© 2004 Plant Management Network.
Resistance to Azoxystrobin in the Gummy Stem Blight Pathogen Documented in Georgia
Katherine L. Stevenson, David B. Langston, Jr., and Kenneth W. Seebold, Department of Plant Pathology, University of Georgia, Tifton 31793
Stevenson, K. L. Langston, D. B., Jr., and Seebold, K. W. 2004. Resistance to azoxystrobin in the gummy stem blight pathogen documented in Georgia. Online. Plant Health Progress doi:10.1094/PHP-2004-1207-01-RS.
Gummy stem blight, caused by the fungus Didymella bryoniae, is the most destructive disease of watermelon in Georgia and in many other watermelon producing areas of the U.S. The QoI fungicide azoxystrobin has been used for gummy stem blight control in Georgia since 1998. As early as 1999, reduced control of gummy stem blight with azoxystrobin was noted in several research sites and commercial fields of cucumber and watermelon in Georgia. Isolates from several of these fields were later confirmed to be resistant to azoxystrobin. To determine how widespread the resistance problem was in Georgia, extensive surveys of watermelon and muskmelon fields and transplant houses were conducted in 2001 and 2002 to determine the frequency of azoxystrobin-resistant isolates in populations of D. bryoniae. Of the 272 isolates collected in 2001, 247 (91%) were resistant to azoxystrobin. In 2002, 82% of the 170 isolates collected were resistant to azoxystrobin, and of the 40 isolates collected from watermelon transplants, all but one were resistant to azoxystrobin, suggesting that resistant isolates in the field may have originated from seed or transplants. Georgia melon growers are now advised to use alternative fungicides that are chemically unrelated to the QoIs for gummy stem blight control.
Gummy stem blight, caused by the fungus Didymella bryoniae (Auersw.) Rehm, is the most widespread and destructive disease of watermelon in Georgia and in many other watermelon-producing areas of the U.S. (Fig. 1). Although watermelon suffers the greatest losses from gummy stem blight, severe epidemics are observed in cucumber and muskmelon each year. Management options for this disease are crop rotation, deep turning to bury diseased tissue, avoiding irrigation that prolongs leaf wetness, and preventive fungicide applications. Of these management options, application of preventive fungicides is the most effective. Fungicides labeled for control of gummy stem blight include: ethylenebisdithiocarbamates (EBDCs such as Dithane, Maneb, Manzate, and Penncozeb); chlorothalonil (Bravo, Echo, Equus); thiophanate-methyl (Topsin M); azoxystrobin (Quadris), a fungicide in the QoI class of chemistry; and a new carboximide fungicide, boscalid, which was labeled for cucurbits in July of 2003. Benomyl, or thiophanate-methyl tank-mixed with EBDCs and alternated with chlorothalonil products, provided good control of gummy stem blight until resistance to the benzimidazoles was observed in the early 1990s (10). Chlorothalonil products have shown good efficacy on gummy stem blight but are not used because they have been implicated in causing phytotoxicity to mature watermelon rinds, as indicated by a warning on the product label. Azoxystrobin provided excellent control of gummy stem blight in the early 1990s (1,9,15,16,22) and was granted Section 18 emergency exemption status in Georgia in 1997 and 1998 specifically for gummy stem blight control. A full Section 3 national label was granted for azoxystrobin use on cucurbit crops in March of 1999, which led to the widespread and routine use of the fungicide to control a broad spectrum of foliar cucurbit diseases. However, compared to previous reports of disease control with azoxystrobin (22), reduced efficacy of azoxystrobin on gummy stem blight was first observed in Georgia as early as 1999 in watermelon field trials (13) and commercial watermelon fields treated with Quadris. Isolates of the pathogen collected in 2000 from watermelon fields in Delaware, Maryland, and Georgia, where disease control was unsatisfactory, were confirmed to be resistant to azoxystrobin in in vitro laboratory assays (17). In 2001 and 2002, extensive surveys were conducted to determine the frequency of azoxystrobin-resistant isolates in commercial watermelon fields in Georgia. Preliminary reports of these results have been published (20,21).
Sample Collection and Fungal Isolations
Isolates of the fungus were obtained from samples of infected leaves, stems, or seedlings of watermelon and muskmelon showing typical symptoms of gummy stem blight (Figs. 2 and 3). In 2001, samples were collected from 26 commercial watermelon fields and research sites in 13 counties in Georgia, as the disease appeared during June through October. In 2002, isolates of the fungus were obtained from infected watermelon and muskmelon during April through October from transplant houses, commercial fields, and research sites from 15 different locations in Georgia, representing at least six different counties. Counties in Georgia where samples were collected in 2001 and 2002 are shown in Figure 4.
Portions of infected tissue were surface disinfested in 10% household bleach, rinsed in sterile distilled water and placed on ¼-strength potato dextrose agar (QPDA). Cultures were incubated at room temperature (23 to 25°C) for up to 1 week until visible characteristic growth of D. bryoniae was observed. Plugs of agar containing mycelium of the fungus were transferred to fresh QPDA and incubated at room temperature under continuous fluorescent light for 2 weeks to encourage sporulation. Three milliliters of sterile distilled water was added to each culture and the surface of the mycelium was gently scraped to release conidia. One drop of the conidial suspension was spread across the surface of QPDA in a petri dish and incubated at room temperature overnight. A single germinated conidium from each culture was transferred to fresh QPDA and incubated at room temperature for 2 weeks to obtain monoconidial isolates for fungicide sensitivity assays.
Fungicide Sensitivity Assays
Sensitivity of each isolate to azoxystrobin was determined using a conidial germination assay on 4% water agar medium amended with azoxystrobin at concentrations of 0.0001, 0.001, 0.003, 0.01, 0.03, 0.1, 0.3, 1.0, 3.0, or 10 µg a.i./ml, or nonamended (0 µg a.i./ml). Technical grade azoxystrobin (Syngenta Crop Protection, Greensboro, NC) was dissolved in acetone, serially diluted to the appropriate concentration, and added to autoclaved water agar cooled to 60°C, such that the concentration of acetone was 0.1% (v/v) in all treatments. The medium was also amended with 100 µg/ml salicylhydroxamic acid (SHAM) to inhibit an alternative respiratory pathway in the fungus that can interfere with the activity of the fungicide.
Each monoconidial isolate was transferred to four petri dishes of QPDA and incubated at room temperature under continuous fluorescent light to encourage sporulation. Conidial suspensions of each isolate were prepared by flooding each culture with 3 ml sterile distilled water and gently scraping the surface of the mycelium to release conidia. The suspensions from all 4 cultures were combined and filtered through 2 layers of sterile cheesecloth to remove mycelial fragments. The suspensions were centrifuged at 5,000 rpm for 10 min to concentrate the conidia and then re-suspended in 5 ml of sterile water. The final concentration of conidia ranged from 1.0 × 105 to 1.5 × 106. Twenty-five microliters of each conidial suspension were transferred to fungicide-amended or nonamended medium in small petri dishes (60 × 15 mm). Two replicate dishes of each fungicide concentration and isolate combination were prepared. After 24 h (in 2001) or 48 h (in 2002) of incubation at room temperature, 50 conidia per dish were examined microscopically and the percentage of germinated conidia was recorded. A conidium was considered germinated if the length of the germ tube was equal to or greater than half the length of the conidium. Relative germination (RG) was calculated as the percentage germination on fungicide-amended medium divided by the percentage germination of the same isolate on medium without fungicide. Percent inhibition (100 minus RG) was converted to a proportion, probit-transformed, and linearly regressed on log10-transformed fungicide concentration. Fungicide sensitivity for each isolate was expressed as the EC50 value (the fungicide concentration that inhibits spore germination by 50% relative to the control), estimated from linear regressions. As reported in the previous study (17), an isolate was considered resistant to azoxystrobin if the EC50 value was greater than 10 µg /ml.
In both years, three different types of dose-response relationships were observed (Fig. 5). A relatively small proportion of the isolates in both years showed a typical S-shaped dose-response relationship between germination inhibition and fungicide concentration, with estimated EC50 values less than 10 µg/ml; these isolates were considered sensitive to the fungicide. Other isolates showed some dose response, but EC50 values could not be accurately estimated because germination was not inhibited by more than 50%, even on the highest concentration tested (10 µg/ml). The EC50 values of isolates in this group clearly exceeded 10 µg/ml and were considered to be resistant to azoxystrobin. A third group of isolates showed no dose-response to the fungicide; germination was not significantly inhibited, even at the highest fungicide concentration tested. This group of isolates was considered to be highly resistant to azoxystrobin.
A total of 272 isolates from 26 fields in 13 counties in Georgia was assayed in 2001 for sensitivity to azoxystrobin. The frequencies of resistant isolates in each field sampled in 2001 are shown in Table 1. Of the 272 isolates tested, 247 isolates (91%) were found to be resistant to azoxystrobin based on the spore germination assay. In contrast to the 2001 season, gummy stem blight was not as severe in 2002. As a result of lower disease incidence throughout the state, only 170 isolates were collected and assayed for sensitivity to azoxystrobin in 2002. The frequencies of resistant isolates in each field sampled in 2002 are shown in Table 2. Of the 170 isolates collected in 2002, 82% were resistant to azoxystrobin. Of the 40 isolates collected from watermelon transplants, all but one isolate were resistant to azoxystrobin. Interestingly, the frequency of resistant isolates ranged from 57 to 64% in the samples from border rows at research sites where fungicide trials had been conducted for many years and included tests of many different fungicides and application schedules. The frequency of resistant isolates was considerably higher (93 to 100%) in all but one of the samples from commercial fields in 2002. A relatively small number of sensitive isolates was detected in both years (25 in 2001 and 31 in 2002). The minimum EC50 values of these sensitive isolates were 0.021 µg/ml and 0.010 µg/ml and the geometric mean EC50 values were 1.476 µg/ml and 0.301 µg/ml, in 2001 and 2002, respectively. Based on results of Fisher's Exact Test (2-tailed, = 0.05), there was no significant association between the number of resistant or sensitive isolates and the use of DMI fungicides during the current season, for isolates collected in both 2001 and 2002.
Table 1. Frequencies of azoxystrobin-resistant isolates of the gummy stem blight pathogen, Didymella bryoniae, in watermelon fields in Georgia sampled in 2001.
y Isolate was considered resistant to azoxystrobin if the EC50 value was >10 µg/ml, based on a conidial germination assay.
z na = no information available on fungicide use.
Table 2. Frequencies of azoxystrobin-resistant isolates of the gummy stem blight pathogen, Didymella bryoniae, in samples of watermelon transplants and field-grown watermelon (W) and muskmelon (M) plants in Georgia in 2002.
y Isolate was considered resistant to azoxystrobin if the EC50 value was >10 µg/ml, based on a conidial germination assay.
z No information available.
Conclusions and Implications for Disease Management
Results of in vitro sensitivity assays conducted in 2001 and 2002 provided evidence of widespread resistance to azoxystrobin in the gummy stem blight pathogen in Georgia. Of the 272 isolates collected in 2001 from 26 fields in 13 counties, 247 (91%) were found to be resistant to azoxystrobin, and of the 170 isolates collected in 2002, 82% were found to be resistant, based on the spore germination assay. It is very difficult to determine with certainty the origin of the azoxystrobin-resistant isolates. However, based on high frequencies of resistant isolates detected in greenhouse transplants and transplanted and direct-seeded field-grown plants, overuse of the product both in the greenhouse and in the field is the suspected cause. Commercial transplant producers in Georgia have indicated that they have used azoxystrobin for years in the greenhouse to suppress diseases before transplants are sold (Langston and Gay, personal communication). Some watermelon growers have also relied too heavily on azoxystrobin to suppress gummy stem blight and other diseases and have made three or more sequential applications in one crop (Langston, personal communication). Another possible source of resistant isolates is inoculum from previous greenhouse crops of infected transplants treated with azoxystrobin. And because D. bryoniae has been isolated from seeds of cucumber, pumpkin, and other cucurbit hosts (3,14), contaminated seed cannot be ruled out as a potential source of resistant pathogen isolates.
The risk of resistance development in pathogen populations treated with QoI fungicides can range from low to high depending on several factors associated with pathogen biology and epidemiology (6). Pathogens with a short generation time, which produce abundant spores that are readily and widely dispersed, are generally associated with greater risk of resistance (2). Indeed, rapid selection of resistance to QoI fungicides in field populations has been documented in several such pathogens, including Magnaporthe grisea (23), Pseudoperonospora cubensis (8), Podosphaera xanthii (=Podosphaera fusca) (8), Blumeria graminis (4), and Mycosphaerella fijiensis (5). In most cases, rapid development of resistance is associated with a single point mutation (G143A) in the cytochrome b gene of mitochondrial DNA in the fungus, which essentially confers immunity to the fungicide (7,11). However, based on recent work with the apple scab pathogen V. inaequalis, mutations other than G143A may cause a more gradual (quantitative) development of resistance to QoIs, and that both types of resistance may develop consecutively in a pathogen population (12). Although the genetic basis for resistance in D. bryoniae was not determined in this study, the rapid development and high level of resistance observed in the isolates we collected are consistent with the G143A mutation documented in other plant pathogens.
Research on QoI resistance in the apple scab pathogen suggests that if initial shifts toward reduced sensitivity develop gradually, in a quantitative fashion, then rotation of QoIs with unrelated fungicides may restore satisfactory levels of disease control by reducing resistant populations before they can reproduce and spread (12). Unfortunately, however, rotation with unrelated fungicides is not likely to restore a satisfactory level of disease control if resistance is the result of the G143A mutation (12). Therefore, to protect their crops, melon growers in Georgia are advised to rely exclusively on alternative fungicides that are chemically unrelated to the QoIs, rather than alternations or tank mixtures, and eliminate any potential sources of inoculum to reduce severity of gummy stem blight epidemics. Mancozeb and chlorothalonil products both suppress gummy stem blight to some degree. Mancozeb products alone are usually marginally effective and chlorothalonil products have been associated with a rind burn when applied within 2 weeks of harvest. A pre-packaged mixture of the new carboximide fungicide boscalid and the QoI, pyraclostrobin was labeled for cucurbits in July of 2003 and has shown very good efficacy against gummy stem blight in field trials (18,19). Like the QoIs, carboximides also inhibit fungal respiration and are site-specific, but the target site is different from that of the QoIs, so cross-resistance between members of these two groups is unlikely, but the risk of multiple resistance remains a concern. Seed companies and producers have been advised to limit the use of boscalid and practice recommended fungicide rotations on seed production melons and production fields, respectively, to reduce the exposure of the fungus to this new product. Transplant growers have been and will continue to be warned of the dangers of using site-specific fungicides on greenhouse transplants. Hopefully, these strategies will help us preserve the efficacy of boscalid for gummy stem blight control.
The authors thank the Georgia Fruit and Vegetable Growers Foundation for partial financial support of this research, and Jason Brock, Doug Denney, and Sydney Goddard for technical assistance.
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