higher than 10%, and no germination was observed either with 100% concentration in radish seeds or with 50 and 100% concentrations in mustard. Mustard seeds were significantly more sensitive [F₍₁₎ = 13.358; P < 0.001] than radish seeds, with lower GI for all dilutions assayed.