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a Dep. of Crop and Soil Sci., Washington State Univ., Pullman, WA 99164-6420
b Program in Statistics, Washington State Univ., Pullman, WA 99164-3144
c Dep. of Plant Pathology, Washington State Univ., Pullman, WA 99164-6430
d Dep. of Botany and Plant Pathology, Oregon State Univ., Columbia Basin Agric. Res. Center, Pendleton, OR 97801-0370
* Corresponding author (kidwell{at}mail.wsu.edu)
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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Abbreviations: AG, anastomosis group • CE, controlled environment • PDA, potato dextrose agar • PNW, Pacific Northwest
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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Attempts at grain production via direct-seeding in the 1970s in the PNW failed due to problems associated with increased fertilizer requirements, rodent damage, and increased pest and disease pressure (Papendick and Miller, 1977; Cook, 1991). Although direct-seed systems have evolved through the years to minimize these risks, root diseases have resulted in tremendous yield and crop quality losses when small grains are direct-seeded into standing stubble of barley (Hordeum vulgare L.) or wheat (Wiese, 1987; Smiley et al., 1989; Cook, 1990, 1991; Cook and Haglund, 1991; Smiley, 1996).
Rhizoctonia root rot, caused by the fungus R. solani AG-8, has been identified as an important, yield-limiting disease in direct-seed systems (MacNish 1985, Rovira, 1986; Weller et al., 1986; Pumphrey et al., 1987; Smiley and Uddin, 1993). Rhizoctonia root rot causes brown, sunken lesions in the root cortex, and serious infection leads to severance of the root, creating what is known as the spear tip symptom in roots. Under acute disease pressure, plants are stunted, creating bare patches in the field that can severely limit grain yield (Pumphrey et al., 1987; Cotterill, 1990; Smiley, 1996).
Increased severity of Rhizoctonia root rot in direct-seed systems has been linked to the presence of volunteers and weeds allowed to grow in the field between harvest and planting, referred to as the green bridge, that can maintain or increase inoculum potential of many plant pathogens, particularly R. solani AG-8 (Smiley et al., 1992). Wheat or barley seeded directly into stubble containing volunteer crop plants or weeds sprayed only 2 to 3 d earlier with glyphosate (phosphonomethylaminoacetic acid) commonly develop severe Rhizoctonia root rot (MacNish, 1985; Pumphrey et al., 1987; Roget et al., 1987). Waiting 2 to 3 wk between preplant glyphosate application and seeding significantly decreases the risk of serious crop damage from Rhizoctonia root rot (Roget et al., 1987; Smiley et al., 1992).
If direct-seed systems are to succeed in the PNW, genotypes adapted to reduced tillage or no-till environments, and associated diseases, must be identified or developed. Before 1995, little research had been conducted in the PNW to investigate differences in response of spring wheat cultivars to conventional vs. direct-seed production systems. Field trial evaluations conducted in Eastern Washington revealed shifts in grain yield rankings among spring wheat and spring barley cultivars under severe pressure from Rhizoctonia root rot, depending on location and management system, suggesting that different levels of avoidance, escape, recovery from, or tolerance to Rhizoctonia root rot may exist among spring cereal cultivars (Kidwell et al., 1998).
The objectives of this study were to (i) determine whether current spring wheat cultivars and advanced experimental breeding genotypes vary in levels of susceptibility to Rhizoctonia root rot in inoculated field trials; and (ii) evaluate whether disease ratings obtained in growth chamber evaluations are predictive of, and consistent with, disease ratings in the field.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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Field Design
A field nursery for disease screening was established at the WSU Spillman Research Farm near Pullman, WA, to assess the response of spring wheat cultivars to R. solani AG-8 in crop years 2000 and 2001. The soil at this site is a Palouse silt loam (fine-silty, mixed, superactive, mesic Pachic Ultic Haploxerolls) (Young et al., 1994). The field had been direct-seeded to spring wheat for two consecutive years (1998 and 1999) before initiation of this study. The field was representative of other Palouse fields that are managed in continuous wheat, where low levels of the pathogens responsible for take-all [caused by Gaeumannomyces graminis (Sacc.) Arx & D. Olivier var. tritici], Pythium (caused by Pythium spp.), and Rhizoctonia root rots were assumed to be present in the soil (Cook et al., 2002). No disease epidemics were apparent within this field before initiation of this study.
The experimental design was a split plot consisting of individual genotypes, with four replicates per genotype, as the whole-plot factor, and two levels (high and low) of R. solani AG-8 inoculum as the subplot factor. The high-inoculum treatment was created by planting 2.4-m-wide drill strips of ‘Madsen’ winter wheat at 133 kg ha-1 the full length of the experimental site in the fall, mixed 3:1 (w/w) with oat grains colonized with R. solani AG8 C-1 (Weller et al., 1986). Each drill strip consisted of eight rows spaced 30 cm apart. In both years, fertilizer was added as a split application with a portion of the N applied in the fall, and the remainder applied in the spring. In 2000, fertilizer was deep-banded below the seed at a rate of 97:8:8 kg ha-1 (N:P:S). Inoculated strips with fall fertilization were alternated with 2.4-m-wide drill strips that received the same deep-band application of fertilizer, but not the mixture of inoculum and winter wheat seed, to create the low-inoculum treatment. The custom-built drill used to establish the treatment is described elsewhere (Cook and Haglund, 1991; Cook, 2000).
Twenty-seven days before planting in the spring of 2000, the drill strips designated as low-inoculum were sprayed with glyphosate at 1.7 kg ha-1 to eliminate volunteer plants. Ten days before planting, the entire plot area was sprayed with glyphosate for weed eradication, and to eliminate the fall sown winter wheat. On 25 April 2000, the entire nursery was fertilized at a rate of 55:11:11 kg ha-1 (N:P:S), using the same drill used for fall planting to inject the fertilizer 10 to 12 cm below the soil surface. Spring wheat genotypes were planted 2 d later using a four-row, double disk cone seeder, with row spacings of 30 cm to match the spacings of the drill used to establish the inoculum treatments the previous fall, and to fertilize in both the fall and spring. Four rows of each genotype were planted side-by-side into the high- and low-inoculum portion of each drill strip, respectively.
The 2001 field evaluations were performed in a similar manner with the following modifications. Replication number was increased from four to six in 2001. Drill strips of fall-seeded (27 Oct. 2000) Madsen winter wheat, mixed 3:1 (w/w) with additional R. solani AG-8 oat grain inoculum, were superimposed on the green bridge drill strips from crop year 2000. Fertilizer was deep banded, at a rate of 83:11:11 kg ha-1 (N:P:S), using the custom-built drill described above. Again, low-inoculum drill strips received fertilizer only. Glyphosate was sprayed at a rate of 1.7 kg ha-1 on the low-inoculum treatment for preseason weed eradication 49 d before planting. Six days before planting, the entire nursery was sprayed with glyphosate at a rate of 1.7 kg ha-1. Each trial entry was seeded at a rate of 133 kg ha-1 into 4.5-m2 plots using the eight-row custom-built cone seeder, where four rows were planted into the high-inoculum and the adjacent low-inoculum subplots simultaneously. Additional fertilizer, at a rate of 83:11:11 kg ha-1 (N:P:S), was deep-banded below the seed at the time of planting. Thirty-eight days after planting, the field was sprayed for weeds with a combination of bromoxynil (3,5-Dibromo-4-hydroxybenzonitrile; Buctril; 1.1 kg ha-1; Aventis, Strasbourg, France), fenoxaprop [(ethyl 2-(4-((6-chloro-2-benzoxazolyl)oxy)phenoxy)propanoate; Puma; 0.7 kg ha-1; Aventis, Strasbourg, France], and MCPA amine (2-methyl-4-chlorophenoxyacetic acid; 1.1 kg ha-1).
Root Disease Measurements
The disease severity ratings on the crown and seminal roots were assessed at Haun Growth Stage 6-8 (6 to 8 fully extended leaves on the main stem) (Klepper et al., 1983). Fifteen plants were randomly collected from each plot for disease rating (Smiley, 1996). Roots were washed with a high power stream of water, and then the roots were scored for disease reaction to R. solani AG-8 based on the following scale: 0 = no lesions evident; 1 = <50% roots with a single brown sunken lesion; 2 = <50% of roots each with a few lesions; 3 = >50% roots each with one or more lesions; 4 = <50% of roots with lesions within 1 cm from the seed; 5 = >50% of roots with brown sunken lesions within 1 cm from the seed; 6 = >50% roots severed by lesions within 3 cm; 7 = >50% roots severed by lesions within 1 cm from the seed; and 8 = all roots completely severed within 1 cm of the seed (Kim et al., 1997).
Plant Characteristic and Grain Quality Assessment
Plant height for each plot was obtained near maturity by calculating the average height of 10 randomly selected primary tillers within each subplot. Heading date, measured in calendar days from January 1, was evaluated on a daily basis as plants approached anthesis. On maturity, the grain was harvested from all four rows within each subplot in 2000 with a Wintersteiger plot combine (Wintersteiger Co., Salt Lake City, UT) and in 2001 with a two-row Suzie harvest-binder (Suzue Manufacturing, Japan) followed by threshing with a Vogel thresher (Bill's Welding, Pullman, WA). Grain was weighed to obtain yield estimates based on 14% moisture. After cleaning the grain free of debris, test weights were determined using a Seedburo filling hopper and a 667-cm3 test weight cup (Seedburo Equipment Co., Chicago, IL). Whole grain protein concentration and moisture levels of grain samples were determined using an Infratec infrared whole grain protein analyzer (FOSS North America, Eden Prairie, MN).
Growth Chamber Analyses
Each genotype also was evaluated for disease reaction to R. solani AG-8 as seedlings in the growth chamber. Two treatments were used: (i) pasteurized soil (60°C moist heat for 30 min) infested with ground oat grain inoculum of R. solani AG-8 isolate C-1; and, (ii) pasteurized soil only. A randomized complete block design with each of two chambers representing separate blocks was used for these tests. Five replicates of each genotype per treatment were randomly distributed within each block. Humidity was maintained at 95% to limit plant transpiration and evaporative water losses from the soil. Growth chambers were programmed with a 14-h photoperiod, with daytime and nighttime temperatures of 23 and 11°C, respectively (Smiley and Uddin, 1993).
The infested treatment was prepared by mixing 0.5 g of ground oat grain inoculum into 1 kg of pasteurized Thatuna silt loam in a standard garbage bag by manual agitation. Plastic tubes (25-cm cell depth; 5-cm cell diameter D40 Deepots, (Stuewe and Sons, Inc., Corvallis, OR), plugged at the bottom with a piece of paper toweling, were filled with 300 mL of medium vermiculite to aid in aeration from the bottom of the tube. Forty grams of the soil/inoculum mixture was placed on top of the vermiculite layer, watered to near saturation (Ogoshi et al., 1990) and incubated, undisturbed, for 1 wk to allow mycelium of R. solani AG-8 to colonize the soil (Weller et al., 1986). The same procedure was followed for the noninfested treatment (without inoculum in the soil). At the end of 1 wk, two seeds of each genotype, pregerminated in Petri dishes, were planted in each plastic tube by covering the seedling roots with an additional 1 cm of noninfested pasteurized soil (Ogoshi et al., 1990). After 3 wk, seedlings were removed from the plastic tubes and washed free of soil and debris using a high-power spray. A fine mesh screen was used to support fragile lateral roots. The roots were scored using the 0 to 8 disease scale described for the field evaluation.
To verify the presence of R. solani AG-8 in infected root tissue, randomly selected roots with putative Rhizoctonia incited lesions were washed and, without surface disinfestation, placed on 1/5-strength PDA amended with Rifampicin to inhibit bacterial growth. Hyphal tips from the Rifampicin-amended media were then transferred to 1/5 PDA for further examination and identification (Smiley and Uddin, 1993).
Data Analyses
A combined ANOVA was calculated using the SAS statistical package (version 6.12, SAS Inc., Cary, NC). Agronomic data were analyzed as a split-split-plot design. The main plot was year, the subplot was genotype, and inoculum level was the sub-subplot factor. All factors were fixed in this analysis. Least square means comparisons were used to determine significant differences between genotype means for high- and low-inoculum treatments for each parameter measured at P = 0.05 (Steele et al., 1997). Least square means were estimated using only three of the four replicates for crop year 2000 due to a missing replications for several genotypes as a result of planting errors. Simple linear correlation analysis was conducted for mean disease rating values across replications within each genotype by inoculum level and year (Steele et al., 1997).
Analysis of variance was conducted on disease rating data from growth chamber evaluations using a randomized complete block design in a factorial arrangement with inoculum and cultivar as the main effects in the model (Steele et al., 1997). All effects were fixed in this analysis. Disease ratings from the field were compared with growth chamber disease ratings through simple linear correlation analysis (Steele et al., 1997).
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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In spite of the low disease pressure at both inoculum levels, significant differences between inoculum treatments were detected for grain yield (P < 0.001), plant height (P < 0.001), heading date (P < 0.05), and grain protein concentration (P < 0.05; Table 2). A statistically significant, negative correlation (r = -0.60, P = 0.0001) was detected between grain yield and disease severity ratings. Grain yield averages were always higher (
10%) for the low-inoculum treatment, suggesting that disease pressure, even if slight, had an impact on yield. The low crop yields in R. solani-infested fields is often attributed to an inability of the plant to obtain adequate water and nutrients from the soil (Cook, 1992).
Plants in the high-inoculum treatment were significantly shorter than plants in the low-inoculum treatment (Table 2); however, the correlation between plant height and disease severity was not significant (r = -0.24, P = 0.1). Kirkegaard et al. (1995)(1999) proposed that the relationship between plant stunting and infection by R. solani may result from a hormonal effect, and that even slight R. solani infection may trigger this effect. This response pathway, if similar among all genotypes used in this study, would produce the same restriction on plant growth regardless of the disease ratings.
A significant delay in days to heading was detected between inoculum treatments (LSD0.1; P = 0.05; Table 2), but was only one-tenth of a day and, therefore, is of little agronomic interest (Cook and Veseth, 1991). The level of R. solani inoculum had no effect on test weight; however, a genotype x year interaction was detected for this yield component. Whole grain protein content was moderately affected by R. solani treatments in 2000 (P = 0.05) but not 2001 (Table 2). The largest source of variation for both test weight and grain protein content was attributed to differences among genotypes, indicating, as expected, that genetic variation for these traits exists within this germplasm (Table 2). The genotype x inoculum factor was not significant for any trait, indicating that performance rankings of genotypes were similar across inoculum levels.
Significant yield differences were detected among the market classes evaluated in this study, based on yield averages across genotypes within each market class. Soft white genotypes (6008 kg ha-1) had a significant yield advantage over hard red genotypes (5547 kg ha-1) in the low-inoculum treatments, as expected. Typically, adapted soft white spring cultivars demonstrate a 5 to 10% grain yield advantage compared with hard red spring cultivars, depending on location (Burns et al., 2002). This difference was dramatically reduced in the high-inoculum treatments where the soft white spring genotypes had a yield average of 5225 kg ha-1 compared with a 5048 kg ha-1 average for hard red genotypes. Even with low disease pressure, soft white genotypes appear to be more severely impacted by the presence of R. solani AG-8, based on grain yield potential, than the hard red genotypes.
Even though yield rankings among varieties did not dramatically shift between treatments, several genotypes evaluated in this study appear to be more tolerant of pressure from R. solani AG-8 than others. Significant grain yield differences between the high- and low-inoculum treatments were detected for several soft white varieties, in one or both years (Table 3). The cultivar ‘Zak’ and experimental line ‘WA007883’ exhibited dramatic yield reductions in the high- compared with the low-inoculum treatment. Although Zak was among the highest yielding entries in the low-inoculum trial in both years, it exhibited dramatic grain yield losses in the high-inoculum treatment, even at relatively modest disease pressure. In contrast, the yield potential of six of the nine hard red spring genotypes evaluated in this study did not differ significantly between inoculum treatments in either year (Table 3). The yield potential of the hard white experimental line WA007900 also was stable across treatments and years. Although a relatively small subsample of germplasm from the region was included in this study, preliminary results indicate that the yield potential of hard spring varieties may be more stable under pressure from Rhizoctonia root rot compared with soft white spring germplasm.
Growth Chamber Evaluations
Analysis of variance for root rot ratings in controlled growth chamber evaluations indicated that disease responses among genotypes did not differ (Table 4). No variation in response to Rhizoctonia root rot was detected among spring wheat genotypes in these evaluations.
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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Although the germplasm evaluated apparently has no physiological resistance to Rhizoctonia root rot based on the growth chamber evaluations, differences in disease tolerance based on field performance was observed among genotypes (Table 3). Further studies directed at identifying specific anatomical or physiological differences between genotypes may be necessary to determine if some genotypes are better able to withstand or recover from Rhizoctonia root rot than others.
Neate (1989) also reported that little variation for resistance to R. solani exists among cultivated cereal genotypes, including wheat, and that identification of tolerant or resistant genotypes may be hindered by season-to-season and field-to-field variability in cultivar performance under pressure from Rhizoctonia root rot. The lack of variation for disease reaction, based on disease severity ratings, among the germplasm evaluated in this study suggests that other potential sources of genetic resistance must be examined. A wider survey of germplasm, including adapted germplasm, synthetics, and if necessary, additional secondary and tertiary gene pool members, may aid in identification of sources of genetic variation for disease reaction to R. solani.
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ACKNOWLEDGMENTS |
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Received for publication February 11, 2002.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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This article has been cited by other articles:
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  J. D. Smith, K. K. Kidwell, M. A. Evans, R. J. Cook, and R. W. Smiley Evaluation of Spring Cereal Grains and Wild Triticum Germplasm for Resistance to Rhizoctonia solani AG-8 Crop Sci., March 1, 2003; 43(2): 701 - 709. [Abstract] [Full Text] [PDF]
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