By Melinda Sullivan

PP 728 – Soilborne Pathogens Class Project

Phytophthora parasitica var. nicotianae causes black shank, a severe root and crown rot of all types of cultivated tobacco. In 1896, Van Breda de Haan named the fungus Phytophthora nicotianae, but he described a mixed culture and failed to give a Latin description for the organism (3). In India, in 1913, Dastur described a similar organism pathogenic to castor bean (but not to tobacco) and named the organism P. parastica. Tucker renamed the fungus that attacks tobacco P. parasitica Dastur var. nicotianae (Breda de Haan) Tucker, and this is the name currently used by most tobacco pathologists (3). In 1963, Waterhouse concluded that the name P. nicotianae had priority and stated that the correct name of the black shank pathogen of tobacco was P. nicotianae Breda de Haan var. nicotianae Waterhouse (3). Both names are used to describe this pathogen, but they should not be considered synonymous (3). Until a detailed taxonomic study is conducted to clarify the current taxonomic confusion, P. parasitica var. nicotianae will be used in this profile.

At least four races of the pathogen are known. Race O is predominate. Other races have been reported from specific geographical locations, developing as a result of specific resistance genes used in cultivars grown in the area (3).

Tobacco (Nicotiana tabacum) is the only known host for this pathogen. The disease was first described by Van Breda de Haan from Java (Indonesia) in 1896 (3). The disease was introduced into the United States and first observed in southern Georgia around 1915 (3). P. parasitica var. nicotianae was first introduced into the flue-cured and burley tobacco areas of North Carolina, Tennessee, and Kentucky in the 1930s and is now widely established in all major tobacco-producing areas in the United States. Black shank occurs worldwide, but the disease is most severe in warmer climates (3).

The fungus can be recovered from soil or plant material by baiting procedures or by use of a selective agar medium (3). An excellent selective medium for the detection and quantification of the pathogen contains either cornmeal agar or 5% V8 juice plus 20 g agar as a basal medium, plus 5 mg pimaricin, 10 mg rifampicin, 250 mg ampicillin, 125 mg a.i. pentachloronitrobenzene, and 50 mg hymexazol, in a liter of water. The colony morphology on this medium is the characteristic dense, rosette pattern. At room temperature, growth can be observed on this medium within two-to-three days. An enzyme-linked immunosorbent assay technique has also been developed for the detection and quantification of P. parasitica var. nicotianae in tissue and soil (3).

P. parasitica var. nicotianae is a soilborne plant pathogen belonging to the class Oomycetes, the order Peronosporales, and the family Pythiaceae. All stages of the organism are diploid. Hyphae are uniform or irregular in width (3-11 µm); tuberculate or smooth; hyaline to light yellow in color; and non-septate but possessing pseudosepta with age (3). P. parasitica var. nicotianae is heterothallic, a condition that generally requires two mating types are required for production of oospores. Oospores are thick-walled sexual spores, smooth, spherical, hyaline-to-straw colored, and average 25 µm in diameter (3). Sporangia are asexual; ovate, ovoid, pyriform, or spherical; usually possessing a single prominent papilla; hyaline to light yellow or golden brown in color; and variable in size (18-70 x 14-39 µm) (3). Zoospores are 7-11 µm in length and biflagellate. Chlamydospores are asexual spores; spherical to ovoid; nonpapillate; thick-walled; hyaline to brown or lemon-to-straw colored; and range from 14 to 60 µm in diameter (3).

Cardinal temperatures for various stages in the life cycle of the pathogen are 10-12^O C minimum, 24-30^O C optimum, and 36^O C maximum (3). Sporangia require high oxygen concentration for maximum production, whereas chlamydospores are not as sensitive to oxygen concentration (3). The pathogen grows well on most culture media, but colony morphology can vary from medium to medium.

The black shank pathogen primarily affects the roots and basal stem region of the tobacco plant, but all plant parts can become infected (3). Symptoms vary with age and weather conditions. Young seedlings are very susceptible and typical "damping-off" symptoms develop during wet and mild weather.

In the field, the first aboveground symptom is a wilting of the leaves of young transplants during the middle of the day, but adult plants can also show the wilting symptoms. Plants usually recover overnight, only to wilt more severely the next day. In contrast to the vascular wilt pathogens (a one-sided wilt), all plant leaves wilt. Depending upon the level of soil moisture and host resistance, leaves begin to turn yellow and hang down the stalk over the next few days to weeks and the plants are stunted in growth(3).
 

If the plants are pulled and the roots are examined, one or more of the large lateral roots are blackened or many of the fine adventitious roots arising from the stem are decayed. As the disease progresses, infection spreads into the stem of plant and often into the entire root system. In the final stages, the stem or shank turns black 30 cm or more aboveground (thus the name black shank). At this point, leaves turn brown, shrivel, and are not marketable. When the stem of a diseased plant is split longitudinally, the pith appears dry, brown-to-black in color, and is usually separated into plate-like disks. Although characteristic of black shank, the disking symptom alone is not diagnostic. Other biotic and some abiotic factors can cause a disking symptom (3).

During periods of rainy weather, the lower leaves of tobacco may become infected by the pathogen as infested soil is splashed onto the leaves. Leaf spots at first are water-soaked and light green. The lesions rapidly expand, turn brown and necrotic, and develop as circular lesions up to 8 cm in diameter (3).
 

Black shank is a warm-weather disease. Soil temperatures above 20^O C are required for infection (1). High soil moisture conditions also enhance disease development. Little or no infection occurs in soil that is not saturated for at least brief periods of time. Soil saturation is required for the release and dissemination of zoospores, the primary infective spore of P. parasitica var. nicotianae (2). Sporangium production is less sensitive to water potential (2, 3).

Black shank occurs in acid and alkaline soils, but the optimum pH for disease development is between 6 and 7. Levels of calcium and magnesium are positively correlated with black shank severity (3). It is unknown, however, if the effects of calcium and magnesium are on the fungus or the host. Tri-valent aluminum available in soil at pH 5 and below is inhibitory to some stages of pathogen development and may effect disease control (3).

Inoculum density of less than one propagule per gram of soil is sufficient to start an epidemic of black shank (3). Infected crop debris and chlamydospores free in the soil serve as primary inoculum of P. parasitica var. nicotianae. In warm, moist soil, chlamydospores germinate to produce one to several germ tubes. The chlamydospores either infect tobacco roots directly or produce a sporangium. Each sporangium germinates indirectly to produce 5-to-30 zoospores or directly by means of a germ tube. Zoospores released during soil saturation are attracted to roots via chemotaxis, where they encyst and germinate within one hour (3). Infection occurs quickly, and the fungus grows rapidly through the epidermis and into the cortex. The fungus later reproduces within the host tissue (thus the pathogen is a hemibiotroph) to produce additional sporangia or chlamydospores (3). The cycle then repeats with the sporangia or chlamydospores germinating to cause further infections. Thus, black shank is a polycyclic disease.

As roots of infected plants decompose, chlamydospores are released into the soil, where they can survive for several years in the absence of a tobacco host. The fungus survives poorly in cold soil, which probably accounts for the low level or absence of the disease in cooler regions where tobacco is grown (3).

The pathogen is spread on infected transplants, in water, or in soil. Infected transplants may not show symptoms, but under favorable conditions the disease continues to develop after transplanting. Rainwater transports infested soil and fungal spores. Soil adhering to farm equipment and even the shoes of workers can also spread the pathogen (3).

The severity of black shank is greatly increased in the presence of root-knot nematodes. Nematodes provide a means of entry for the fungus, and the fungus rapidly colonizes the giant cells that the nematode induces (3). Cultivars resistant to black shank lose much of their resistance if infected by root-knot nematodes.

Management of black shank requires an integrated approach including: crop rotation, fungicides, nematode control, cultural practices, and resistant varieties (3). The fungicide metalaxyl/mefanoxum has been effective at reducing disease losses to black shank, especially when used in conjunction with other management strategies. Soil fumigation to control nematodes may also be important where root-knot nematode is of concern. Crop rotation is very effective at reducing inoculum density. Because the pathogen causes more damage in warm, wet weather and can survive in infected plant debris, cultural practices such as sanitation and proper drainage can be effective in disease control. Host resistance is an effective strategy for black shank disease management. Selection of the proper variety, however, is a critical management decision, because resistant varieties vary significantly in their level, type of resistance, and overall yield capability (3).

1. Jacobi W.R., Main C.E., and Powell N.T. 1983. Influence of temperature and rainfall on the development of tobacco
    black shank. Phytopathology 73: 139-143.

2. Shew H.D. 1983. Effects of soil matric potential on infection of tobacco by
    Phytophthora parasitica var. nicotianae. Phytopathology 73: 1090-1093.

3. Shew H.D., Lucas G. B. 1991. Compendium of Tobacco Diseases. Amer. Phytopath. Soc., St. Paul, Minnesota.

Melinda Sullivan wishes to acknowledge the help of Dr. H. David Shew from the Department of Plant Pathology at NC State University.  Dr. Shew provided much of the information and all of the pictures used in this pathogen profile.

Black Shank

NC State Research

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                                                                                                                                                           Posted Spring 2001