Use of green manures to reduce inoculum production of Fusarium graminearum on wheat residues

Final Report for GNC05-054

Project Type: Graduate Student
Funds awarded in 2005: $10,000.00
Projected End Date: 12/31/2006
Grant Recipient: University of Minnesota
Region: North Central
State: Minnesota
Graduate Student:
Faculty Advisor:
Dr. Ruth Dill-Macky
University of Minnesota
Faculty Advisor:
Dr. Linda Kinkel
University of Minnesota
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Project Information

Summary:

This study investigated the use of green manures for reducing Fusarium graminearum survival in association with wheat residues in greenhouse and field experiments. Green manures promoted the development of higher densities and Fusarium-antagonistic abilities of F. graminearum-antagonists in soils. The use of green manures did not significantly impact the survival of F. graminearum in wheat residue. However, streptomycete densities and F.graminearum-antagonist densities were significantly positive correlated with reduced survival of Fusarium. These results suggest that the use of green manures can enhance populations of indigenous soil microorganisms antagonistic to the survival of F. graminearum in wheat residue.

Introduction:

Fusarium head blight (FHB or scab) is one of the most economically devastating diseases of wheat (Triticum aestivum L.) (Windels, 2000). Studies have documented a significant increase in the frequency of FHB epidemics worldwide over the last 15 years (McMullen, 2005; Windels, 2000). Some of the proposed causes for the upsurge of FHB epidemics include an increase in the frequency of wet weather, especially between anthesis and harvest; the planting of highly susceptible wheat cultivars; and the widespread adoption of conservation tillage practices (Dill-Macky, 1996; McMullen et al., 1997).

Fusarium head blight is caused by several species from the genus Fusarium. F.graminearum Schwabe (Gibberella zeae (Schwein.) Petch [telemorph]) is most frequently associated with this disease in the United States (Wilcoxson et al., 1988; Wong et al., 1992). Crop residues are reported as the major source of ascospores, which are considered the primary inoculum in FHB epidemics (Reis, 1988; Sutton, 1982). The residues of corn (Zea mays L.) and small grain cereals (Champeil et al., 2004; Dill-Macky and Jones, 2000; Sutton, 1982; Wiese, 1987), are particularly important contributors of the inoculum initiating FHB epidemics. Although F. graminearum readily produces conidia on the infected spikes of cereal plants, FHB is generally regarded as a monocyclic disease (Bai and Shaner, 1994; Khonga and Sutton, 1988). Because residues are the principal inoculum reservoir, inoculum densities are associated with the amount of crop residue and the degree of infestation (Shaner, 2003; Dill-Macky and Jones, 2000; Pirgozliev et al., 2003). Pereyra and Dill-Macky (2004) concluded that the node tissues of wheat, being slower than other tissues to decompose, provide an ideal site for inoculum production over extended periods. The survival of F. graminearum in residues will however be affected by other microbes, either directly by competition, parasitism and predation, and/or indirectly by influencing the rate of residue decomposition.

Several studies have focused on reducing the inoculum produced by crop residues (Dill-Macky and Jones, 2000; Khonga and Sutton, 1988; Pereyra and Dill-Macky, 2004; Sutton and Vyn, 1990). A relatively new approach to reduce pathogen populations in crop residues is managing indigenous soil microbial communities (Peters et al., 2003; Sturz et al., 1997; Weller et al., 2002; Zaitlin et al., 2004). Organic amendments may be used to increase the activity of indigenous microbes that act as biocontrol agents (Lockwood, 1988). Bossio et al. (1998), Mazzola (2004) and Govaerts et al. (2006) reported that cultural practices, soil type, plant species, and plant genotype are significant factors determining the composition of soil microbial communities. The soil community requires carbon energy sources, and different sources may promote some microorganisms over others (Bailey and Lazarovits, 2003). Cover crops and green manures have been reported as especially effective in increasing the proportion of pathogen inhibitory microorganisms within indigenous microbial populations (Davis et al., 1994; Lupwayi et al., 1998; Mazzola et al., 2001; Wiggins and Kinkel, 2005a,b).

The incorporation of green manures has been shown to increase the density and diversity of microbes in soil, particularly the density and the pathogen-inhibitory activity of bacteria such as fluorescent Pseudomonas spp. (Mazzola et al., 2001; Bulluck and Ristaino, 2002), non-pathogenic Fusarium spp. (Davis et al., 1994; Alabouvette et al., 1996), streptomycetes and actinomycetes (Mazzola et al., 2001; Wiggins and Kinkel, 2005a, b). Several authors have reported effective disease control using green manures to promote the increase of Streptomyces populations acting as biocontrol against diseases caused by other Streptomyces (Wiggins, 2003; Lazarovits et al., 1999), Phytophthora (Jones and Samac, 1996; Xiao et al., 2002), Pythium (Chamberlain and Crawford, 1999; Jones and Samac, 1996) and Rhizoctonia (Chamberlain and Crawford, 1999; Wiggins, 2003). Furthermore, Chamberlain and Crawford (1999) demonstrated that inoculation with Streptomyces hygroscopicus effectively controlled Fusarium oxysporum Schlechtendahl amend. Snyder & Hansen in turfgrass production. Wiggins (2003), found that survival of Fusarium oxysporum was significantly lower in soils following the incorporation of a buckwheat green manure compared to control treatments and that greater pathogen inhibitory activity of the streptomycete population was consistently associated with this result. There is however no literature on the use of green manures to promote indigenous streptomycete populations to control FHB.

Fusarium head blight is one of the most difficult diseases to control due to the lack of effective genetic resistance, the wide host range of the pathogen, and the long distance dispersal of inoculum. The available control measures, including the use of moderately resistant varieties, fungicides and cultural practices are only partially effective (Luz et al., 2003). Due to the lack of a single effective practice to control FHB, the integration of several different management practices appears necessary. By improving our knowledge of how FHB-antagonists might be manipulated using green manures, it may be possible to expand the available management options.

Project Objectives:
  • Determine the effects of green manures on the frequency of soilborne antagonists inhibitory against pathogenic F. graminearum, and on the intensity of their F. graminearum inhibition.

    Quantify the impacts of green manures on the survival of F. graminearum on wheat residues.

    Evaluate the effects of green manures on the rate of decomposition of F. graminearum-infected residue.

Cooperators

Click linked name(s) to expand
  • Ruth Dill-Macky
  • Linda Kinkel

Research

Materials and methods:

A series of greenhouse and field experiments were conducted to evaluate the effects of green manures on the frequency and intensity of Fusarium-inhibitors, wheat residue decomposition and Fusarium survival in residue by soil microbes.

In each experiment, wheat was grown and wheat residues, naturally infested by Fusarium, were incorporated into the soil. Two green manures, sorghum-sudangrass [Sorghum bicolor (L.) Moench-S. bicolor (L.) Moench var. sudanense (Piper)] hybrid and common buckwheat (Fagopyrum esculentum Moench) were evaluated and compared to a fallow (no green manure) treatment. Changes in total bacterial density in the soil, the density of streptomycetes in the soil, the density of F. graminearum-antagonists in the soil, and the inhibitory activity of F. graminearum antagonists were evaluated before and following the incorporation of the green manures. Soil characteristics (pH, organic matter, K, P) were examined along with the frequency of Fusarium spp. especially F. graminearum, in the wheat residue. The decomposition rate of the infested residue was also evaluated over time.

Greenhouse experiments:

Five experiments were conducted in the greenhouse facilities at the University of Minnesota, Saint Paul, MN.

Experiment 1. Waukegan silt loam soil (fine-silty over sandy, mixed Typic Hapludoll) was collected at the Minnesota Agricultural Experiment Station, Saint Paul, MN (44 degrees 59’ N, 93 degrees 10’ W). The field plot where the soil was collected had been in a soybean-wheat rotation over the last ten years, and had been planted to wheat in the previous cropping season. The site has a history of moderate to severe FHB. Soil was stored in plastic containers at room temperature until needed (approx. 45 days). Wheat residue (cv. Oxen) was collected from a field plot in Baker, MN and stored at -20 degrees C until used (approx. 55 days). F. graminearum was recovered from 72% of nodes at the start of the experiment.

Field soil was sieved using a 6 mm mesh sieve to remove large pieces of plant material. Stem pieces, each 2.5 cm long and containing a node, were cut from the collected residue. Residue pieces were mixed thoroughly with the collected soil and perlite (5:1, vol/vol) at a rate of 100 pieces per 7 kg of soil using a cement mixer. The soil-perlite-residue mix was then transferred into 11-L plastic pots (approx. 25 cm diameter).
The experimental design was a randomized complete block with 15 replicates and three treatments. Seeds of sorghum-sudangrass (cv. Excel, 30 seeds per pot) and buckwheat (cv. Mancan, 20 seeds per pot) were planted 1-cm deep and distributed uniformly across the pot.

Pots were watered and weeded by hand as necessary. The greenhouse was maintained at 20-22 degrees C throughout the experiment. Two weeks after planting, pots were thinned to 20 sorghum-sudangrass plants and 15 buckwheat plants per pot, respectively. Six weeks after planting, the aboveground biomass of the green manure treatment from each pot was harvested, cut into approximately 1-cm long pieces and incorporated by hand into the soil of the pot in which it had grown. Soil from fallow-treated pots was similarly disturbed although no residues were incorporated.

Experiment 2. Waukegan silt loam soil was prepared as for Experiment 1 described above. Wheat residue (cv. Wheaton), naturally infested with F. graminearum, was collected from field plots at the Minnesota Agricultural Experiment Station at Rosemount, MN and stored at -20 degrees C. Isolation tests indicated that 55% of nodes were infested by F. graminearum at the initiation of the experiment. The residue was incorporated into the soil, as described for Experiment 1. The experiment was established following the methods described for Experiment 1.

Experiment 3. Elmville fine sandy loam soil was collected from a field plot in Baker, MN (46 degrees 43’ N, 96 degrees 33’ W) that had been maintained in a corn-soybean-wheat rotation and planted to wheat in the previous season. Soil was stored in sealed plastic containers at room temperature until needed (approx. 90 days). Wheat residue (cv. Alsen) was collected from field plots on the Minnesota Agricultural Experiment Station at Saint Paul and stored at -20 degrees C until needed. Residue was naturally infested by F. graminearum; 58% of nodes were infested at the start of the experiment. The experiment was established following the methods described for Experiment 1.

Experiment 4. Tallula silt loam soil (coarse-silty, mixed, mesic Typic Hapludoll) was collected from the Minnesota Agricultural Experiment Station at Rosemount, MN (UMore Park, 44 degrees 42’ N, 93 degrees 06’ W). Soil was stored in sealed plastic containers at room temperature until needed (approx. 50 days). The field had been maintained in a corn-soybean rotation and wheat was grown in the last season. The wheat crop had been inoculated twice with F. graminearum (Zadoks stage 65 and 75 respectively; Zadoks et al., 1974) to ensure the presence of infected residue using a suspension of macroconidia (2.0 x 105 macroconidia per ml). Inoculum was produced following the methods described by Evans et al. (2000), and sprayed using a gasoline-powered backpack sprayer. The macroconida suspension of F. graminearum was mixed with Tween 20 (polysorbate; Fisher Biotech, Fair Lawn, NJ) and applied at a rate of 17.5 l of F. graminearum suspension and 194 ml of Tween 20 per 1,000 m2 each inoculation date. The inoculum was prepared using a mixture of 12 F. graminearum isolates (#10102027, 10102030, 10102042, 10102066, 10102116, 10102117, 10103006, 10103007, 10103015, 10103043, 10103054, 10103055) from the collection at the Small Grains Pathology Laboratory, Department of Plant Pathology, University of Minnesota. Residue from this wheat crop (cv. Oxen) was collected and stored at -20 degrees C until needed. The experiment was established following the same methods described for Experiment 1.

Experiment 5. This experiment used the same soil, residue and experimental design as Experiment 4, except that the experiment was established one month later.

Field experiments:

Field experiments were used as scaling factor. Two adjacent field experiments were simultaneously established at the Minnesota Agricultural Experiment Station in Rosemount, MN (UMore Park). Plots (3 x 5 m) were established in a randomized complete block design with six replicates. Plots were separated by 4 m within a block and 5 m between blocks. Experiments were separated by 20 m. Wheat (cv. Olsen) was planted, and inoculated (as described above).

The experiments were both plowed using a chisel plow (John Deere 1610; John Deere Co., Moline, IL). The green manures were planted on August 18 for Experiment A and September 1 for Experiment B. Sorghum-sudangrass (cv. Excel; planting rate, 93 kg of viable seed/ha) and common buckwheat (cv. Mancan; planting rate, 109 kg of viable seed/ha), were planted in 21-row plots, 5 m long and with 17 cm inter-row spacing, using a 7-row drill planter (Kincaid Equipment Manufacturing Corporation, Haven, KS). No fertilizer was applied.

On October 6, the green manures were incorporated in both experiments using a rotovator (Kuhn 690; Kuhn Farm Machinery Inc., Vernon, NY) that incorporated the green manure to a 10 cm depth. The rotovator was also used in fallow plots to produce similar soil disturbance as in the green manure treatment plots.

Data collection

Greenhouse experiments. The aboveground biomass (fresh weight per m2) of each green manure treatment was determined at the time of incorporation in all experiments except Experiment 1. Fresh weights were converted to dry weights, assuming 18% of dry matter for sorghum-sudangrass hybrid and 25% of dry matter for buckwheat, and expressed as kg.ha-1. The percentage of dry matter for each green manure species was estimated from fresh and dry weights obtained following a sampling of the aboveground biomass of each green manure species in the two field experiments. Biomass was collected the day prior to green manure incorporation from three arbitrarily placed 0.5 x 0.5 m quadrats in each plot. Biomass samples were placed in plastic bags and stored at 4 degrees C until processed. Samples were subsequently dried with forced air at 35 degrees C for 48 h to determine dry matter weight.

Soil and wheat residue samples were collected from every pot when each treatment was established, at the time of green manure incorporation, four weeks after green manure incorporation, and 12 weeks after green manure incorporation. For each pot, three cores of 2.5-cm diameter were taken from the top 20 cm of soil, bulked, and stored in plastic bags at 8 degrees C until they were processed. Twenty nodes were arbitrarily collected at each sampling time and stored in paper bags at -20 degrees C until processed. The remaining soil was returned to the pot after each sampling.

Field Experiments. Soil samples were collected when the plots were established, at the time of incorporation of green manures, five weeks after incorporation, and six months after incorporation. Soil samples consisted of five cores of 2.5-cm diameter collected from the top 20 cm of soil per plot. The five cores, collected at five sites distributed evenly across each plot, were bulked and stored in plastic bags at 8 degrees C until processed.

Wheat residue samples were collected when the green manures were planted, five weeks after the incorporation of green manures and six months after incorporation. Fifty residue pieces, each with a node, were collected from each plot at each sampling time. Specifically, 10 arbitrarily selected nodes were taken at each of the five locations where soil sampling was conducted within a plot. Only residue pieces where the node was below the soil surface were collected. Node pieces were placed in paper bags and stored at -20 degrees C until processed.

The day prior to green manure incorporation, aboveground plant biomass was assessed. Due to the high frequency of volunteer wheat plants in all plots, green manure biomass (sorghum-sudangrass or buckwheat) and wheat biomass were separated and the fresh and dry matter weight of each component determined as described above.

In field experiments, weather data were obtained from the University of Minnesota’s Climatology Working Group, which maintains a weather station at the Minnesota Agricultural Experiment Station, Rosemount, MN (44 degrees 55’ N, 93 degrees 07’ W) (http://climate.umn.edu). Parameters included air temperature, precipitation, and the date of the first frost for the year.

Soil microbial assessment

Total cultivable soil streptomycete and bacterial population. Streptomycete densities for all soil samples were estimated using a modification of Herr’s method (Herr, 1959) as described by Wiggins and Kinkel (2005a). Briefly, soil samples were dried overnight and 5 g of soil were placed in 50 ml of distilled water and shaken for 1 h on an orbital shaker (Environ-Shaker 3597, Lab-line Instruments, Inc., Melrose Park, IL) at 175 rpm and 4 degrees C. Soil suspensions were serially diluted (1:10) three times and 100 μl of the diluted suspensions were plated onto water-agar (WA) medium (10 g of Bacto agar per liter dH2O) in Petri plates. Plates were then overlaid with 5 ml of molten starch casein agar (SCA) medium (10 g of starch, 0.3 g of casein, 0.02 g of CaCO3, 2.0 g of KNO3, 2.0 g of NaCl, 2.0 g of K2HPO4, 0.05 g of MgSO4.7H2O, 0.01 g of FeSO4.7H2O, 0.001 g of ZnCl2, and 15 g of agar per liter of dH2O), and allowed to solidify. Plates were incubated at 28 degrees C for seven days in darkness. This double-layer agar method (WA/SCA) results in the preferential growth of streptomycetes. All soil samples were also diluted as described above, and plated onto oatmeal agar (OA) medium (20 g of oatmeal, 15 g of Bacto agar, and 1 g of casamino acid per liter of dH2O) amended with cycloheximide (100 μg/ml), polymyxine (5 μg/ml) and penicillin (1 μg/ml). Oatmeal agar plates were incubated at 28 degrees C for seven days in darkness. Two replicate plates per sample were prepared on each medium at each dilution. Streptomycetes and total bacteria were counted on plates of WA/SCA and OA, respectively, and expressed as the number of colony-forming units (CFU)/g of dry soil.

Activity of streptomycete F. graminearum antagonists. The assessment of the antagonistic activity of soil streptomycetes against pathogenic F. graminearum was performed using a modification of Herr’s method (Herr, 1959) as described by Wiggins and Kinkel (2005a). Soil dilutions were performed as described above, and plated onto Petri plates containing 15 ml of WA. Plates were subsequently overlaid with 5 ml of molten WA and incubated at 28 degrees C for three days in darkness. On the third day, each plate was overlaid with potato dextrose water agar (PDWA) medium (2.4 g of potato dextrose broth, and 10 g of Bacto agar per liter of dH2O), and inoculated with a F. graminearum-potato dextrose agar (PDA) suspension prepared as follows.

A seven-day old culture of F. graminearum on PDA medium (39 g of potato dextrose agar per liter of dH2O) was cut into approximately 1 cm2 pieces with a sterile knife and blended with 100 ml sterile water in a sterile flask in a blender (Model 31BL92; Waring Commercial Blendor, New Hartford, CT) for three 5 sec and one 10 sec periods, sequentially. All assays were performed with a single F. graminearum isolate (University of Minnesota, Small Grains Pathology accession 10102067), known to be highly aggressive on wheat. Following blending, 4 ml of the F. graminearum-PDA suspension were added to 100 ml molten (50-55 degrees C) PDWA and mixed thoroughly. Ten milliliters of the resulting suspension were overlaid on the top of each dilution plate and allowed to solidify. Plates were then incubated at 28 degrees C for four days in darkness. Each soil sample was plated onto three replicate plates.

Streptomycetes were considered antagonistic if they produced a clear inhibition zone (no F. graminearum growth) of at least one millimeter surrounding the colony edge. Antagonist densities were expressed as CFU/g dry soil. The proportion of streptomycetes antagonistic against F. graminearum in each soil was expressed as a percentage (number of colonies identified as antagonists divided by the total number of streptomycete colonies x 100). The diameter of each zone of inhibition against F. graminearum was calculated by subtracting the diameter of the colony from the inhibition zone diameter including the colony size. For each Petri plate the mean inhibition zone size per antagonist colony was determined.

Soil chemical analyses. Soil tests were performed at the Soil Testing Laboratory at the University of Minnesota (Methods used in the laboratory are available at: http://soiltest.coafes.umn.edu/methods.htm).
Soil pH, phosphorus (P2O5), potassium (K+) and organic matter content were determined for the soil samples of six arbitrarily selected replicates from each of Experiments 1 and 3 (collected four weeks after green manure incorporation), and on all samples from field Experiment A (collected five weeks after green manure incorporation).

Residue decomposition. The decomposition of wheat residue was quantified by measuring losses in dry weight over time. The collected residue for each sample was rinsed in tap water to remove adhered soil, and dried with forced air at 35 degrees C for 24 h prior to weighing. Twenty residue pieces from each plot or pot in greenhouse and field experiments (one centimeter long each containing a node) were analyzed at each sampling time, except at the last sampling time (12 weeks after green manure incorporation) in greenhouse experiments, when fewer residue pieces were recovered.

Presence of F. graminearum/G. zeae in residue. Samples were processed to determine the presence of F. graminearum according to the protocol developed by Pereyra (2000). Briefly, residue pieces were rinsed in tap water to remove adhered soil and air dried at 35 degrees C for approximately 24 h. Node pieces were then surface-disinfected in 70% ethyl alcohol for 30 sec, rinsed once in sterile water, immersed in 0.6% sodium hypochlorite for 45 sec, then rinsed three times in sterile distilled water and blotted dry on sterile filter paper. Surface-disinfested node pieces were plated on Komada’s medium (KM: 1 g of K2HPO4, 0.5 g of KCl, 0.5 g of MgSO4.7H2O, 0.01 g of Fe-Na-EDTA, 2.0 g of L-Asparagine, 20.0 g of D(+)-Galactose, 15.0 g of agar, 1.0 g of pentochloronitrobenzene, 0.5 g of Oxgall, 1.0 g of Na2B4O7.10H2O, and 6 ml of streptomycin sulfate solution (300 ppm) per liter of dH2O) selective for Fusarium spp. (Komada, 1975). Twenty node pieces were processed from each plot. Node pieces on KM were incubated at 22 degrees C with 12 h light and dark cycles for 15 days. Following incubation, salmon-pink to white colonies were considered Fusarium spp. Gibberella zeae colonies were identified by transferring Fusarium colonies to carnation leaf-piece agar (CLA) medium (three to five sterile carnation leaf pieces [5 to 7 mm in diameter] per 15 ml of 1.5% water agar) (Fisher et al., 1982) where G. zeae readily forms perithecia. Colonies on CLA were evaluated for perithecia production following incubation for 15 days at 22 degrees C with 12 h light and dark cycles.

Statistical analyses. Data were subjected to analysis of variance (ANOVA) using the Generalized Linear Model procedure (PROC GLM) of SAS (release 9.1; SAS Institute, Inc., Cary, NC). When the F test was significant (P<0.05) the treatment means were compared using Fisher’s least significant difference (LSD) at P=0.05. Relationships among dependent variables were tested using Pearson correlation coefficients. The assumptions used in the ANOVA were tested using PROC UNIVARIATE (release 9.1; SAS Institute, Inc., Cary, NC) and the Box-Cox technique (Oehlert, 2000) was used to determine the transformation that best fit when needed. The binomial test was used to analyze the consistency of patterns observed for individual variables among treatments over time. Binomial probabilities were estimated using the binomial approximation of the normal distribution (http://faculty.vassar.edu/lowry/binomialX.html, Lowry, 2006; DeGroot, 1975).

Research results and discussion:

Green manure biomass varied significantly among treatments and experiments. In the greenhouse, buckwheat biomass at the time of incorporation was equivalent to 500-900 kg/ha, while sorghum-sudangrass biomass was equivalent to 90-150 kg/ha. Green manure biomass in field experiments was confounded by the presence of a substantial biomass of volunteer wheat plants in all plots, including fallow. Late-planted green manures accrued less biomass than early-planted plots.

There were substantial differences in soil characteristics among the greenhouse and field experiments. However, there were no significant differences among treatments in field or greenhouse experiment (data not shown).

Microbial densities: Bacterial and streptomycete densities varied significantly among experiments, and over time in all experiments. In greenhouse Experiments 1-3, bacterial densities increased after planting and incorporation of green manures, and remained higher than at planting 12 weeks after incorporation. In contrast, in Experiments 4 & 5 the effects of green manures on bacterial densities over time were more variable, and all soils had substantially lower densities of bacteria 12 weeks after incorporation than at the start of the experiment. In field experiments, there was no consistent effect of green manures on bacterial densities (data not shown) likely reflecting the confounding effect of volunteer wheat plants in all plots; bacterial densities increased over time in all treatments.

At individual sampling times, green manure-treated soils generally had higher bacterial densities than fallow soils in the greenhouse experiments. Specifically, within individual sampling times (excluding the first, pre-treatment sample time), sorghum-sudangrass-treated soils had higher bacterial densities than fallow soils in 13 of 15 cases (P=0.005, binomial test), though differences were statistically significant at only 4 time points. Likewise, buckwheat-treated soils had higher bacterial densities than fallow soils in 12 of 15 cases (P=0.019, binomial test), and differences were statistically significant at 7 of the 15 sample times. Overall, green manures had more substantial and consistent effects on bacterial densities over time in Experiments 1-3 than in Experiments 4 & 5.

Streptomycete densities also increased after planting and incorporation of green manures in greenhouse trials, and remained higher than at planting 12 weeks after incorporation in Experiments 1-3. As with total bacterial densities, green manures appeared to have a less consistent and long-lasting effect on streptomycete population densities in Experiments 4 & 5. Though densities increased immediately after planting and incorporation, streptomycete populations were lower at 12 weeks after incorporation than at the start of the experiment. There were no consistent effects of green manure treatments on streptomycete population densities over time in field experiments (data not shown), again likely reflecting the confounding effect of volunteer wheat plants.

At individual times, green manure-treated soil generally had higher streptomycete densities than fallow soils in the greenhouse experiments, though these differences were not significant. Specifically, at 9 of 15 (sorghum-sudangrass, P=0.302, binomial test) and 12 of 15 (buckwheat, P=0.019, binomial test)) time points, green manure-treated soils had higher streptomycetes densities than fallow soils. Effects were greatest in Experiments 1-3.

Pathogen antagonist populations: Densities of F. graminearum antagonists varied substantially over time and among greenhouse experiments. Antagonist densities increased after green manure planting and/or incorporation in Experiments 1, 3, 4, & 5, although increases were generally not sustained through 12 weeks post-incorporation. Within individual samples, green manure treatments generally increased densities of F. graminearum antagonists relative to fallow soils; in 12 of 15 (sorghum-sudangrass, P=0.019, binomial test) and 13 of 15 (buckwheat, P=0.005, binomial test) cases, green manure-treated soils had higher densities than fallow soils, though differences were rarely significant. There were no significant effects of green manures on antagonist densities in field trials (data not shown).

The inhibitory activity (mean inhibition zone size) of F. graminearum antagonists varied widely among experiments and over time. There were no clear effects of green manure treatment on inhibitory activities over time.

Relationships among microbial community measures: Pearson’s correlation coefficients calculated for greenhouse and field experiments indicate that larger bacterial communities generally had higher densities of streptomycetes and F. graminearum antagonists as evidenced by the consistent positive correlations between bacterial and streptomycete densities (12 of 14 cases, P=0.008, binomial test) and between bacterial density and the density of F. graminearum antagonists (10 of 14 cases, P=0.090, binomial test). Further, bacterial densities were also positive correlated with the proportion of streptomycetes that were antagonistic against F. graminearum (11 of 14 cases, P=0.031, binomial test); in larger communities, a greater proportion of the streptomycetes were inhibitory against F. graminearum. Finally, the density of F. graminearum antagonists was positively correlated with the intensity of pathogen inhibition (mean inhibition zone size, 12 of 14 cases, P=0.008, binomial test); communities that have lots of F. graminearum antagonists had antagonists that are better at inhibiting the pathogen than communities with fewer antagonists.

Wheat residue decomposition: In greenhouse experiments, dry weight of wheat residues declined significantly over time (data not shown). Twelve weeks after green manure incorporation, less than 50% of the weight of the original wheat residue biomass was recovered from any treatment.

Fallow-treated soils had significantly less wheat residue biomass than green manure-treated soils in 6 of 15 sample times; differences between green manure and fallow treated soils were not statistically significant at the remaining 9 sample times. Residue decomposition occurred more slowly in the field experiments than in the greenhouse; more than 80% of the wheat residue biomass was recovered from soil six months after green manure incorporation. There was no significant effect of green manures on wheat residue decomposition in field trials (data not shown).

Fusarium survival on wheat residue: Recovery of Fusarium spp. from wheat node tissue varied significantly over time in greenhouse experiments, generally declining from 70-86% to 5-14% 12 weeks after green manure incorporation. There was no consistent effect of green manures on the survival of Fusarium spp. on node tissue (data not shown). Likewise, recovery of F. graminearum varied over time, and was not consistently influenced by the green manure treatments.

Recovery of Fusarium spp. and F. graminearum on wheat nodes also declined significantly over time in field trials, though declines were more gradual than was observed in the greenhouse experiments. Six months after green manure incorporation, recovery of Fusarium spp. and F. graminearum was 20% and 5% of original values, respectively. As in greenhouse experiments, there were no consistent effects of green manures on Fusarium survival.

Relationships between residue decomposition and Fusarium survival: Survival of F. graminearum was positively correlated with residue biomass in 10 of 14 samples (P=0.090, binomial test), suggesting that larger residue biomass is more conducive to F. graminearum survival. However, there was no consistent association between residue biomass and recovery of other (non- graminearum) Fusarium spp.
Correlation between soil microbial communities, residue decomposition, and Fusarium survival: Streptomycete densities and F. graminearum antagonist densities were significantly negatively correlated with the survival of Fusarium spp. and F. graminearum on nodes. In addition, streptomycete densities and F. graminearum antagonist densities were negatively correlated with residue biomass when Pearson’s correlation coefficients were determined over all experiments: higher streptomycete and antagonist densities where associated with lower Fusarium survival and lower residue biomass. No significant correlation was found between the intensity of F. graminearum inhibition (mean inhibition zone size) and Fusarium survival or residue biomass.

DISCUSSION

Sorghum-sudangrass and buckwheat green manures generally increased the densities of F. graminearum antagonists relative to fallowed soil, though the differences were not always statistically significant. Among all experiments, the significant negative correlation between F. graminearum antagonist densities and survival of Fusarium on wheat residue suggests that increases in antagonist densities offer potential as a means for controlling FHB. However, the increases in F. graminearum densities following green manure treatments were insufficient for inducing consistent reductions in F. graminearum survival in wheat residue in field and greenhouse trials.

This is the first report demonstrating the potential of soil microbial communities to reduce infestation of wheat residue by F. graminearum. Though a single cycle of green manure was insufficient to significantly impact F. graminearum survival, our results suggest that further investigation on the use of strategies for enhancing soil antagonist populations using green manures are warranted.

The impact of green manures on soil microbial populations is generally thought to result from nutrients made available to microbes after the incorporation of the green manures. Shifts in bacterial densities observed in the green manure-treated soils of this study reflected the effects of plant roots on soil microbial activity prior to incorporation. It is likely that root exudates during the growing period of the green manures increase bacterial populations in soil as has been reported by other researchers (Abawi and Widmer, 2000; Kirkegaard et al., 2004; Marschner et al., 2001; Westover et al., 1997).

High soil microbial populations increase the competition for nutrients among microorganisms and are likely to facilitate increases in the density of antibiotic-producing bacteria (Alabouvette et al., 1996). In our study, high soil bacterial densities were associated with higher streptomycete densities, and in turn higher densities of F. graminearum antagonists. Thus, green manures increased the total bacterial population along with the density of F. graminearum antagonists, though these increases were not always significant. Increases in soil microbial diversity and microbial activity in soil, specifically streptomycetes and actinomycetes, in response to the incorporation of green manures have been reported previously (Mazzola et al., 2001; Wiggins and Kinkel, 2005a,b).

The strong positive correlation between the inhibitory activity of F. graminearum antagonists and the increased density of antagonists suggests that more competitive isolates, such as those that produce antibiotics, dominate the population when population densities are high. Wiggins (2003) reported a similarly strong positive correlation between the density of streptomycete antagonists against Verticillium dahliae and the mean inhibition zone size produced by individual isolates. The results of these studies indicate that a density-dependent directional selection on soil microbial communities may be facilitated by green manures.

The initial density and intensity of F. graminearum antagonists and their response to treatments varied among the soils tested. However, the characteristics of soils evaluated in this study (i.e. pH, P2O5, K+ and OM) did not explain the differences in F. graminearum antagonist populations, since significant differences in soil characteristics were not found among treatments (data not shown). Many factors can affect soil microbial communities, including soil texture, cultural practices and plant species present (Bossio et al., 1998; Mazzola, 2004). It is likely that these factors contributed to the variable response to treatments found among soils in this work.

The rapid decomposition of wheat residue in the greenhouse experiments suggests that conditions for residue decomposition were more favorable in the greenhouse than in the field conditions of this study. Parr and Papendick (1978) and Blevins et al. (1994) reported that warm temperatures and moist soil conditions accelerate residue decomposition. In our study, greenhouse temperatures (averaging 22C) and soil moisture likely promoted the rapid decomposition of the wheat residues. Decomposition was slower in field experiments than in greenhouse experiments, and was consistent with previous work (Pereyra and Dill-Macky, 2004), showing slow decomposition over winter months due to reduced microbial activity in frozen soils. In our study, residue decomposition was generally greater in fallowed soils than in the green manure-treated soils under greenhouse conditions. Soil moisture in the fallow treatment tended to be higher than the soil moisture in the soils treated with green manures, presumably as growing plants were actively removing water from the soil through transpiration. Thus, greenhouse experiments might be inappropriate to detect the effects of microbial communities on residue decomposition.

The substantially lower survival of Fusarium spp. observed in this study as compared with previous studies appears to be unprecedented. Pereyra and Dill-Macky (2004) reported that the proportion of Fusarium-infested node pieces recovered from field experiments remained greater than 80% for two years after harvest. The reduced survival observed in the greenhouse experiments reported here may be associated with the water content of the soil and temperature which may have been more favorable for residue decomposition, and/or those saprophytes that compete with Fusarium spp. In field experiments, the substantial decline in the percentage of nodes colonized by Fusarium spp. might be related to unfavorable environmental conditions for FHB infection that resulted in a limited colonization of residues. The degree of colonization of tissues may influence the ability of Fusarium spp. to compete over time with other saprophytes.

The lack of significant a significant effect of green manures on Fusarium spp. survival in field experiments may well have been confounded by the substantial biomass of volunteer wheat. It is feasible that the biomass from volunteer wheat plants was as effective as the green manures in promoting the activity and proliferation of F. graminearum antagonists. A preliminary greenhouse study indicated that young wheat plants incorporated into the soil had a beneficial effect on F. graminearum antagonists (data not shown). The incorporation of wheat biomass increased F. graminearum antagonist densities in soils and their inhibitory activity (mean inhibition zone size per isolate) in 43% and 35% respectively, compared to those in fallow-treated plots. These results support the hypothesis that volunteer wheat plants acted as green manure and thus likely confounded the experimental results in field experiments.

Similarly, the large amount of volunteer wheat biomass likely also confounded the effect of green manures on residue decomposition. The decomposition of wheat straw is known to be closely related to the nitrogen concentration and or carbon:nitrogen (C:N) ratio, as reported by Janzen and Kucey (1988). Additionally, Ehaliotis et al. (1998) found that the decomposition rate of residues with high C:N ratio, such as those of wheat, can be increased by including crops with low C:N ratio in the cropping sequence. The use of rotations enables microbial decomposers to utilize nitrogen from one residue to decompose other residues present. Young wheat plants have a lower C:N ratio than wheat residue where most of the N was already transported to the harvested grains.

It is therefore likely that the large biomass of young volunteer wheat plants, with lower C:N ratio than wheat residue, has enhanced the decomposition rate of the latter even in the fallow treatments. Further studies are required to demonstrate that the incorporation of volunteer wheat plants before winter as a green manure can accelerate wheat residue decomposition and increase the activity of F. graminearum antagonists.

The observed reduction in the proportion of F. graminearum of the total Fusarium species recovered from Fusarium-colonized nodes is in agreement with the findings of previous studies where other species, including F. equiseti, F. oxysporum, F. solani, and F. sporotrichioides, are reported to have greater saprophytic ability than F. graminearum (Burgess, 1981; Garret, 1970; Ocamb, 1991; Pereyra and Dill-Macky, 2004). The reduction in the proportion of F. graminearum-colonized nodes recovered from Fusarium-colonized nodes was consistently greater in green manure-treated soils, compared to fallow-treated soils, in all seven experiments in this study. It was in green manure-treated soils that competition among microorganisms was likely to have been greatest and thus where competition would likely have had the greatest impact on less competitive microbes, such as F. graminearum.

Although we were able to demonstrate an impact of green manures on the soil microbial community and on F. graminearum survival, significant correlations between these parameters were not found when experiments were analyzed individually. Significant correlations were found in the combined analysis across all seven experiments. A significant negative correlation between streptomycete densities or F. graminearum antagonist densities and the survival of Fusarium or the residue recovered indicates a beneficial effect of high streptomycete densities on reducing Fusarium survival. The lack of a stronger correlation between F. graminearum antagonist densities and F. graminearum survival, along with the reduction in bacterial density, suggest that the effects of a single green manure cycle on the soil bacterial community may be weak and transitory. The beneficial effects of green manures on the soil microflora might be enhanced by the incorporation of green manures in succession as reported by Cookson et al. (1998) and Workneh and van Bruggen (1994). Multiple green manure cycles are likely to produce more substantial and sustained changes to the soil microbial community structure and pathogen inhibitory activity (Davis et al., 1996; Wiggins and Kinkel, 2005a). Further investigation is necessary to confirm this hypothesis.

The small biomass of the green manures incorporated in field experiments following a short growing season is perhaps the greatest limiting factor to the inclusion of green manures into cropping systems in Minnesota. The lack of adaptation of buckwheat for late summer planting and the high density of volunteer wheat plants in the field limited our ability to assess the impact of green manures on soil microbial populations in the field. A green manure species better adapted to late summer planting may be required to optimize the potential of green manures to be effective in Minnesota.

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Participation Summary

Educational & Outreach Activities

Participation Summary

Education/outreach description:

PEREZ, C.; DILL-MACKY, R.; KINKEL, L. 2004. Use of Green Manures to Inhibit Fusarium graminearum on Wheat Residues. Proceeding of 2nd International Symposium on Fusarium Head Blight. Incorporating the 8th European Fusarium Seminar. Orlando, FL, USA. 11-15 December, 2004. p. 364.

PÉREZ, C., DILL-MACKY, R., AND KINKEL, L. 2005. Evaluating the potential for green manures to enhance indigenous soil antagonists and reduce inoculum of Fusarium graminearum in wheat residues. Phytopathology 95:S82. APS Annual Meeting, July 30-August 3, 2005, Austin, Texas, USA.

PEREZ, C. 2005. Use of green manures to reduce Fusarium graminearum survival in wheat residues. Master Thesis. Department of Plant Pathology. University of Minnesota. 117 p.

PEREZ, C.; DILL-MACKY, R.; KINKEL, L. Use of green manures to reduce Fusarium graminearum survival in wheat residues. Being sent to Plant and Soil.

Project Outcomes

Project outcomes:

This study is the first report to examine the use of green manures to manage FHB. The results support the use of green manures to increase the population of indigenous soil microorganisms with a capacity to antagonize the survival of F. graminearum in wheat residue. A directional selection to increase the indigenous population of biocontrol agents in soil may be achieved by including green manures in the crop sequence. Further field studies are required to develop strategies for reducing the risk of FHB epidemics through the use of green manures.

Despite the complexity of the soil environment, our results indicate a potential tool to manage the indigenous microbial populations and the diversity of those populations. In addition to their benefits to soil organic matter and nutrients, improving soil structure, and contributing to weed and erosion control (Abdallahi and N’Dayegamiye, 2000; Blackshaw et al., 2001), green manures may be considered in the management of some plant diseases. The findings of this study may be used as a model to examine the survival of F. graminearum in small grain residues, and for other pathosystems in which soilborne antagonists influence the densities and activities of plant pathogens.

Since this is the first report using green manures to manage FHB, further investigation is needed before giving an estimation of benefit on the sustainability of wheat growers. Although some adjustments of this approach is required (e.g. adaptation of green manures for late planting dates), we can anticipate that making this technology more effective, a reduction of the risk of severe FHB epidemics is expected, contributing to the sustainability of the production system.

Economic Analysis

Further investigation is required before making an economic analysis. Seeds cost and tillage (green manure seeding and incorporation) are the only additional costs when using green manures.

Farmer Adoption

Further investigation is required before being adopted by growers. Repetition of the experiment under different year-conditions and technology adjustments are needed. This is a new approach to controlling FHB and only one year of experimentation is not enough to generate a new technology. However, this research allowed exploring the potential of the use of green manures to reduce FHB inoculum, and identify some limiting factor when using this technology.

Green manure species more adapted to late summer planting dates in Minnesota have to be evaluated since biomass production of buckwheat and sorghum-sudangrass showed to be not sufficient stimulating microbial populations in soil under field conditions.

Recommendations:

Areas needing additional study

As mentioned above, experiment replication and new green manure species evaluation is warranted since greenhouse experiments suggest that the use of green manures can enhance populations of indigenous soil microorganisms antagonistic to the survival of F. graminearum in wheat residue.

A green manure species better adapted to late summer planting may be required to optimize the potential of green manures to be effective in Minnesota.

Any opinions, findings, conclusions, or recommendations expressed in this publication are those of the author(s) and do not necessarily reflect the view of the U.S. Department of Agriculture or SARE.