The University of Georgia College of Agricultural & Environmental Sciences
Cooperative Extension Service

Tobacco Disease Control

Paul Bertrand

Crop Rotation
Nitrogen Management
Plant Spacing
Early Stalk and Root Destruction
Varieties
Blue Mold
Black Shank
Brown Spot
Fusarium Wilt
Soreshin
Target Spot
Angular Leafspot
Granville (Bacterial) Wilt
Hollow Stalk (Soft Rot, Barn Rot)
Tobacco Etch Virus (TEV) and Potato Virus Y (PVY, Vein Banding)
Tomato Spotted Wilt Virus (TSWV)
Root-Knot Nematode

Disease control in tobacco involves an integrated program of numerous practices. Useful practices include crop rotation, root and stalk destruction, nitrogen management, plant spacing, varieties, and chemicals. Growers should never allow themselves to fall into the trap of letting chemicals carry the entire disease control burden. Total reliance on chemicals is environmentally unsatisfactory. Annual pressure on chemicals increases chances of control failure due to loss in activity, pathogen build-up, or errors in application. Even where chemicals are effective, disease control can be improved by making use of all available practices. This in turn increases yield, quality and profit. There are a few common diseases that cannot be controlled with chemicals. Table 1 lists common tobacco diseases and practices known to be useful in controlling each one.

Crop Rotation

Crop rotation is a particularly useful practice in dealing with pathogens that live in the soil. Rotation causes pathogen populations to decline naturally in the absence of suitable food sources. The longer tobacco can stay out of a field, the more beneficial the program will be. In general, small grains and forage grasses are the best overall crops to rotate with tobacco. In the case of root-knot nematodes, selection of the best rotational crop depends upon which root-knot species are present. Cotton is a good choice where Javanese and/or peanut root-knot is present, but a poor choice where high levels of Southern root-knot are present. Corn, though always better than continuous tobacco, is not always ideal due to variation in how corn varieties respond to different nematode populations. Soybeans vary greatly in susceptibility to root-knot nematodes and it is fairly easy to select a variety that would be more beneficial in a particular case. As with other diseases, small grains and forage grasses work well. In most cases, even native weeds would be better than continuous tobacco.

Nitrogen Management

Excess nitrogen causes a nightmare of sucker problems and favors several diseases. Use the full amount of nitrogen needed to produce a top quality crop, but not a bit more. Even small amounts of excess nitrogen can result in a significant, usually negative response in tobacco.

Plant Spacing

A field spacing of about 20-22 inches between plants has been shown to produce the best tobacco. Tighter spacing puts more plants per acre and restricts air movement between plants. Restricted air movement slows drying and increases suitable pathogen habitat by favoring both infection and spread of diseases. Increasing plant population by decreasing in-row spacing to compensate for loss from tomato spotted wilt virus is not necessary and is not recommended. Following such a practice would lead to increased sucker control problems every year and increased losses from blue mold, target spot, angular leafspot and hollow stalk in wet years. Reduced plant spacing will also lead to more rapid and extensive spread of tobacco mosaic virus.

Early Stalk and Root Destruction

This practice, also known in North Carolina as R-9-P for reduce nine pests, is very important. Root and stalk destruction as soon as possible after harvest reduces overwintering populations of pathogens infesting these plant parts. The root destruction part of the program also eliminates fall and winter suckers which feed infested roots and serve as sources for build-up of foliar diseases. Blue mold and viruses can carry over during a mild winter in these suckers.

The four steps involved in a good root and stalk destruction program are outlined below:

• Cut stalks into small pieces with a rotary or flail mower as soon as possible after harvest.
• Plow or harrow the stubble immediately after stalk cutting. Be sure to pull roots completely out of the soil.
• Cross harrow the field two to three weeks after Step 2.
• When roots are fully dried out and dead, plant a winter cover crop to reduce soil erosion.

Varieties

Resistant varieties are useful for control of some tobacco diseases. Growers should not rely on the same variety for disease control year after year. Pathogens can adapt through build up of initially minor species or races and in time overcome any variety. This has been clearly demonstrated for black shank and root-knot nematodes. Changing varieties every now and then is a good idea and should slow down development of "new problems." Changing varieties after a pathogen has overcome an old favorite will not change the situation back. Table 4 in the Variety Section of this book lists common tobacco varieties and their disease resistance properties.

Blue Mold

Blue mold caused by Peronospora tabacina is spread by spores carried long distances by wind. Ideal conditions for blue mold would be night temperatures above 50 degrees F, day temperatures about 70 degrees F along with fog, rain or dew to keep leaves wet. Races of blue mold resistant to Ridomil were found in Georgia in 1991. The incidence of Ridomil resistance in the blue mold pathogen population entering Georgia has increased steadily since 1991. Fields should be regularly checked during blue mold favorable weather. Any blue mold found should be reported to your county agent at once.

Long distance spore movements are tracked by the North Carolina Blue Mold Forecast System. This system can give up to 48 hours warning of blue mold spore movement. This information can be accessed via the world wide web at: http://www.ces.ncsu.edu/depts/pp/bluemold/

These forecasts may be used to trigger treatment or intense scouting depending on the level of risk each individual wishes to assume. These forecasts are useful to predict first movement of blue mold into an area. Once blue mold is present, local weather conditions should be used to make further treat/don't treat decisions.

Table 2. Blue Mold Control

Note: Above directions are for dilute sprays. If concentrate sprays are used, adjust rate and volumes proportionally, e.g. for mist blows, use 2X concentrate and ½ the spray volume.

Begin applications when the Blue Mold Advisory states that conditions favor development of blue mold, and before the onset of disease infection. Continue applications on a 5-7 day spray schedule until weather conditions favoring infection and sporulation decrease. Discontinue sprays when and if the threat of blue mold subsides. Restrictions (Field Applications): DO NOT exceed 2.5 lbs/acre per application. DO NOT exceed 10 lb/acre of ACROBAT MZ per season. In flue cured tobacco, DO NOT spray after the appearance of first button, or within 21 days of harvest.

Black Shank

Black Shank is caused by the fungus Phytophthora parasitica var. nicotianae. DO NOT GROW TRANSPLANTS IN ANY FIELD WHERE BLACK SHANK IS KNOWN TO HAVE OCCURRED. METHYL BROMIDE WILL NOT ERADICATE THIS FUNGUS OR GUARANTEE DISEASE FREE TRANSPLANTS. Black shank infection is favored by wet spring weather. Highest losses then occur during dry summer periods as rotted roots fail to keep up with water demands. Typically black shank results in extensive root rotting, pith disking and decomposition, and blackening on the outer surface of the stalk. Resistant varieties show less obvious symptoms in the pith.

USE RESISTANT VARIETIES! No varieties are immune to black shank. Some varieties show a degree of resistance. The degree of resistance shown by any variety will vary from field to field depending on the natural and not predictable variation in the local black shank fungus and how often the variety is planted. The field to field variation in two varieties with moderate black shank resistance is shown in Table 4. Neither of the resistant varieties had been grown in these fields previously. WHEN CHOOSING A RESISTANT VARIETY, KEEP LOCATION TO LOCATION VARIATION IN RESISTANCE IN MIND. The more often a variety is planted, the lower the resistance it will have (Table 5). A Complete listing of variety response to black shank is found in Table 4 of the Variety Section of this book.

BACK-UP ANY RESISTANT VARIETY WITH A SOUND MEFENOXAM PROGRAM(Table 6.)

FOLIAR APPLICATIONS OF RIDOMIL GOLD/ULTRA FLOURISH

FOR BLACK SHANK CONTROL ARE ILLEGAL AND USELESS!

1) Very little mefenoxam is taken up by leaves.

2) Almost none of the mefenoxam taken up by a leaf moves out of that leaf.

3) All mefenoxam movement in plants is upward away from the roots where the black shank fungus enters.

Results of Black Shank on Tobacco

Brown Spot

Brown spot, caused by Alternaria alternata, is a foliar disease that usually develops from mid season to harvest. It begins in the lower leaves and works up the plant. Brown spot is favored by wet weather, excess nitrogen and tight plant spacing. Fungicides are not effective for brown spot control.

Fusarium Wilt

Fusarium wilt, caused by Fusarium oxysporum f.sp. nicotianae, is not a common problem. It is a serious problem where it occurs. Symptoms usually develop on one side of the plant with distinctive leaf yellowing and drying. Peeling the outer bark will reveal brown to black discoloration in the woody stem cylinder. Initially there is little root rot. The best control for Fusarium wilt is to abandon an infested site. Where this is not practical long rotations with forage grasses or small grains are recommended. Root-knot nematodes will make Fusarium wilt much worse. In Fusarium infested sites, treatment with multi-purpose fumigants and RKN resistant varieties are useful.

Soreshin

Soreshin is caused by the fungus Rhizoctonia solani AG-4 and usually develops during the first 4-6 weeks after transplanting. This disease is favored by rough handling transplants and cool, wet weather. Sand blasting stems in wind storms often leads to soreshin. There is no chemical control for soreshin.

Target Spot

Target Spot is frequently seen in plantbeds. It caused damage in Georgia tobacco fields for the first time in 1991. It is caused by races of R. solani in group AG-3 which are different from those causing soreshin.

In plantbeds the symptoms are small brown greasy looking spots that occur primarily in wet areas of the bed. In the field it occurs first on lower leaves and in very wet seasons can move to some extent up the plant. Symptoms begin as small spots similar to what is seen in plantbeds. On field tobacco the spots enlarge, become somewhat circular, light colored, and papery with a target like pattern of concentric bands. Target spot is very difficult to distinguish from brown spot or black shank leafspot by symptoms alone. Target spot is favored by long periods of leaf wetness and continuous moderate temperatures (68 degrees F - 86 degrees F). These are similar to the conditions that favor blue mold. The R. solani races that cause target spot have always been present in our soils. The growing season weather will regulate future occurrence of target spot.

Fair target spot control can be obtained in plantbeds using spray programs in Table 1 in the Transplant Production Section of this book.

Angular Leafspot

Angular leafspot, caused by Pseudomonas syringae pv. tabacina, is a bacterial disease favored by wet weather, excess nitrogen, excess lime (high pH), low topping and tight plant spacing. Any handling of plants in infected fields during wet periods will spread the disease. Streptomycin sprays have been recommended in various sources for angular leafspot control. The value of these sprays is questionable. It is not economical to apply them every year, yet bacterial diseases become very hard to control once they are present in the field. The disease increases during wet weather. A warm, dry period that would dry the field out and allow for spraying usually halts spread of the disease. The program usually recommended is a solution containing 200 ppm Streptomycin applied at 25-35 gallons per acre every 7 days.

Granville (Bacterial) Wilt

Granville wilt, caused by Pseudomonas solanacaerum, is not a common problem in Georgia. However, it is a very serious problem for the growers who have it. Tobacco variety resistance to Granville wilt is given in Table 4. of the Variety Section of this book. Corn, cotton and peanuts are poor choices for rotational crops in fields where Granville Wilt is a problem. Use soybeans, small grains or weeds in these fields. In fields with a known history of Granville wilt use a multi-purpose fumigant such as Chlor-o-pic at 3 gpa or Telone C-17 at 10.5 gpa.

Above ground symptoms of Granville Wilt are virtually identical to those of Fusarium Wilt. In early stages before secondary deterioration begins, internal symptoms are somewhat different. Granville wilt usually shows a general darkening. Fusarium wilt will show dark streaks in the vascular tissue while the pith remains white. The best way to verify Granville wilt is to suspend one end of a stem section in warm (not hot) water for two to 10 minutes. Set the stem section so that one end is about one inch under water, two inches or so above the bottom of the water reservoir. If Granville wilt is the problem, a distinct cloudy fluid or material will usually flow out of the underwater stem end.

Hollow Stalk (Soft Rot, Barn Rot)

These diseases are caused by the bacteria, Erwinia carotovara var. carotovora. Disease is favored by tight plant spacing and wet weather, particularly during topping season. When the disease is present, tops and suckers should not be pulled while the plants are wet. High rates of contact sucker control chemicals can contribute to disease in wet seasons. When disease is present, harvest should be avoided while the leaves are wet. Packing wet infested leaves into boxes or racks can result in further losses from barn rot.

Tobacco Etch Virus (TEV) and Potato Virus Y (PVY, Vein Banding)

TEV and PVY are aphid transmitted viruses. The virus is carried as a contaminant on the mouth parts. The virus is picked up in feeding on infected plants. Virus can be transmitted to a healthy plant if an aphid feeds as briefly as 10 seconds. Nearly all the virus particles the aphid carries are wiped off of its mouth during the first feeding after picking up virus. Insecticide sprays will not help control these or other aphid borne diseases.

Tomato Spotted Wilt Virus (TSWV)

TSWV first appeared in Georgia tobacco in 1986. Leaf symptoms are quite variable and include necrotic banding along and around the main veins, target-like ring spots, leaf twisting with symptoms on only one side of the midrib, and/or general necrosis of bud leaves. Stalk symptoms are also somewhat variable. Early in the season the lower stalk may show a dark, somewhat sunken, area resembling soreshin. Near topping time, parallel black necrotic bands (curved or straight) may be seen moving down the stalk from infected leaves. These bands are different from the usual russet streaks that come from contact sucker control products. Large plants may develop symptoms (stalk and leaf) on one side or the entire stalk may become necrotic causing a rapid wilt, leaf yellowing and death. Near harvest, stalks of infected plants seem to blacken and rapidly deteriorate.

TSWV is spread by thrips. In tobacco the major carrier of TSWV is the tobacco thrips Franklinella fusca. TSWV must be picked up by juvenile thrips feeding on infected plants. TSWV may be spread by both juvenile and adult thrips. In tobacco, adult thrips are believed responsible for most infection.

Weeds provide the source of TSWV. About 30 species of common broad leafed weeds have been found to host TSWV in Georgia. Tobacco thrips acquire the virus in native weed communities and bring it to tobacco as they move about in search of feeding sites. Weeds near tobacco may be more important than weeds further away but thrips are active flyers and move whatever distance is necessary to find suitable habitat.

SPOTTED WILT MANAGEMENT

I. TRANSPLANT DATE

The response to transplant date varies from year to year and from farm to farm in any particular year. No single best time to plant or not plant can be identified. However, in combining the results of all transplant date trials the data shows the risk of most spotted wilt is twice as great in tobacco planted before 7 April as compared to planting after 7 April.

II. ADMIRE 2F

Admire 2F was labeled for TSWV suppression in 1997. Numerous trials have been conducted comparing Admire treated to nontreated plants. TSWV suppression with Admire has ranged from 0-60%. It is reasonable to expect 25-35% TSWV suppression with Admire when it is correctly applied. It has been found that when correctly applied higher rates of Admire give improved TSWV suppression.

HOUSE PLANTS

Apply Admire as a "spray-on rinse-off" tray drench. The rinse off must be complete before product dries on the plants. If treated plants look chalky the product did not rinse off and must be re-applied. Avoid over rinsing. If a milky scum is seen on the float water under trays just after rinse-off the product has been washed through the root medium and should be reapplied. NOTE: Where plants are grown in trays on benches great care must be taken with post treatment watering not to leach the Admire out of the trays.

BARE ROOT PLANTS

Mix Admire 2F directly with transplant water or in a separate front mounted tank where it can be metered into the transplant furrow.

"Spray-on rinse-off" treatments with Admire in plant beds have failed to reduce TSWV.

III. ACTIGARD 50WG

Approval for use of Actigard in plant houses and plant beds has been granted through a third party special local needs registration. Flue cured stabilization will be the third party administrator for this use. Product labels for this use must be requested through flue-cured stabilization. Third party registrations are used in cases where there is some risk of plant damage associated with product use. Under these registrations the user of the product assumes ALL liability.

Spray Actigard over plants in houses or beds 5-7 days before transplanting. Potential for plant injury depends largely on plant size at the time of treatment. The following guidelines are suggested:

PLANT BEDS 0.5oz/32,000 plants.

PLANT HOUSES with 338 cell trays 0.5oz/50,000 plants.

In houses using 242 or 288 cell trays adjust rate between 0.5oz/32,000 and 0.5oz/50,000.

Use Actigard with Admire in a joint program.

CONTROL OF TSWV WITH AN ACTIGARD/ADMIRE PROGRAM

USE ACTIGARD VERY CAREFULLY:

1. DO NOT exceed 0.5oz/31,000 plants.

2. DO NOT treat plants with Actigard a second time.

3. DO NOT hold Actigard treated bed plants more than a day after pulling. Treated plants which are held do not perform as well as fresh plants.

4. Use transplant water.

Even when Actigard is used carefully some plant damage in the form of slow early season growth may occur. How well plants recover depends on field environment the first few weeks after transplanting.

1. Use transplant water.

2. DO NOT plant in dry soul

3. Irrigate before plants become stressed.

The TSWV research project in Georgia has lead to three general conclusions about this disease in tobacco.

• Plant Beds/Plant Houses Are Not a Significant Source of Disease.
• TSWV can occur in plant beds or plant houses, but several observations and research projects have made no connection between plant source and the final incidence of disease.
• Secondary Spread of TSWV Is Not Significant.
• Secondary spread would be due to juveniles feeding on infected plants in the field and furthering disease spread as they move about as infectious adults. Results to date suggest disease arising in this manner is not significant.
• Most Infections That Lead to Symptom Development and Plant Death Occur Shortly after Transplanting.
• The number of plants showing symptoms of TSWV continues to increase throughout the season. This does not mean that the actual infections are occurring continuously. TSWV, in fact, is known to have a variable and often quite long period between infection and symptom development. Research results from insecticide trials, planting date trials, and plotting disease increase over time support the hypothesis that most infections that lead to symptoms occur shortly after transplanting.

Should Tobacco with a High Incidence of Tswv 3-5 Weeks after Transplant Be Reset?

This sounds like a good idea, however, our data from transplant date trials suggest there is some risk involved. The data (Table 6) show that in some years earliest transplanting leads to most disease. In these years, re-setting would be useful. However, it also shows that in other years, latest transplanting leads to most disease. Resetting an early planting with "too much" TSWV these years would make the situation even worse. While early planted tobacco seems to suffer the greatest losses most often, we CANNOT predict how a particular season will turn out.

Root-Knot Nematode

Root-knot nematode (RKN) is the major nematode problem facing Georgia tobacco growers. Control of RKN is becoming more complicated each year. A virtual complete shift to varieties resistant to race 1 & 3 of Southern RKN has resulted in selection of previously minor species and races of RKN and elevating these to major pests. As a result, Javanese, race 2 and/or 4 of Southern and, to a lesser extent, peanut RKN are now major tobacco pests. There are no varieties resistant to these species or races leaving root destruction, crop rotation and chemicals as the only control options. Control of Javanese and peanut RKN must be more thorough than with the traditional Southern RKN because as individuals these species are much more damaging to tobacco.

Root and Stalk destruction should be done thoroughly.

Crop Rotation

In selecting rotational crops the grower must know which RKN species and races are present. For example, cotton is a good rotational crop for Javanese and Peanut RKN, but a poor one for Southern RKN. Soybeans vary widely in their resistance to various RKN species. Choice of best variety depends upon the specific RKN present.

Nematode Sampling

The only way to know what is present is to sample the field. Samples taken in the fall or winter especially following crops other than tobacco are almost worthless. When taking nematode samples, follow the guidelines outlined below:

• Sample during the tobacco growing season (15 June - 15 July).
• Sample moist soil; do not add water to dry samples.
• Sample the entire field.
• Sample problem areas separately from non problem areas.
• Keep samples in a portable cooler between collection and delivery to the county extension office. This will keep the samples from overheating which kills and rots nematodes very quickly.
• Sample early in the week so samples will arrive at the lab in Athens the same week as collected.
• Fill out the nematode sample form completely including tobacco variety information.
• Check tobacco roots for nematode galls. These can be used to determine RKN species present. Knowing which kinds are present is much more useful than knowing how many.
• Sampling crops other than tobacco the year before growing tobacco is generally a waste of time.

Root Examination

Root examination is very useful in evaluating nematode problems. From mid June until mid July, dig up a few stalks at various points in the field and look at the roots for galls. If RKN is present some galls may be seen on any variety regardless of what chemical nematicide is used. However, if root systems are consistently 25% or more galled, some loss is occurring and the control program should be reviewed. Always look at root systems in areas of any field where the tobacco does not grow well. Once the RKN population has been defined, the best rotational crops and chemical nematicide can be chosen.

Root-Knot Nematode Species Identification

The Extension Nematology Lab has technology to enable accurate identification of RKN species. At present there is no charge for the procedure. The procedure is expensive so growers taking advantage of this service are asked to:

• Use it only where reasonable suspicion of a problem exists (numerous galls on RKN resistant varieties).
• Keep good records so as to not continually re-sample fields where problems have been identified. Nematode populations vary with crop history and season, but they do not go away.
• Handle samples for root-knot nematode species identification through the county agent as with conventional nematode samples. Sampling involves mid-June to mid-July collection of plant roots. Samples are to be collected in the following manner.
• Map out the field and dig rather than pull up sample plants.
• Gently shake off most soil.
• Select for testing plants with moderate to severely galled root systems that show NO sign of secondary rotting or gall breakdown. The test requires live female nematodes to be present in the galls.
• Collect specimen plants from several areas in a field. Take a separate sample for each 10 acre block in a field.
• Place galled roots (tops should be cut off) in plastic bags containing moist field soil and store in a cooler. It is not necessary to send the whole root system. Individual roots with galls on them are good enough. Clip roots rather than pulling roots apart to avoid stripping or damaging root galls.DO NOT ADD WATER.
• Do not allow samples to heat or dry out.
• Collect samples early in the week (Monday or Tuesday) so they can be shipped and received in the same week.

Root-Knot Nematode Chemical Control

Resistant Varieties

Root-knot nematode resistance in tobacco is specific for races 1 and 3 of Southern root-knot nematode. No varieties are available with resistance to Javanese, Peanut or races 2 and 4 of Southern root-knot nematode. For a listing of varieties resistant to races 1 and 3 of Southern RKN refer to Table 4 in the Variety Section of this book.

Attention: Pesticide Precautions

1. Observe all directions, restrictions and precautions on pesticide labels. It is dangerous, wasteful and illegal to do otherwise.
2. Store all pesticides in original containers with labels intact and behind locked doors. "Keep pesticides out of the reach of children."
3. Use pesticides at correct label dosage and intervals to avoid illegal residues or injury to plants and animals.
4. Apply pesticides carefully to avoid drift or contamination of non-target areas.
5. Surplus pesticides and containers should be disposed of in accordance with label directions, to contamination of water and other hazards will not result.
6. Follow directions on the pesticide label regarding restrictions as required by State and Federal Laws and Regulations.
7. Avoid any action that may threaten an Endangered Species or its habitat. Your county extension agent can inform you of Endangered Species in your area, help you identify them and, through the Fish and Wildlife Service Field Office, identify actions that may threaten Endangered Species or their habitat.

Trade names are used only for information. The Cooperative Extension Service, the University of Georgia College of Agricultural & Environmental Sciences, does not guarantee or warrant published standards on any product mentioned; neither does the use of a trade or brand name imply approval of any product to the exclusion of others that may also be suitable.

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Issued in furtherance of Cooperative Extension work, Acts of May 18 and June 30, 1914, The University of Georgia College of Agricultural and Environmental Sciences and the U.S. Department of Agriculture cooperating.

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