Stephen J. Toth, Jr. and Thomas A. Melton, editors

The information and recommendations in this newsletter are applicable to North Carolina and may not apply in other areas.

Date: September 19, 2003

From: Dr. Frank Louws, Dr. Leonor Leandro, Rosemary Schwegel, and Jim Driver, Department of Plant Pathology

Anthracnose problems have been widespread this year in plug operations. We have isolated the pathogen from many different sites and have consistently identified Colletotrichum acutatum. This pathogen is particularly aggressive in the spring when it causes ripe fruit rot. Clearly, it also is aggressive on young green and succulent tissue, causing lesions on leaf petioles and plant collapse. In the last 2 years, we have been able to isolate it from infected roots also.

Can you treat plug trays and reduce disease levels? To determine the effect of C. acutatum on young plug plants and to determine if fungicides could have a curative effect, we conducted a trial last fall. Four week old plug plants (in 50 cell trays) were inoculated with a spore suspension of C. acutatum and covered with plastic for 48 hrs (to allow for infection). Several trays were not inoculated (non-inoculated controls). Seven days later, plants were sprayed until runoff and trays were drenched from above until soil saturation with a fungicide solution. Plants were field set the following morning. No additional fungicides were applied until 25 March 2003. Data on plant mortality, vigor, and yield were collected. Young and old leaves were collected in February and assayed for the presence of C. acutatum.

RESULTS: Lesions began to form on leaf petioles 7 days after inoculation. The inoculation level was moderate. Some plant mortality occurred soon after planting, and by the spring differences were noted (Figure 1). Captan treatments increased plant mortality. C. acutatum alone (with no treatments) did not significantly reduce plant stand compared to non-inoculated controls.

The pathogen caused stunting of inoculated plants (Figure 2). All plants inoculated with C. acutatum were distinctly stunted (6 Mar 2003) compared to plants not inoculated. The plant growth in each plot of inoculated plants was variable i.e. some plants were larger than others. Therefore, we used a weighted scale using values from 1 to 6, rating each plant and then weighting this rating based on the number of plants in each category. No fungicide “cured” the plants from this stunting symptom and Captan tended to increase stunting problems.

Ten to 12 newly emerged and old leaves (that emerged after inoculation) were collected 6 Feb. Leaves were surface sterilized and then treated with paraquat (GramoxoneTM) to kill plant tissue, and sprayed with RovralTM to inhibit saprophyte growth. The leaves were incubated in moist chambers to allow C. acutatum to grow out, if present in the leaf. On average 27% and 45% of the new (range = 4% to 50%) and old leaves (range = 10% to 64%), respectively, were infected by the pathogen. Non-inoculated controls tended to have the lowest number of infected leaves. These data suggest that the pathogen had successfully spread from plot to plot and colonized new leaves during the winter months.

RECOMMENDATIONS: Based on this trial, recommendations to treat infected plug plants cannot be provided. The fungicides in this trial offered no advantage. (Fall fungicide applications in the field were not evaluated in this trial). We have good data that demonstrate the benefits of spring fungicide programs and therefore suggest it may be best to save all fungicide uses for spring applications.

Can you cull out infected plug plants and plant the rest? Last year, many growers culled out infected and surrounding trays, planting the remainder of plants. This appears to be a successful strategy for growers who cannot secure an alternative source of plants. The main factor that determines the success of culling is the level of disease in the plug trays. Early problems diagnosed in August of 2003 were severe. We mapped “hot spots” in an area of 60,000 trays (Figure 3). This map represents hotspots where plant symptoms such as stunting and desiccation were clearly visible.

The site also had 21 sprinklers evenly distributed throughout the area. We collected 5 petioles from four sites around each sprinkler (N, S, E, and W). These petioles were placed in sterile tubes, then taken to the lab. The tissue was frozen (-20 C) for 2-3 hrs and then surface sterilized in a bleach solution. The petioles were incubated for 7 days and scored for presence of C. acutatum using microscopes . Two sites were sampled in a similar fashion. Up to 15% and 7% of the samples at Sites 1 and 2, respectively, had the pathogen present. The majority of this tissue had no symptoms.

RECOMMENDATIONS: Infestation levels as high as those observed at these sampling sites were too high for culling to be an effective control strategy. These growers secured an alternative source of plants. A short while later, at two other grower sites in the mountains, the level of infestation based on the presence of hot spots was lower. However, leaf petioles and (especially) runners with diagnostic lesions were scattered throughout the trays.

There has been some indication that later shipments of tips have not been as heavily infested. In such cases, and given the shortage of plants, the only option a grower may have is to cull out hot spots and surrounding trays. Remember that although surrounding plants may show no symptoms, the pathogen may still be present. In such cases, limit overhead watering once plants are rooted and do not handle plants when wet. Plant sooner rather than later.

Are bare-root plants from the same source a viable alternative? Dr. Poling has provided extensive details on his trip to Canada and the sampling of strawberry tissue. Dr. Poling walked 2 fields of strawberry plants (Field A & B) that were destined for supplying North Carolina. A systematic sampling scheme was developed and 400 leaf petioles from the mother plants were collected from each field (roughly estimated to represent 1.5% of the mother plants). Forty petioles, sampled from each of 10 rows, were placed in ziplock plastic bags and delivered to our lab. One bag was previewed (Field A, Bag 8), based on Dr. Poling’s suggestion, since lesions were present on 2 of the petioles. The lesions were diagnostic for anthracnose and characteristic spores were confirmed in one of the lesions.

All of the bags were placed in the -20 C freezer for 6 hrs in an attempt to kill the green tissue (as we did above). The petioles were then surface sterilized and placed on wire racks in incubation chambers that contained wet paper towels. Such high relative humidity chambers allow the pathogen to grow out and presence of the pathogen can be scored using microscopes. The petioles remained green, even after 6 days. The petiole tissue from the field nursery, unlike those from plug houses, was not susceptible to the freeze-thaw method of desiccation. The samples were then treated using a paraquat technique and scored again after 4 days (13 Sep) and again after 7 days (16 Sep). Most of the petiole tissue remained green. The tissue needs to be killed for the pathogen to grow out. On the tissue that did die, no evidence of C. acutatum was observed.

RECOMMENDATIONS: Field A had symptoms and spores consistent with C. acutatum. There is a chance that the bare-root plants will have anthracnose associated with them. The pathogen will multiply during the winter and early spring – producing more inoculum on the green tissue. Then, with the warm temperatures and favorable wet weather in the spring, flower blight and fruit rot can occur. The vigilant use of fungicides in the spring, according to the schedules we have developed, should help minimize these crop losses.

Should the bare-root plants be dipped in Quadris? Growers do have an option to dip their bare-root plants in Quadris. It is now a labeled use. Research in California and Florida has shown this to be of benefit if the bare-root plants are heavily infected. In one study, bare-root transplants were inoculated with C. acutatum and incubated for 4 days. The transplants were then dipped in fungicide solution for 5 minutes directly before hand-planting into the field and strawberry plants were assessed for mortality, vigor and yield. The use rate for Quadris was 4 oz/100 gal. Treatments were also complemented with a spring fungicide program or not treated further.

Inoculated plants treated with no fungicide had a mortality rate of 83% - 88%. Quadris dip treatments reduced this level to 5% - 15% mortality. Likewise, Quadris dip treatments boosted plant vigor. Quadris dip treatments, combined with a spring fungicide program that included Quadris, Switch and Captan, were among the best in the trial.

RECOMMENDATIONS: If there is strong evidence of a heavy infestation of anthracnose present in the bare-root plants, a Quadris drip treatment could be beneficial for suppression of root and crown rot caused by Colletotrichum species. Use 5-8 fluid oz per 100 gallons. Where practical, wash and remove excess soil prior to dipping; completely immerse planting stock in solution; DO NOT reuse water solution (ideally, each set of plants should be dipped in fresh solution); dip to expose plants for a minimum of 2 to 5 minutes; plant immediately.

There are some concerns and issues. First, the tips with the anthracnose problem have also had a moderate level of angular (bacterial) leaf spot. The question arises: “Will the bare-root plants have the same disease?”. Dip treatments will cause this pathogen to spread from plant to plant. Thus, it would be best, if practical, that fungicide solutions not be reused. Alternatively, growers may wish to explore how they can treat only the roots in the dip process. Removal of free soil has in itself reduced anthracnose problems in Florida and helps to keep the Quadris fungicide active. Workers must be properly protected when dipping and planting. The solution must be properly disposed of.

IN SUMMARY: Anthracnose concerns have been high for 2003, primarily from one nursery source. It appears that tips, especially in early shipments, have had a very high incidence of infection. This high level of disease precludes culling and there is a serious concern about plant establishment and plant vigor if infected plants are used. Fungicide applications to plug trays have not proven useful and no data is available concerning the utility of fall applications in the field. There is considerable data that indicates spring fungicide programs work and should be the focus of anthracnose management. Bare-root plants from the same nursery may have anthracnose but the likelihood of disease development is unknown. Assays were not effective for determining a rough estimate of incidence. If the data suggest a high infestation level in the nursery plants, Quadris root dips could be implemented. Otherwise, growers may choose to focus on good plant establishment and management with a focus on spring fungicide use.

DISCLAIMER:

From: The University Office of Legal Affairs (Aug. 29, 2003)

1. The strawberry specialist inspected fields that are of concern to inspect for proper disease management practices and visual signs of the disease. A representative sample of plants was obtained upon which laboratory examinations were conducted to determine if the disease was possible.

2. By the nature of the disease, it may be hidden at the time of visual and/or laboratory examination, and thus there is some risk that the disease may be present but not detected. Therefore, the risk of a false negative is relatively high.

3. We have done the in-field and laboratory screening to help reduce the risk to growers, but because the risk of false negatives is relatively high, the grower assumes risk for any disease that develops subsequent to the screening.

4. Therefore, our screening process in no way states or implies that plants are free from disease or the risk of developing disease.

Employment and program opportunities are offered to all people regardless of race, color, national origin, sex, age or disability. North Carolina State University, North Carolina A&T State University, U.S. Department of Agriculture, and local governments cooperating.

Last modified on September 22, 2003 by Stephen J. Toth, Jr.