Compendium of Rhododendron and Azalea Diseases and Pests, Second Edition

Compendium of

Rhododendron and Azalea Diseases and Pests SECOND EDITION

Part I. Diseases Caused by lnfectious Agents Diseases Caused by Fungi Phytophthora Root Rot Phytophthora root rot (also known as rhododendron wilt) is a serious disease that affects many species in the genus Rhodo­ dendron and several other genera in the family Ericaceae. The disease was first reported on cultivated rhododendron in 1929 in New Jersey. By that time, it had caused losses in nursery stock in both western and eastern regions of the United States where rhododendron are produced. Since then, occurrence of this disease has been reported in many parts of the world where rhododendron are produced and grown.

Phytophthora root rot is most severe on rhododendron, but it also occurs commonly on azalea and other ericaceous species, including blueberry, Chilean wintergreen, erica, heather, Irish heath, kinnikinnick, leucothoe, mountain laurel, pieris, salal, and strawberry tree. Interestingly, Phytophthora root rot has not been reported in native populations of ericaceous plants in the Appalachian Mountain region of the eastern United States, even though Phytophthora cinnamomi (P. cinnamomi var. cin­ namomi) is common in the forest soils of this region. Attempts to reintroduce Chapman’s rhododendron (R. chapmanii (Alph.

Wood) A. Gray)—­a rare and endangered species—­to coastal habitats on sandy soils in the southeastern United States had limited success, in part because of the plant’s high susceptibility to P. cinnamomi.

Phytophthora root rot occurs most commonly in nurseries on 1-­and 2-­year-­old plants, particularly those produced in containers. The disease also occurs on field-­grown nursery plants

Fig. 1. Phytophthora root rot and wilt in the field, caused by Phy­ tophthora cinnamomi. (Cour­tesy H. A. Hoitink)

and plants grown in landscapes when soils are poorly drained or subject to prolonged periods of saturation (Figs. 1 and 2).

The distribution of Phytophthora root rot on rhododendron and azalea plants in the United States depends on the species of Phytophthora present in the soil. For instance, P. cinnamomi, the most common species attacking these plants, is found in landscapes where rhododendron and azalea grow in the southeastern United States (i.e., from the mid-­Atlantic states to Connecticut) and on the west coast of North America (i.e., from California to British Columbia). P. cinnamomi does not occur above 40°N latitude (i.e., southern Ohio) in the eastern United States because it cannot survive at low temperatures, but the pathogen does occur in northern Ohio on container-­grown rhododendron in nurseries where plants are protected from low winter temperatures by overwintering structures.

Symptoms Roots of infected rhododendron plants develop necrosis, and then leaves become chlorotic and wilt. As leaves begin to wilt, they roll downward parallel to the midribs (Fig. 3). Eventually, foliage on affected plants turns brown as tissues become necrotic. In comparison, foliage on infected azalea becomes chlorotic and then necrotic, but wilting is rare. Necrotic older leaves eventually drop to the ground, giving the plant a partially defoliated appearance (Fig. 4). Azalea leaves on infected

Fig. 2. Mortality of rhododendron in the landscape (center plant) from Phytophthora root rot, caused by Phytophthora cinnamomi.

(Cour­tesy R. G. Linderman)

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plants are noticeably smaller, too. Highly susceptible 1-­and 2-­year-­old rhododendron plants may wilt and die within 14 days after infection.

Colonization may advance into all or only parts of the root system, depending on many factors. On older plants, roots frequently regenerate in the well-­drained upper layer of the soil or potting mix (Fig. 5). As the pathogen advances into the stem, cambium tissue becomes colonized first and turns dark brown.

Adjacent phloem and xylem tissues turn brown later. By that time, permanent wilt symptoms have usually developed in the foliage. Eventually, cankers are visible at the base of the stem in 1-­and 2-­year-­old plants (Fig. 6). Older plants usually do not develop visible cankers on the lower stems.

Rhododendron spp. with dense leaf hairs on the undersides of leaves, along with some moderately resistant cultivars, may not develop wilt symptoms until all of the roots have died. On such plants, yellow and eventually necrotic areas form in interveinal tissues near the midribs of the youngest leaves during the summer. Affected plants may become colonized by various secondary invaders, including weak pathogens, which results in dieback. Large plants in old field plantings frequently do not

Fig. 3. Wilting of rhododendron with Phytoph­ thora root rot (right), caused by Phytophthora cinnamomi, compared to noninoculated control (left). (Cour­tesy R. G. Linderman)

Fig. 5. New roots regenerated from the crown of a field-produced rhododendron affected by Phytophthora root rot. (Cour­ tesy H. A. Hoitink)

Fig. 4. Comparison of healthy (noninoculated) azalea cultivar Hinodegiri (top) and plant with severe Phytophthora root rot (bottom), 4 months after inoculation with Phytophthora cinnamomi.

Note the necrotic roots and much smaller root system on the inoculated plant. (Cour­tesy D. M. Benson)

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Fig. 6. Canker on a rhododendron crown stem, caused by Phy­ tophthora citricola. (Cour­tesy R. G. Linderman)

have aboveground symptoms, other than mild chlorosis and lack of growth. A few noninfected roots in an otherwise rotted root system may support such a plant until other stress factors or secondary pathogens result in its death.

In old landscape plantings and in 3-­and 4-­year-­old field-­ produced plants, symptoms of root rot are often present for 1 year or more before plants die. On such chronically infected plants, the fungus Botryosphaeria dothidea (Moug.:Fr.) Ces. & De Not. frequently causes shoot and branch dieback. This opportunistic pathogen may cause dieback on ericaceous plants weakened by other stress factors, as well.

Causal Organisms Phytophthora spp. that cause root rot and those that cause dieback are fungal-­like microorganisms in the phylum Oomycota and closely related to the heterokont algae or Chromista (Straminipila), rather than true fungi (see also later in Part I, the section Phytophthora Dieback). Phytophthora spp. and the Oomycota, in general, are different from true fungi because they contain cellulose and beta glucans in the cell wall (compared to chitin in true fungi) and have a diploid vegetative hypha that lacks cross walls (compared to septate hyphae and haploid nuclei in true fungi), among other differences. Motile zoospores possess two flagella—­one a tinsel type and the other a whiplash type—­unlike true fungi, for which motile spores possess one flagellum. Thus, Phytophthora spp. usually respond to fungicide classes differently from true fungi.

P. cinnamomi Rands (Oomycota, Pythiaceae) is the most frequently isolated and most aggressive pathogen associated with root rots on plants in the Ericaceae. Although P. cinnamomi is the most important species associated with Phytophthora root rot of rhododendron in the United States and other parts of the world (e.g., Australia, Denmark, England, France, Germany, Japan, and the Netherlands), P. cambivora (Petri) Buisman is one of the first species associated with root rot of rhododendron in the United States. Other species of Phytophthora may be more common or more aggressive on some species or genera in this large family of plants (Table 1). Several authors have examined differences in host specificity among isolates of P. cinnamomi from ericaceous plants but found no evidence for the existence of races. However, isolates of P. cinnamomi from camellia of the A1 mating type (which is uncommon in the United States) caused less severe root rot on azalea than isolates of the A2 mating type from azalea. Isolates of P. cinna­ momi from other nonericaceous hosts were also not as aggressive on Rhododendron cultivar Purple Splendor as on isolates from hybrid rhododendron.

In the southeastern United States, P. nicotianae Breda de Haan (syn. P. parasitica Dastur) is commonly isolated from roots of azaleas growing in nurseries and landscapes in the states in the Gulf Coast, whereas P. cinnamomi is commonly found on roots of azaleas from South Carolina up through the mid-­Atlantic states to Connecticut and in states along the West Coast. P. nicotianae is also a common root rot pathogen on greenhouse azalea; on container-­grown rhododendron, it has been reported only as a root rot pathogen in North Carolina and Virginia.

P. citricola Sawada and P. cryptogea Pethybr. & Laff. cause Phytophthora root rot of rhododendron, but they are considered minor pathogens. Several other species of Phytophthora have been recovered in nursery irrigation water and caused root rot in artificial inoculation of azalea, including the recently identified species P. hydropathica Hong & Gallegly, P. irrigata C. Hong and M. Gallegly, and P. tropicalis Aragaki & J. Y. Uchida. P. hydropathica and P. tropicalis have been isolated from rhododendron foliage, but P. irrigata has been recovered only from water samples. It is unknown whether these species or the other common species that cause Phytophthora dieback can cause root infections on plants growing in the nursery or the landscape.

Epidemiology Rhododendron and azalea are produced almost entirely from stem cuttings or through tissue culture; few species, except native ones, are raised from seed. P. cinnamomi is rarely isolated from stem cuttings in these crops. In contrast, the species of Phytophthora that cause dieback and shoot blight are frequently introduced into a new crop by infected cuttings. Alternatively, plants produced by tissue culture are free of these pathogens.

P. cinnamomi survives largely as chlamydospores and mycelium in infected roots, lower stem parts (including the crown), and infested host debris in soil. In the northern United States, the pathogen does not overwinter well in soil in the absence of protected host plants, as mentioned earlier. However, in temperate regions with mild climates, such as North Carolina and South Carolina, the pathogen survives in the soil and in the groundcover base on which plants in containers are placed.

Presumably, during periods of favorable soil moisture conditions, sporangia are produced and release zoospores, which infect roots. Roots and soil samples can be baited to isolate Phytophthora spp. (Figs. 7 and 8).

Inoculum of P. cinnamomi is most likely introduced into a new crop of rooted cuttings via infested irrigation water, container media, or host debris, or it can be splashed from infested surfaces on which plants have been placed. The pathogen has been isolated only infrequently from ponds and irrigation water into which surface water drained from infected rhododendron plantings, but it has been recovered from runoff water leaving infested fields. Therefore, irrigation water is not a common source of primary inoculum for P. cinnamomi. In the state of Ohio, sampling of irrigation ponds fed by groundwater did not isolate the pathogen when surface water from infested stock blocks was excluded. Other species of Phytophthora that cause

Fig. 7. Baiting a potting mix sample for Phytoph­ thora spp. with leaf disks of camellia. (Cour­tesy R. G. Linderman)

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root rot of rhododendron and azalea are more commonly isolated from irrigation ponds and retention basins when irrigation runoff is collected for reuse.

The surface or substrate on which container-­grown plants are placed affects the incidence of Phytophthora root rot. The spread of P. cinnamomi from infected plants to nearby healthy plants is greatest when containers are set on black, polyethylene plastic (which prevents drainage away from the bottoms of containers) and least when they are set on a layer of coarse gravel (which allows rapid drainage). However, splash dispersal from the surface of the container medium to nearby plants has been reported for azalea infected with P. cinnamomi and may play a role in limiting the effectiveness of a well-­drained substrate.

The population dynamics of P. cinnamomi in landscape beds vary with the severity of the root rot. In North Carolina, for example, inoculum density ranged from 2 to 18 propagules per gram of soil over a 21-­month period in the presence of a living infected host (azalea). Once the plants had been killed, however, inoculum density was 1 propagule per gram of soil or less and gradually declined to an undetectable level over the next 6–10 months.

P. cinnamomi produces chlamydospores, sporangia, zoospores, and oospores. Because the fungus is heterothallic and the A1 mating type is relatively rare in the United States, oospores are not commonly produced in nature. Any of the other propagules may play a role in infection, however. Soil moisture plays a key role in the infection process. Sporangia are produced over a wide range of matric potentials, but zoospores are released most readily in soil water at matric potentials greater than –5 millibars (mbar) (i.e., –5 to 0 mbar). Therefore, the most rapid buildup of inoculum occurs when free moisture is present (e.g., in puddles, at the water table in soil or the bases of pots, and as standing water on the soil surface).

Soil pH affects sporulation by P. cinnamomi. For example, at the low pH value of 3.3, sporangium production and zoospore release are inhibited. At pH 4.0, sporangium formation is still reduced, but zoospore release is not. Therefore, it is not surprising that maintaining soil pH at 4.0 controls Phytophthora root rot of rhododendron. Maintenance of such a low pH is not practical in a nursery, however, because growth is too slow at that level and plants may suffer nutrient deficiencies. During mist propagation of cuttings, the medium is usually at pH 3.5–4.5.

At these levels, the pH may affect sporulation.

Soil temperature has a dramatic effect on root rots caused

by P. cinnamomi. Detailed studies have been published of the effect of temperature on Phytophthora root rots of several crops, but little is known about how temperature may affect root rot on plants in the Ericaceae. On rhododendron in the Northern Hemisphere, the disease is most serious during the warm months of June, July, and August. Temperatures of soils and container media during these months range from as low as

Fig. 8. Isolates of Phytophthora spp. growing out from leaf disks of camellia used to bait an infested potting mix. (Cour­tesy R. G. Linderman)

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15°C in soils to as high as 50–55°C at the sun-­exposed walls of black or green plastic containers. On avocado, the effect of temperature on infection was found to correlate positively with effects on growth of and sporangium formation by P. cinna­ momi. Soil temperatures above 15°C are necessary for infection, but avocado plants grow as well in infested soil as they do in noninfested soil at temperatures higher than 33°C. On Cape heath in England, root rot develops more rapidly at temperatures above 17°C than at lower temperatures. Field observations of the disease in rhododendron therefore appear to support the results of published studies of the effects of temperature on this disease in other crops.

Management The technology for producing woody ornamental plants has become increasingly sophisticated. Several species of Ericaceae are now propagated almost exclusively by tissue culture.

Various indexing procedures used for floricultural crops are being used in the production of woody plants, including species of Ericaceae. Nonetheless, some nurseries still produce rhododendron from seed and in field soil. This section therefore addresses various strategies for integrated pest management (IPM), as no single strategy provides complete control. Five principal approaches are summarized and integrated: prevention, chemical control, cultural practices, biological control, and host resistance.

Prevention. The best and most effective strategy for managing root rot is prevention. To accomplish this, all sources of inocula must be eliminated. Cuttings used in propagation must be taken from disease-­free stock plants, and no soil particles should adhere to their leaves or stems. In addition, cuttings must be rooted under high standards of sanitation. The misting water used during propagation should be chlorinated, pumped from deep wells on site, or obtained from the local municipality (i.e., has been treated for human consumption). Weeds and algae in irrigation ponds should be controlled to maintain water quality. Irrigation water or misting water that has a high pH or salinity will increase disease problems during both propagation and production and therefore must be avoided.

For production of ericaceous plants in containers, following various preventive and sanitation procedures will reduce the incidence of root rot. Containers should not be reused unless they have been sanitized by aerated steam or with a liquid disinfestant. In addition, containers should be placed on gravel or a similar coarse, neutral substrate (e.g., woven plastic fabric) on a sloping base to avoid standing water and periodic flooding. Following this practice will restrict zoospore movement and splash dispersal during heavy rainfall and irrigation and ultimately decrease the number of infections. Host debris, such as prunings and abscised infected leaves, should be removed from the production area between crops to reduce the survival of inoculum. Components to be used in container media should be stored on a concrete pad, and care should be taken not to contaminate stockpiles of media with field soil or runoff water from the nursery.

Chemical control. The fungicides fosetyl-­aluminum (i.e., fosetyl-­Al), mefenoxam (and its precursor, metalaxyl), and etridiazole have been used for many years as sprays or drenches for Phytophthora pathogens to prevent root rot of container-­grown azalea and rhododendron. Recently, several fungicides with new chemistries have proven effective as drenches in disease prevention, including ametoctradin + dimethomorph, cyazofamid, dimethomorph, fenamidone, fluopicolide, and mandipropamid.

Newer formulations of phosphorus acid derivatives (e.g., phosphonate and phosphite fungicides) are also effective. Chemicals in this group of compounds can be applied as sprays or drenches to prevent root rot and are the only fungicides that are truly systemic (i.e., moving both upward and downward in the plant).

Fungicide application rates and methods vary with the crop and the product used. Multiple applications are needed each

season from the time the container medium temperature approaches 15°C in the spring until it drops below this level in the fall. None of these fungicides is therapeutic, however, and so will not cure infected plants. To produce a healthy crop, it is thus critical to begin applications before infection occurs.

Many of the currently used fungicides prevent infection and limit growth and development of the pathogen in infected plants. This means that it is possible that infected but symptomless plants and noninfected plants growing in infested potting mix may be purchased and planted in the landscape. These plants may eventually succumb to root rot in the landscape once residual activity from the nursery-­applied fungicides is no longer effective. However, an 18-­month residual activity against Phytophthora root rot of azalea was observed in mefenoxam-­ treated landscape beds naturally infested with P. cinnamomi.

With the loss of fumigants such as methyl bromide (which was banned because of its ozone-­depleting effect), options are limited for treating field soil in nurseries and landscape beds infested with Phytophthora spp. and other pathogens. Formulations containing chloropicrin, dazomet, and metam sodium are available, but safety concerns and site restrictions may limit fumigation activity in unsecured residential and public areas.

Fumigated beds often become reinfested when runoff water from adjoining areas reintroduces the pathogen to the treated soil, so practices such as ditching and creating raised beds are needed to prevent this from occurring.

Cultural practices. Sandy soils with low pH are used widely in nurseries in the eastern United States for field production of ornamental plants in the Ericaceae. The pH value that effectively limits Phytophthora root rot (≤4.0) is below the optimum pH for growth (5.0–5.5); however, soil readings of pH 3.8–4.5 are not uncommon in some of these nurseries. Low pH and other edaphic factors that prevent saturation, flooding, and puddle formation are therefore used with some success to reduce the incidence and severity of Phytophthora root rot. In heavy soils that retain water, ericaceous plants are typically produced in raised beds to improve drainage and reduce disease pressure.

Although concise research is lacking, specific guidelines for the physical properties of soilless container media have been developed to reduce the incidence of Phytophthora root rot. These guidelines are based on observations of disease incidence in nurseries and measurements of physical properties related to drainage of media. Rapid drainage of container media is the key to preventing the production and release of root-­infecting zoospores by Phytophthora spp. and thus preventing the initiation of root rot. Sand, silt, or clay should not be used in any medium for growing rhododendron, because these fine-­textured particles tend to migrate to the bottom of the container and inhibit drainage. Prolonged periods of excess moisture render any medium conducive to zoospore activity and predispose roots to infection by P. cinnamomi. Container media prepared with very fine bark particles, for example, can also be conducive to root rot. To ensure adequate drainage, the percolation rate of a medium must be approximately 2.5 cm per minute throughout the production cycle of a crop.

Woody ornamentals are largely produced in containers in which the column height of the soil or medium is 9–10 cm (in azalea pots) or 15–19 cm (in typical 1.0-­and 2.5-­gallon containers for large plants). In container mixes with these column heights, air-­filled pore spaces of 15% and 20–35%, respectively, at container capacity are associated with low losses from root rot in nurseries, irrespective of the organic components of the container media. Given this, both the air-­filled pore space and the percolation rate should be monitored to reduce losses from Phytophthora root rot.

In the landscape, using raised beds and ditching are effective strategies for minimizing the impact of Phytophthora root rot on azalea and rhododendron. Raised beds improve drainage around the root system and shorten the time that soil remains saturated after rain or irrigation. Drainage in beds with heavy

soils can be improved by amending the soils with coarse substrates, such as tree bark. Although plants may become infected in naturally infested beds, they can tolerate infection by Phy­ tophthora spp. and remain aesthetically acceptable for several years, depending on the cultivar (Figs. 9 and 10).

Temperature treatments cannot generally be used on a practical scale to control disease development. Hot-­water treatment can be used to eradicate P. cinnamomi from infected rootstock, but this option is not widely applied by growers. Theoretically, freezing can be used in cold climates to reduce the survival of P. cinnamomi in soil in the absence of host plants. However, other species that can cause root rot on rhododendron (e.g., P. cactorum and P. citricola) can survive very low temperatures in the soil, often in association with host debris.

Biological control. General suppression of Phytophthora root rot of azalea and rhododendron growing in tree bark-­based container media results from beneficial microbes associated with the bark and breakdown products of the bark itself in addition to the drainage benefit. Tree bark tissues may release compounds into the water phase of container media that inhibit zoospores and sporangia of Phytophthora spp. Canadian sphagnum peat, another common ingredient in soilless container mixes, does not release such inhibitors. Without exception, all of the barks examined to date—­including those of eucalyptus, Monterey pine, a mixture of pines from the eastern United States, and various hardwoods (e.g., oaks and maples)—­ release these inhibitors into container media when freshly removed from trees. However, the length of time over which the

Fig. 9. Hinodegiri azaleas planted in ground-level landscape beds died from Phytophthora root rot, caused by Phytophthora cinnamomi, after one growing season. (Cour­tesy D. M. Benson)

Fig. 10. Hinodegiri azaleas planted in raised landscape beds grew vigorously, even though the bed was infested with Phytoph­thora cinnamomi. (Cour­tesy D. M. Benson)

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inhibitors continue to be released after planting varies from season to season and with the species of tree from which the bark was removed. Also, the effect of the inhibitors varies with the tree species if the bark is composted before use. Hardwood bark (primarily red oak) may release inhibitors for up to 1 year after potting; the most toxic components in red oak bark have been identified as breakdown products of waxes.

Specific suppression of Phytophthora spp. that cause root rot can be accomplished by amending container media with beneficial microorganisms such as Trichoderma harzianum Rifai and T. hamatum (Bonord.) Bainier. Under highly conducive disease conditions or heavy inoculum pressure, however, these beneficial microbes may not prevent root infections by zoospores.

Host resistance. Ericaceous plants vary in susceptibility to P. cinnamomi, but most commercially produced rhododendron species and cultivars are highly susceptible. Among 336 cultivars of hybrid rhododendron tested, cultivars Caroline, Professor Hugo de Vries, and Red Head are the most resistant.1 A strain of cultivar English Roseum and several other hybrids are moderately resistant. Among 198 rhododendron species tested for resistance, R. davidsonianum Rehder & E. H. Wilson, R. delavayi Franch., and R. pseudochrysanthum Hayata are the most resistant. Feeder roots of these resistant plants do become infected, but new roots regenerate from the crowns of affected plants in the well-­drained upper layers of soil or container medium.

Extremes of soil moisture affect the susceptibility of rhododendron to P. cinnamomi. Cultivar Caroline, which is otherwise relatively resistant to Phytophthora root rot, develops severe symptoms of root and crown rot if it is exposed to drought stress (with a leaf water potential of –16 bar) or if the root system is flooded for 24–48 h before inoculation. These extreme conditions do not occur in all of the areas in which rhododendron are produced or grown, however. In nurseries, these conditions are controlled through proper management of irrigation and container media. Failure to control these conditions may explain the variability in resistance observed among different ericaceous plants in different parts of the world.

Cultivars in 10 hybrid azalea groups vary from being highly resistant to highly susceptible to root rot caused by P. cinna­ momi.2 The most resistant cultivars are Corrine Murrah, Fakir, and Formosa. As groups, the Southern Indica hybrids are the most resistant and the Carla hybrids are the most susceptible.

The azalea species R. poukhanense H. Lev. has good resistance, is cold hardy, and is commercially acceptable.

Growers should use host resistance more widely when planting rhododendron and azalea in the landscape to reduce losses from Phytophthora root rot, particularly in sites where P. cinnamomi is known to occur. Resistance is a management strategy that can be effective, sustainable over time, and environmentally sound. However, in the long term, sustainable disease management can best be achieved when all five strategies—­ prevention, chemical control, cultural practices, biological control, and host resistance—­are used together in an integrated approach.

Benson, D. M., and Blazich, F. A. 1989. Control of Phytophthora root rot of Rhododendron chapmanii A. Gray with Subdue. J. Environ.

Hort. 7:73-­75.

Benson, D. M., and Cochran, F. D. 1980. Resistance of evergreen hybrid azaleas to root rot caused by Phytophthora cinnamomi. Plant Dis. 64:214-­215.

Benson, D. M., and Parker, K. C. 2007. Efficacy of registered and unregistered fungicides for control of Phytophthora root rot of azalea, 2006. Plant Dis. Manag. Rep. doi:10.1094/PHP-2011-0512-01-RS. Benson, D. M., Shew, H. D., and Jones, R. K. 1982. Effects of raised and ground-­level beds and pine bark on survival of azalea and population dynamics of Phytophthora cinnamomi. Can. J. Plant.

Pathol. 4:278-­280.

Blaker, N. S., and MacDonald, J. D. 1981. Predisposing effects of soil moisture extremes on the susceptibility of rhododendron to Phytophthora root and crown rot. Phytopathology 71:831-­834.

Bush, E. A., Hong, C. X., and Stromberg, E. L. 2003. Fluctuations of Phytophthora and Pythium spp. in components of a recycling irrigation system. Plant Dis. 87:1500-­1506.

Englander, L., Merlino, J. A., and McGuire, J. J. 1980. Efficacy of two new systemic fungicides and ethazole for control of Phytophthora root rot of rhododendron, and spread of Phytophthora cinnamomi in propagation benches. Phytopathology 70:1175-­1179.

Ferguson, A. J., and Jeffers, S. N. 1999. Detecting multiple species of Phytophthora in container mixes from ornamental crop nurseries.

Plant Dis. 83:1129-­1136.

Gerlach, W. W. P., Hoitink, H. A. J., and Schmitthenner, A. F. 1976.

Survival and host range of Phytophthora citrophthora in Ohio nurseries. Phytopathology 66:309-­311.

Hoitink, H. A. J., and Schmitthenner, A. F. 1974a. Relative prevalence and virulence of Phytophthora species involved in rhododendron root rot. Phytopathology 64:1371-­1374.

Hoitink, H. A. J., and Schmitthenner, A. F. 1974b. Resistance of rhododendron species and hybrids to Phytophthora root rot. Plant Dis. Rep. 58:650-­653.

Hoitink, H. A. J., VanDoren, D. M., Jr., and Schmitthenner, A. F. 1977.

Suppression of Phytophthora cinnamomi in a composted hardwood bark potting medium. Phytopathology 67:561-­565.

Linderman, R. G., and Zeitoun, F. 1977. Phytophthora cinnamomi causing root rot and wilt of nursery-grown native western azalea and salal. Plant Dis. Rep. 61:1045-1048.

Ownley, B. H., and Benson, D. M. 1991. Relationship of matric water potential and air-­filled porosity of container media to development of Phytophthora root rot of rhododendron. Phytopathology 81:936-­941.

Spencer, S., and Benson, D. M. 1982. Pine bark, hardwood bark compost, and peat amendment effects on development of Phytophthora spp. and lupine root rot. Phytopathology 72:346-­351.

Vegh, I., and Frossard, C. 1980. Study of some factors of variation of French Phytophthora cinnamomi strains as parasites of ornamental shrubs. Phytopathol. Z. 99:101-­104.

White, R. P. 1936. Summary of nine years’ experience with rhododendron wilt. Plant Dis. Rep. 20:204-­207.

Selected References

Armillaria Root Rot

Benson, D. M. 1984. Influence of pine bark, matric potential, and pH on sporangium production by Phytophthora cinnamomi. Phyto­ pathology 74:1359-­1363.

Benson, D. M. 1987. Residual activity of metalaxyl and population dynamics of Phytophthora cinnamomi in landscape beds of azalea.

Plant Dis. 71:886-­891.

Benson, D. M. 1990. Landscape survival of fungicide-­treated azaleas inoculated with Phytophthora cinnamomi. Plant Dis. 74:635-­637.

Armillaria root rot is caused by Armillaria mellea and possibly other Armillaria spp. These basidiomycete fungi (order Agaricales, subdivision Basidiomycota) produce spores in fleshy fruiting bodies (i.e., mushrooms), so the disease is sometimes called mushroom root rot. In addition, because the pathogen produces elongated, vegetative strands of rhizomorphs that have dark coatings or sheaths and look like shoestrings, the disease is sometimes called shoestring root rot. In the U.S. state of California and other areas, the disease often occurs on oak trees and is therefore sometimes known as oak root fungus root rot.

The first report of A. mellea on rhododendron was in New York State in 1891, but the disease has also been reported in

or the complete list of cultivars tested, see Hoitink and Schmitthenner F (1974b).

2 For the complete list of cultivars tested, see Benson and Cochran (1980).

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(Prepared by H. A. J. Hoitink, D. M. Benson, and A. F. Schmitthenner; revised by D. M. Benson and S. N. Jeffers)