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Botrytis Blight of Flowering Potted Plants

Margery L. Daughtrey, Cornell University, Long Island Horticultural Research Laboratory, Riverhead, NY 11901; Robert L. Wick, University of Massachusetts, Amherst, MA 01002; and Joseph L. Peterson, Rutgers University, New Brunswick, NJ 08901

Posted 5 June 2000. Plant Health Progress doi:10.1094/PHP-2000-0605-01-HM.

Reproduced, with permission, from Compendium of Flowering Potted Plant Diseases, pp. 11-14, 1995. The American Phytopathological Society, St. Paul, MN, U.S.A.

Botrytis cinerea has a worldwide distribution and is ubiquitous in greenhouses. Botrytis blight is one of the most common diseases of greenhouse crops.


Fig. 1. Lesions on hydrangea leaves caused by Botrytis cinerea during propagation. (Courtesy S. H. Davis)

B. cinerea causes a range of symptoms including spots and blight on leaf or petal tissues (Fig. 1), crown rot (Figs. 2 and 3), stem cankers (Figs. 4-7), cutting rot, and damping-off. Storage tissues such as roots, corms, or rhizomes are also susceptible. Wounded or senescent tissues are especially susceptible to invasion, but healthy tissues may also become colonized. Lesions caused by B. cinerea are often identified in the field by the characteristic gray, fuzzy sporulation (Fig. 8). However, spores develop only under humid conditions. Leaf lesions often develop a zonate pattern. Flower petals may have tiny flecks of discoloration or become completely blighted. Stems may die back from cutting wounds or develop tan to brown cankers that originate at the bases of petioles of blighted leaves.

Fig. 2. Botrytis crown rot of calceolaria. (Courtesy A. R. Chase)

Fig. 3. Botrytis canker of lisianthus (Eustoma grandiflorum).

Fig. 4. Botrytis stem canker on vinca (Catharanthus roseus). (Courtesy M. A. Hansen)

Fig. 5.  Sporulation of Botrytis cinerea on a stem lesion on Impatiens wallerana. (Courtesy J. A. Matteoni)

Fig. 6. Sporulation of Botrytis cinerea on hydrangea stems. (Courtesy J. A. Matteoni)

Fig. 7. Stem rot of kalanchoe, showing sporulation of Botrytis cinerea. (Courtesy D. Karasevicz)

Fig. 8. Sporulation
of Botrytis cinerea
on hydrangea stems. (Courtesy J. A. Matteoni)

Poinsettia (Euphorbia pulcherrima Willd. ex Klotzsch) is susceptible at all stages of production. During propagation, leaves may become blighted and cuttings may rot. On potted poinsettias, tan to brown lesions form on leaves, stems, and bracts. Extensive tan cankers can form on stems when B. cinerea enters via blighted petioles or shoots (Fig. 9). Lesions begin at the margins of bracts, turning darker as they expand (Figs. 10 and 11). Latex may exude from the undersides of lesions. Sporulation develops on the necrotic areas under humid conditions (Fig. 12).

Fig. 9. Tan stem lesion on poinsettia caused by Botrytis cinerea.

Fig. 10. Lesions caused by Botrytis cinerea developingat he edge of poinsettia bracts. (Courtesy M. F. Heimann)

Fig. 11.  Botrytis cinerea lesion at the tip of a poinsettia bract.

Fig. 12.  Botrytis cinerea sporulation on poinsettia flower parts. (Courtesy M. K. Hausbeck)

Cyclamen (Cyclamen persicum Mill.) petals, petioles, and leaves are susceptible to B. cinerea. On petals, the initial flecks enlarge to 14 mm, progressing from water-soaked lesions to tan, necrotic spots. Colored petals show rings of intensified color around the lesions. The petioles and flower stalks developing beneath the canopy may become infected and rot, causing portions of the plant to collapse. Subsequently, the infected petioles and pedicels typically become covered with grayish brown B. cinerea sporulation. Symptoms of Fusarium wilt or bacterial soft rot of cyclamen are quite similar; however, these diseases cause characteristic vascular discoloration and a soft, mushy rot of the corm, respectively.

Fig. 13. Botrytis cinerea sporulation on tan canker at the base of a Persian  violet (exacum) stem. (Courtesy J. A. Matteoni)

Exacum (Exacum affine Balf. f.) is particularly susceptible to Botrytis blight. Young seedlings are susceptible to damping-off. A mature plant, which suddenly wilts, may have a reddish brown Botrytis canker at the base of the stem, originating at a node where leaves were buried at transplanting (Fig. 13). This symptom may be confused with the basal canker produced by Fusarium solani (Nectria haematococca) and with the black stem canker caused by impatiens necrotic spot tospovirus (INSV). Initially, a stem canker on exacum caused by B. cinerea has a water-soaked appearance and may be bordered by a white halo. Both B. cinerea and INSV cause pale tan cankers on the secondary stems. Leaves infected with B. cinerea develop tan spots approximately 3 mm in diameter or bleaching or necrosis of the leaf margin. Lesions may coalesce to form large, tan, zonate, membranous lesions. Infected flowers exhibit water-soaked flecking on the petals or may become completely colonized and then collapse and turn brown. Flower blight frequently leads to colonization of the pedicel and stem.

On African daisy (Gerbera jamesonii H. Bolus ex Adlam), B. cinerea can cause damping-off, spotting and blighting of leaves and flowers, and crown rot. Leaves develop zonate lesions, and flower petals show tan spots and tip necrosis or are entirely blighted. B. cinerea may be seedborne in African daisy.

Fig. 14. Dieback of florist's geranium caused by Botrytis cinerea. (Courtesy M. K. Hausbeck)

Fig. 15. Leaf lesions on florist's geranium resulting from Botrytis cinerea-infected flowers falling on the foliage.

Geraniums (Pelargonium spp.) are very susceptible to B. cinerea. Annual losses for 1985 were estimated at $5.17.6 million worldwide. Losses are particularly high in vegetatively propagated P. hortorum L. H. Bailey. Stock plants are grown close together and have multiple branches and a dense canopy that is created with growth regulators in order to maximize cutting productivity. Consequently, lower leaves of stock plants typically senesce and become colonized by B. cinerea, resulting in a massive buildup of inoculum. Stock plants are wounded when cuttings are harvested, often resulting in stem blight, which progresses down toward the main stem (Fig. 14). B. cinerea infections may also develop at the cutting bases or at leaf scars during propagation under mist. Flower blight is common. P. hortorum grown from seed is also susceptible to B. cinerea. Brown, zonate leaf lesions are often initiated after petals drop from the flowers or from a hanging basket crop above (Fig. 15). Geraniums sometimes show ghost spots indicative of Botrytis lesions whose development has been arrested.

Causal Organism

Fig. 16. Botrytis cinerea sporulation. (Courtesy J. A. Matteoni)

Botrytis cinerea Pers.:Fr. is a hyphomycete with straight, brown, septate conidiophores, which are simple or, more commonly, alternately branched. The conidiophores bear botryose clusters of hyaline conidia, which appear grayish brown in mass (Fig. 16). Conidia are budded off from a swollen, sporogenous cell at the tip of the conidiophore. The conidia (814 69 m) are ellipsoid to ovoid. Botrytis spp. are pleomorphic: the conidial anamorph is in the form-genus Botrytis Pers.:Fr., the microconidial anamorph is Myrioconium H. Sydow, and the sclerotial anamorph is a Sclerotium Tode. The teleomorph for B. cinerea is Botryotinia fuckeliana (de Bary) Whetzel, which is only rarely observed forming apothecia on sclerotia.

Colonies of B. cinerea on potato-dextrose agar are at first off-white and become gray to brown with the development of spores. The optimum temperature for growth is reported to be 2428C, but some growth occurs from 0 to 35C. Sclerotia are black, hard, tightly appressed to the substrate, irregular in size and shape, and 115 mm long or confluent. Sclerotia have a dark, pseudoparenchymatous rind of nearly isometric cells approximately 510 m in diameter enclosing a medulla of tightly knit, hyaline, thick-walled hyphae.

Host Range and Epidemiology

Fig. 17A.

Fig. 17B.

Fig. 17C.

Fig. 17.  Infection of a geranium cutting wound by Botrytis cinereaA, Spore germination 3 hr after inoculation (1,000x); B, germ tube elongation 6 hr after inoculation (3,200x); and C, tissue invasion 12 hr after inoculation (100x). (Courtesy M. K. Hausbeck)

B. cinerea has a very broad host range, especially among dicots, and can cause "gray mold" disease within a wide range of temperatures. All of the potted plants covered in this compendium are susceptible to Botrytis blight.

Infection may occur directly or through natural openings or wounds (Fig. 17) by means of conidial germ tubes or by hyphal growth from colonized dead plant parts or organic debris that contacts healthy tissue. Infection by B. cinerea is stimulated by nutrient depletion of leaves and pollen deposition. Fruits and flowers of plants are generally more susceptible to infection than healthy, nonsenescent leaves. Bracts and flowers of poinsettias are particularly prone to infection by B. cinerea (Fig. 11).

Conidia are released by a hygroscopic mechanism in association with a rapid change in relative humidity and require air currents or splashing water for dispersal within the greenhouse. In geranium stock-plant production areas, peak concentrations of Botrytis conidia have been associated with worker activity (Fig. 18). Harvesting cuttings, spraying pesticides, cleaning plants, and even drip-tube watering increases the number of B. cinerea conidia in the greenhouse atmosphere. Releases of 5041,297 conidia per cubic meter per hour have been recorded during P. hortorum cutting harvest, a time at which both cuttings and stock plants are freshly wounded.

Fig. 18. Release of Botrytis cinerea spores in geranium stock correlated with worker activity. Numbers in parentheses are daily total spore counts (reprinted, by permission, from Hausbeck and Pennypacker, 1991) (click image for a larger view).


Once dispersed, conidia can germinate in a water film that contains solutes; the more conidia in a water droplet, the greater the likelihood of an aggressive infection. African daisy flowers kept at 1825C and 100% relative humidity for only 5 hr develop small, necrotic lesions from single-conidium infections of the ray florets encompassing one to several epidermal cells. Exposure of open African daisy flowers to conidia and subsequent periods of high relative humidity during shipment and storage will lead to the formation of small, necrotic lesions of the florets. Even after having been maintained under dry conditions for as long as 14 months, B. cinerea conidia retain their ability to infect African daisy florets once a film of moisture is available. Germination and lesion development on African daisy have been observed at 425C, but not at 30C. Poinsettias grown at constant 10C show more Botrytis blight incidence than those grown at various 10C night and 17C day split-night temperature regimes or those grown at constant 17C. Presumably, this is an indirect effect of higher relative humidity resulting from lower temperatures rather than a direct temperature effect.

Cultural practices often create opportunities for B. cinerea infections. For example, making cutting wounds on stock plants and stripping the lower leaves from cuttings facilitate infection. When crops of flowering plants are grown in hanging baskets over a crop susceptible to Botrytis blight, the fallen petals may serve as an energy source for the fungus and facilitate infection of adjacent healthy tissue.

Sclerotia formed in colonized plant tissues are important for long-term survival of B. cinerea in soil; under appropriate conditions, they will germinate and produce conidial (or, possibly, ascospore) inoculum.


Control of B. cinerea is challenging because of its abilities to survive as a saprophyte, rapidly invade host tissues, and quickly produce abundant conidia that are easily distributed by air currents. Sanitation alone is not sufficient for minimizing Botrytis blight losses in the greenhouse. Prevention should be the main focus of a Botrytis blight management program. An integrated strategy combining environmental management, cultural practices, and fungicides will most effectively manage this omnipresent threat in the greenhouse.

Concerns about potential injury from fungicides, unsightly residues, limited registered fungicides, and the development of fungicide resistance make environmental management the obvious first line of defense against Botrytis blight. Effective control requires careful attention to managing leaf wetness duration and relative humidity. Providing ventilation and heat at sunset will help drive moisture-laden air out of the greenhouse, reducing the opportunities for infection during the night. Several cycles may be necessary. Adequate spacing between plants should be maintained, and open-mesh benching and horizontal airflow systems that improve air circulation in the greenhouse should be used.

For optimum and efficient control of Botrytis blight on geraniums, it has been recommended that growers regulate the environment and apply fungicide treatments at times when inoculum is high and wounds are available. Lowering relative humidity within a geranium canopy to less than 93% and controlling the occurrence and duration of free moisture on foliage are important environmental modifications.

Chemical control has been complicated by the appearance of widespread benzimidazole (including benomyl and thiophanate-methyl) resistance and a less pervasive dicarboximide (iprodione and vinclozolin) resistance in populations of B. cinerea in the greenhouse flower industry in Europe and North America. Insensitive strains are exchanged among greenhouses along with the exchange of plant material. After a greenhouse B. cinerea population acquires benzimidazole or dicarboximide resistance, it appears that the trait remains within the population after the fungicide is no longer being applied.

Benzimidazole and dicarboximide fungicides are useful only on nonresistant strains of B. cinerea. Other fungicides registered in the United States for control of Botrytis in greenhouses, including chlorothalonil, copper hydroxide, copper sulfate pentahydrate, and mancozeb, are generally more effective against B. cinerea. The use of tank mixtures that include a single-site and a multisite fungicide at reduced dosages has been proposed to avoid control failure caused by fungicide resistance. Product labels provide information about approved uses and tank mixtures on flowering potted plants.

Fungicide phytotoxicity may present a particular problem for growers attempting Botrytis control on crops during the finishing stages. Chlorothalonil smoke dust has been reported to injure poinsettia bracts and the flowers of several potted plants. Injury to poinsettias is often cultivar specific. High temperature and moisture on plant surfaces contribute to chlorothalonil smoke dust injury. Of several fungicides tested on exacum, only chlorothalonil gave effective control of B. cinerea with no phytotoxic effects.

Sprays of Trichoderma harzianum Rifai, a bioantagonistic fungus, have shown some efficacy against Botrytis on cyclamen, but the effect is inferior to control with iprodione. More widespread occurrence and recognition of greenhouse strains of B. cinerea resistant to some of the currently effective fungicides will encourage the development of biocontrol agents in the future.

Cultivars often vary remarkably in their susceptibility to Botrytis, but major gene resistance has not been identified for any plant species.

Selected References

Hausbeck, M. K., and Pennypacker, S. P. 1991. Influence of grower activity and disease incidence on concentrations of airborne conidia of Botrytis cinerea among geranium stock plants. Plant Dis. 75:798-803.

Jarvis, W. R. 1980. Epidemiology. Pages 219-250 in: The Biology of Botrytis. J. R. Coley-Smith, K. Verhoeff, and W. R. Jarvis, eds. Academic Press, London.

Jarvis, W. R. 1992. Managing Diseases in Greenhouse Crops. American Phytopathological Society, St. Paul, MN.

McCain, A. H. 1983. Gray mold control of cyclamen using Trichoderma harzianum and reduced rates of vinclozolin. Pages 1-2 in: Calif. Plant Pathol. 63.

Moorman, G. W., and Lease, R. J. 1992. Benzimidazole- and dicarboximide-resistant Botrytis cinerea from Pennsylvania greenhouses. Plant Dis. 76:477-480.

Moorman, G. W., and Lease, R. J. 1992. Residual efficacy of fungicides used in the management of Botrytis cinerea on greenhouse-grown geraniums. Plant Dis. 76:374-376.

Orlikowski, L., Hetman, J., and Tjia, B. 1974. Control of seed-borne Botrytis cinerea (Pers. ex Fr.) on Gerbera jamesonii Bolus. HortScience 9:239-240.

Pappas, A. C. 1982. Inadequate control of grey mould on cyclamen by dicarboximide fungicides in Greece. Z. Pflanzenkrankh. Pflanzenschutz 89:52-58.

Tompkins, C. M., and Hansen, H. N. 1948. Cyclamen petal spot caused by Botrytis cinerea, and its control. Phytopathology 38:114-117.

Trolinger, J. C., and Strider, D. L. 1984. Botrytis blight of Exacum affine and its control. Phytopathology 74:1181-1188.

Trolinger, J., and Strider, D. L. 1985. Botrytis diseases. Pages 17-101 in: Diseases of Floral Crops. D. L. Strider, ed. Praeger, New York.

Vali, R. J., and Moorman, G. W. 1992. Influence of selected fungicide regimes on frequency of dicarboximide-resistant and dicarboximide-sensitive strains of Botrytis cinerea. Plant Dis. 76:919-924.