Control of Soilborne Fungi with Biofumigation

The experiment was conducted at the Clemson University Coastal Research and Education Center, Charleston, SC. The soil was Yonges loamy sand with a pH of 6.3 in 2004 and 6.1 in 2005. Previously, the field was planted with muskmelon (Cucumis melo L.) and watermelon (Citrullus lanatus) in the spring of 2003. To increase the inoculum density of F. oxysporum f. sp. niveum, watermelon cv. Black Diamond was direct-seeded 15 September 2003 and 11 and 25 August 2004 in all treatment plots. At the time of seeding, 280 kg/ha ammonium sulfate 21-0-0 and 560 kg/ha of 10-10-10 N-P-K was applied in 2003 and 2004, respectively. In 2003, Black Diamond plants were mowed and the field was disked twice on 19 November. After seeding in 2004, emergence was poor due to excessive rainfall from Hurricane Charley, Tropical Depression Bonnie, and Tropical Storm Gaston. The field was not replanted and it was disked twice.
The experimental design was a randomized complete block with six replications. Treatment plots measured 24.4 by 0.9 m with a 3-m fallow space between plots. Six treatments were evaluated: 1) non-amended control soil covered with black polyethylene mulch at transplanting; 2) soil fumigated with methyl bromide at 448 kg/ha and black polyethylene mulch applied a month before transplanting watermelon; 3) canola (Brassica napus L. cv. Dwarf Essex) (Columbia Grain, Clarkston, WA) incorporated into soil and black polyethylene mulch applied 1 month later when watermelons were transplanted; 4) canola incorporated into soil and soil covered immediately with black polyethylene mulch at incorporation time; 5) mustard (B. juncea [L.] Czern cv. Cutlass) (Dr. K. Downey, Saskatoon Research Center, Saskatoon, Canada) incorporated into soil and black polyethylene mulch applied 1 month later at transplanting; and 6) mustard incorporated into soil and black polyethylene mulch applied immediately at incorporation time.
On 24 November 2003 and 3 November 2004, 672 kg/ha of 10-10-10 N-P-K fertilizer was applied followed by direct seeding of canola and mustard at 2.24 and 4.48 kg/ha, respectively (Hair, 2001) using a Brillion turfmaker junior seeder (Brillion Iron Works Inc., Brillion, WI). The field was watered as needed with overhead irrigation. In 2005, 672 kg/ha ammonium sulfate mixed with 1 kg/ha boron (Solubor®) was applied 3 months after seeding (Hair, 2001).
To determine the biomass of canola and mustard, plants were dug from two randomly chosen 0.25-m2 areas per plot (Gardiner et al., 1999), approximately 1 week before incorporation on 15 March 2004 and 30 March 2005. The plants were counted, washed, and oven-dried at 50oC for 1 week. The plants were then cut at the crown to separate roots and shoots, and dry weights of each were measured.

A month after the incorporation of mustard and canola, seedless watermelon cv. Tri-X® 313, susceptible to Fusarium wilt, was transplanted into single-row plots. Transplants were spaced 30 cm apart within rows. ‘Black Diamond’ pollenizer plants were transplanted into rows between plots. Plots were watered and fertigated using drip irrigation. During the growing seasons, recommended preventative fungicides were applied to manage foliage diseases. On 28 June to 13 July 2004, and 20 July to 5 August 2005, fruits were harvested from a 12-m length in the middle of each plot and weighed.
The number of plants in each plot was counted weekly until canopies of adjacent plants joined. To identify pathogens causing damping-off and wilt, diseased watermelon plants were collected on 25 May 2005 from all treatments and washed under running tap water and portions of necrotic lesions on stems and roots were cultured on semi-selective media. The recovered fungi were identified.
To rate the percentage of wilted foliage in each plot, plots were divided into quarters and disease severity assessments were made in each quarter plot on the canopy just before the first harvest and then on weekly intervals using a modified Horsfall-Barratt 15-point rating scale (Keinath and DuBose, 2004).
In 2005, 20 hyphal tips from colonies of Pythium spp. randomly selected from soil dilution plates to represent all treatment plots were transferred to cornmeal agar (CMA). Nineteen Pythium isolates recovered from watermelon roots in 2005 also were transferred to CMA. Isolates were identified to species using a grass-leaf blade culture technique (Van der Plaats-Niterink, 1981). At desired time intervals (2 to 4 days for asexual structures and 3 to 10 days for sexual structures) leaf samples were placed on a slide, stained with 0.05% trypan blue with lactophenol, and examined with a compound microscope at 400× and 1000× (Martin, 1992).
On July 15 2004, four vine pieces were collected from each treatment plot. The vines were cut into 1-cm sections, disinfected in 0.6% NaOCl for 1 minute, rinsed in sterile distilled H2O for 1 minute, and placed on Komada’s media with five segments from one vine on each plate (four plates per plot). From these plates, 43 isolates of F. oxysporum were recovered. On May 29 2005, 36 isolates were obtained from wilted 2-month-old watermelon plants isolated as previously described. Three agar plugs from 1-week-old single-spore F. oxysporum isolates were placed in 50 mL of potato dextrose broth in a 250-mL flask and shaken at 150 rpm at ambient temperature (22 to 24 oC) for 4 days. Cultures were decanted through sterile cheesecloth in a sterile funnel into sterile 50-mL centrifuge tubes and the fungal mats were washed with 5 to10 mL sterile distilled water to dislodge conidia. Culture filtrates were centrifuged at 5,000 rpm for 10 min to pellet spores, and spores were resuspended in 10 mL of sterile distilled water. Spore concentration was determined by counting with a hemocytometer and concentration was then adjusted to 106 microconidia/mL. Isolates with low concentrations of conidia were not used in pathogenicity and race tests.
Pathogenicity tests were set up in a completely randomized experiment with five and three replications, respectively. A replication consisted of four seedlings in a 15-cm diameter pot. Black Diamond was used for all pathogenicity tests. Watermelon was seeded in flats containing 1:1 v/v sand: vermiculite. Two-week-old seedlings were removed and their roots were washed under running tap water. To inoculate plants, roots were dipped into the conidium suspension and transplanted into 15-cm diameter pots containing a 4:1:1 (v/v/v) mix of sand: peat: vermiculite (Egel et al., 2004). Roots of negative-control plants were dipped in sterile distilled water. Two isolates of F. oxysporum f. sp. niveum race 2, FON 997 and FON 998 (isolated from watermelon cv. Stars and Stripes, Colleton County, SC), were included as positive controls. Wilted plants were counted weekly starting 2 weeks after inoculation for a total of 6 weeks. Temperature was set at 28oC in the greenhouse during experiments, but ranged from 16 to 40C over the course of the experiment.
Analysis of variance was performed with PROC GLM of SAS (SAS Institute, Cary, NC; release 8.2 for personal computers). All data sets were checked for normality, and Hartley’s test for equality of variance was calculated. In both years colony forming units (CFUs) of F. oxysporum, Pythium spp., and fluorescent Pseudomonas spp. were transformed to logarithm values. Pre-planned comparisons using single-degree-of-freedom contrast statements were used to compare effects of treatment on microbial populations. For pre-treatment microbial densities, means of similar treatments (i.e., two non-planted, two canola, and two mustard treatments) were pooled before comparisons were made. Treatment means for final incidence of damping-off, Fusarium wilt severity, and yield were compared using Fisher’s Protected LSD (P=0.05). For glucosinolate concentrations, means were pooled within species and standard errors calculated. Single-degree-of-freedom contrast statements were used to compare total glucosinolate concentrations.

To determine pre-incorporation inoculum density of F. oxysporum, R. solani, Pythium spp., and S. rolfsii and population density of flourescent pseudomonads, soil samples were taken on 9 March 2004 and 25 and 26 March 2005. Post-incorporation inoculum densities were assayed on 27 April 2004 and 18 April 2005. In 2005, F. oxysporum was also quantified at the end of the season on 16 August. Using a 2.5-cm diameter soil probe, 30 soil cores were taken to a depth of 15 cm from each plot. Soil samples from each plot were thoroughly mixed before assaying for microorganisms.
To estimate the inoculum density of Pythium spp., 10 g of soil from each plot was added to 200 mL of 0.3% water agar and shaken for 30 seconds. Three aliquots (0.5 mL on each plate) were spread on plates of pimaricin-ampicilin-rifampicin-pentachloronitrobenzene (PCNB) medium prepared with pimaricin at 5 mg/L (P5ARP) (Jeffers and Martin, 1986). Plates were incubated at 20oC for 20 h and then adhering soil was washed away. Plates were reincubated at 20oC and large-sized colonies (ca. > 1 cm) were counted at 36 to 48 h after plating (randomly selected colonies were later identified to species level using morphological techniques, described later). Colony counts were expressed as colony forming units (CFU) per gram of soil dried for 24 h at 100oC.
To estimate the inoculum density of Rhizoctonia solani, 300 g of soil from each plot was wet-sieved through a # 18 mesh sieve (1-mm pore openings) to recover organic matter (van Bruggen and Arneson, 1986). The organic matter was air-dried at 22 to 24oC overnight. Dried organic matter from each sample was placed in 10 small heaps of equivalent size on ethanol-potassium-nitrate agar, prepared with 2% ethanol and 20 µL/L prochloraz (Trujillo et al., 1987). Total number of heaps with and without colonies of R. solani growing from them was recorded after incubating 3 to 4 days in the dark at 23 to 25oC. Hyphae of selected colonies were stained in situ with 3% KOH and alkaline safranin O stain then examined at 200× to 400× using a compound microscope to determine the number of nuclei per cell (Bandoni, 1979).
To estimate the population density of flourescent Pseudomonas spp., 10 g of soil from each plot was added to 90 mL sterile distilled water and shaken for 30 seconds. Further ten-fold dilutions were made and three 0.1-mL aliquots of 10-1 and 10-2 dilutions were spread on S-1 medium (Gould et al., 1985). S-1 plates were incubated in the dark at 23 to 25oC for 3 to 4 days and colonies of flourescent Pseudomonas spp. were counted under long-wave ultraviolet light. All colony counts were expressed as colony forming units (CFU) per gram of soil dried for 24 h at 100oC.
To estimate the inoculum density of Sclerotium rolfsii, 300-g aliquots of soil from each plot were air dried overnight in aluminum pans (30 × 24 × 4 cm) at ambient temperatures (22 to 24 oC). Dried soil was spread evenly in pans and moistened with 75 mL of 1.33% (v/v) methanol (Rodriguez-Kabana et al., 1980). Aluminum pans then were put into large Ziploc® plastic bags and incubated at 30oC, 3 to 4 days. Germinated sclerotia were counted by closely examining the soil surface visually for discrete colonies of S. rolfsii.
To estimate the inoculum density of F. oxysporum, 10-g of soil from each plot was added to 90 mL of sterile distilled water and shaken for 1 minute. Further ten-fold dilutions were made and three 0.1-mL aliquots of 10-1 and 10-2 (an aliquot per plate) were spread on Komada’s medium (Komada, 1975). The plates were held under diurnal light (16-hr photoperiod) at ambient temperature (22 to 24 oC). Colony counts were expressed as colony forming units per gram of soil dried for 24 h at 100oC.

To quantify glucosinolate content in the canola and mustard crops, five additional plants were dug from each plot 1 week prior to incorporation, put into plastic bags, and held on ice in a cooler chest. In 2004, the canola and mustard plants were immediately taken to the laboratory where roots and shoots were cut separately into small pieces. Two 5-g subsamples from each plot were frozen in liquid nitrogen before being freeze-dried. In 2005, the plants were stored at -80oF for 12 months before freeze drying. Freeze-dried plant parts were ground using a Wiley mill equipped with a 1-mm mesh sieve. The powder was kept in a screw-cap test tube and stored at 5oC in a refrigerator until it was assayed for glucosinolates 2 weeks later.
Glucosinolates were extracted from 300 mg of freeze-dried root and shoot tissues using the procedure described by Magrath et al. (1993) with modifications employed by Kirkegaard and Sarwar (1998). To quantify desulphoglucosinolates, 2-propenyl and benzyl glucosinolates were used as internal standards. Levels of other glucosinolates were determined using response factors for desulphoglucosinolates published by the European Economic Community (Commission Regulation [EEC] No 1864/90) (Kirkegaard and Sarwar, 1998). Desulphoglucosinolates were analyzed by injecting a 30 µL sample into Hewlett Packard 1090 high pressure liquid chromatograph (Agilent Technologies, Wilmington, DE). Peaks were identified using pure standards. Forty-eight and 24 samples were analyzed in 2004 and 2005, one sample per replication and one sample from half of the replications, respectively. To calculate glucosinolate concentration incorporated per square meter, glucosinolate concentrations per gram were multiplied by biomass per square meter (Morra and Kirkegaard, 2002; Sarwar and Kirkegaard, 1998).

Canola and mustard plants were mowed and then rototilled (Ferguson Tilrovator, Ferguson Mfg. Co., Suffolk, VA) 15 cm deep into the soil on 17 March 2004 and 5 April 2005. In 2004, 50% of canola and 10% of mustard plants had flowered whereas approximately 90% of canola and 50% of mustard plants had flowered in 2005. Most of the non-flowering plants had unopened flower buds.
After incorporation, soil samples were taken 0 to 15 cm deep using a 2.5-cm diameter soil probe at 6 time intervals: 0 h and 6 h, and 1, 2, 12, and 26 days after incorporation in 2004 and 0 h and 6 h, and 1, 2, 4, and 12 days in 2005 . Soil cores were placed in a 50-mL Pyrex centrifuge tube containing 5 mL of 0.2 M CaCl2 and 12 mL of dichloromethane with 0.1% cyclohexane as an internal standard (Gardiner et al., 1999). A total of 216 samples (6 treatments x 6 replications x 6 sampling times) collected in 2004 and 90 samples (6 treatments ×3 replications x 5 sampling times) collected in 2005 were analyzed. Fewer samples were analyzed from 2005 because isothiocyantes were not detected at the later sampling times in 2004. Gardiner’s et al. (1999) methods were used for extraction. Samples were analyzed on a Hewlett Packard 5890 gas chromatograph (GC). External standards used for quantification included allyl isothiocyante (Lancaster Synthesis, Inc., Pelham, NH), N-propyl, phenyl, benzyl, methyl, and 2-phenylethyl isothiocyanate and goitrin (oxazolidinethione) (Alfa Aesar, Ward Hill, MA).