Compendium of Ornamental Palm Diseases and Disorders

Preface The editors sincerely thank the many people who assisted in the preparation of this compendium. Authors besides the editors (R. M. Giblin-Davis, University of Florida, IFAS, Fort Lauderdale Research and Education Center; N. A. Harrison, University of Florida, IFAS, Fort Lauderdale Research and Education Center; and P. Jones, Institute of Arable Crops Research, Rothamsted, U.K.) contributed to the compendium.

Likewise, slides and figures were obtained not only from the editors but also from the following individuals, and we thank them for their contributions: D. Caldwell, A. Chase, R. Cullen, J. Downer, R. Giblin-Davis, N. Harrison, G. Holcomb, C. Kadooka, A. Meerow, D. Ogata, H. Ohr, J. Ooka, P. Roberts, S. Vann, and F. Wong. Also thanked are reviewers Minoru Aragaki and Gordon Holcomb and APS PRESS editor Gary Moorman, who helped us bring this project through to its successful conclusion. The staff of APS PRESS also deserves a thank you for their tolerance, patience, and excellent editing capabilities.

Palms are beautiful ornamental plants, and we sincerely hope this compendium will be useful in growing and maintaining these “princes� of the plant world.

This compendium is a revision of Diseases and Disorders of Ornamental Palms, edited by A. R. Chase and T. K. Broschat, and first published in 1991 by APS PRESS. Material has been updated and expanded, and the format has been changed to that of an APS PRESS compendium, including the addition of selected references. Insects of palms have not been included since this topic is thoroughly covered elsewhere.

The compendium is divided into two parts: Infectious Diseases and Physiological Disorders. Diseases and disorders may look very similar in palms. For example, potassium deficiency in some palms is expressed as leaf spots on old leaves, hence the reference to confluent orange spotting or orange blotch in the literature. Fungi can be isolated from leaf spots associated with nutritional disorders, but isolation is not necessarily indicative of the cause of the leaf spots, since many saprobic (nonpathogenic) fungi quickly invade and grow on tissue weakened or killed by nutritional deficiencies. Furthermore, since some nutritional deficiencies or excesses produce symptoms on leaves of different ages, it is also important to note the leaf age of affected palms before making a diagnosis. Distribution and types of spots or symptoms are also clues to the cause. Fronds exhibiting only leaflet tip death indicate a root or nutritional problem and not a foliar disease. Thus, this compendium attempts to provide a broad view of disease, nutritional, and environmental problems that plague palms in the ornamental production and landscape industries.

M. L. Elliott T. K. Broschat J. Y. Uchida G. W. Simone

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Contents 43 43 45 46 47 47

Introduction 1 4 4

Palm Anatomy and Growth Palm Species Diseases and Disorders of Palms

Part I. Infectious Diseases

Diseases Caused by Nematodes Red Ring Burrowing Nematode Other Nematodes Diseases Caused by Protozoans Hartrot and Marchitez Sorpresiva

Part II. Physiological Disorders

5 Disease Caused by Algae 5 Algal Leaf Spot 6 Diseases Caused by Bacteria 6 Bacterial Blight 7 Bacterial Bud Rot 8 Sudden Decline 8 Diseases Caused by “fungi�

9 Annellophora Leaf Spot 9 Bipolaris and Exserohilum Leaf Spots 12 Botrytis Leaf Spot and Blight 12 Calonectria Leaf Spot (Cylindrocladium Leaf Spot) 14 Colletotrichum Leaf and Fruit Spot 15 Damping-Off 17 Diamond Scale 17 Fusarium Wilt 17 Canary Island Date Palm Wilt 19 Date Palm Wilt (Bayoud Disease) 21 Oil Palm Wilt 22 Ganoderma Butt Rot (Basal Stem Rot) 25 Gliocladium Blight (Pink Rot) 26 Graphiola Leaf Spot (False Smut) 27 Pestalotiopsis Diseases 29 Phytophthora Diseases 32 Pseudocercospora and Cercospora Leaf Spots 33 Rachis Blight 35 Stigmina Leaf Spot 35 Tar Spot 37 Thielaviopsis Diseases (Black Scorch, Stem Bleeding, Dry Basal Rot, Trunk Rot, Heart Rot) 39 Disease Caused by a Phytoplasma 39 Lethal Yellowing 41 Diseases Caused by Viroids and Viruses 41 Cadang-Cadang and Tinangaja 42 Viral Diseases 42 Coconut Foliar Decay 42 Ringspot 42 Viral Diseases Caused by Potyviruses

49 49 50 50 51 51 52 52 52 53 54 54 54 55 55 56 56 57 57 57 57 58 58 58 58 59 59 60 60 60 61 61 61

Nutritional Disorders Boron Deficiency Boron Toxicity Calcium Deficiency Chlorine Deficiency Copper Deficiency Copper Toxicity Fluoride Toxicity Iron Deficiency Magnesium Deficiency Manganese Deficiency Molybdenum Deficiency Nitrogen Deficiency Phosphorus Deficiency Potassium Deficiency Sulfur Deficiency Zinc Deficiency Other Physiological Disorders Air Pollution Albinism Cold Injury Excessive Water Uptake Foliar Salt Injury Hapaxanthic Flowering Herbicide Toxicity High Level of Soil Soluble Salts Improper Planting Depth Lightning Injury Power Line Decline Root Suffocation Sunburn Water Stress Wind Damage

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Index

Color Plates (following page 32)

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Introduction Linnaeus named palms “Principes”, the princes of the plant world. This noble status is reflected in the size of some species, with the largest leaf and seed known in the plant kingdom found in the palm family. The leaves of Raphia regalis can reach 82 feet (~25 m) long. The famous double coconut seed of Lodoicea maldivica can weigh 50 pounds (~23 kg). Besides their stature, palms are one of only a few plant families that provide both food and shelter for people, while at the same time are admired and collected for aesthetic reasons.

Palms belong to a natural but distinctly separate family of plants called Arecaceae (synonym = Palmae). These plants are highly diverse morphologically and ecologically, and they are common in tropical, subtropical, and Mediterranean climatic regions of the world. Palms are a source of food and oil (e.g., Bactris, Borassus, Butia, Cocos, Elaeis, Euterpe, Metroxylon, Phoenix, Sabal, and Salacca spp.), fiber (e.g., coir from Cocos nucifera, cords from Washingtonia spp., and rope from Borassus flabellifer), baskets/hats/mats (e.g., Borassus, Cocos, Livistona, Pritchardia, Raphia, Sabal, and Washingtonia spp.), rattan (e.g., Calamus spp.), tannin (e.g., Areca and Sabal spp.), lumber (e.g., Cocos and Oncosperma spp.), and thatch for roofing material (e.g., Borassus, Cocos, Livistona, and Sabal spp.).

Exotic, lesser-known uses of palms include making wine or other drinks (e.g., Arenga, Borassus, Caryota, and Jubaea spp.) and providing a narcotic high (e.g., Areca catechu).

Besides this value to humans, palms are also widely employed in the landscape and in many national and international tropical gardens. The palm family offers the tall, stately Roystonea regia, the striking red petioles of Cyrtostachys renda, salt-tolerant Cocos nucifera and Phoenix dactylifera, Dypsis lutescens for hedges, and many species used in interiorscapes (e.g., Caryota, Chamaedorea, Howea, and Rhapis spp.). Specimen plants are grown in private collections and at public institutions, with impressive palm collections housed in enormous greenhouses in almost any climate. Their value is reflected in the high cost of maintaining the tropical environment required by palms in temperate areas.

In the United States, primarily in California, Florida, and Hawaii, palms are produced for the ornamental industry as potted, greenhouse-grown specimen plants or as container- and fieldgrown plants. This is in contrast to the plantation fields of palms grown for food, oil, and other commercial uses. Ornamental palms are also grown in other parts of the United States, southern Europe, Central and South America, Japan, and Australia.

While palms are naturally distributed on both sides of the equator, human activity, especially by palm enthusiasts, has transported palms to new locations atypical of their native habitat. For example, while Phoenix dactylifera is believed to have originated in the Persian Gulf area and is commonly grown in semiarid regions, it is now ubiquitous throughout subtropical Florida as landscape centerpieces. This movement from one environment to another has special consequences for palm health. For palm genera used for food, breeding programs have developed cultivars adapted to the regions in which they are grown. For ornamental palms, however, new cultivars are not bred for new environments, and palms are transported to new locations in which they might not thrive or survive.

Movement of palms to many new environments poses great challenges to ornamental palm growers. Thus, this compendium broadly addresses both nutritional and other environmental disorders as well as diseases of palms. The compendium focuses on palm production for the ornamental market and not as a plantation crop, although information developed from plantation crops, such as disease management techniques, is included.

Since ornamental crops have a high aesthetic value, spots that are of minor consequence in a plantation are a major problem for the ornamental nursery industry. Thus, prevention of leaf spots is highly crucial in ornamental palm management, and loss of a single specimen palm can be extremely costly.

Palm Anatomy and Growth Palms belong to the division of flowering plants known as monocotyledons or monocots. However, few monocot species attain the size of large palms. Thus, while palms are often referred to as “trees”, they have none of the plant characteristics common to broadleaf trees (e.g., oak, maple, poplar, and eucalyptus), which are dicotyledons. Palms have more in common with a corn plant than with an oak tree. Figure 1 is a diagram of a representative solitary (single-stemmed) palm.

Palms develop in phases. After seed germination, palms go through an establishment or juvenile phase wherein the apical meristem often remains at or below ground level. Seedling leaves are produced, followed by mature leaves. Most importantly, the palm stem increases in diameter before elongating vertically. In the adult or mature vegetative phase, the stem continues to elongate (if solitary) or basal branching occurs (if clustering). Leaves produced in this growth phase are a constant size. Some palm species do not develop a conspicuous aerial trunk for a number of years. Thus, one palm species may remain in the establishment or juvenile phase for more than 10 years (e.g., Sabal palmetto), whereas another species may develop to the point of reproduction during the same amount of time (e.g., Chamaedorea spp.). In other words, reference to palm age in years is not a good indication of palm maturity.

The palm root system develops from the stem base in the root initiation zone. Thus, all palm roots are adventitious roots.

Palm roots emerge from the stem at maximum thickness and, like the stems, are incapable of secondary thickening growth.

However, palm roots can branch, and this root branching is defined by orders. First-order roots are primarily lateral or descending. Second-order roots may ascend or descend. Third- and fourth-order roots are the primary absorbing organs and develop extensively in nutrient-rich soils. Palm roots do not produce root hairs. For many palms, especially Phoenix spp., the root initiation zone extends well above ground. Palms should not be transplanted any deeper than they were originally growing.

Otherwise, a slow decline and eventual death of the palm results.

As monocots, palm stems have no vascular cambium, a specialized layer of cells in dicots that separates vascular tissue (xylem and phloem) and from which new vascular tissue 1

arises. It is this production of new vascular tissue that increases the trunk diameter of dicot trees. In palms, the vascular tissue is in bundles scattered throughout the internal tissues of the stem (trunk). This vascular tissue survives the lifetime of the palm.

Each vascular bundle has xylem and phloem cells but no vascular cambium. Without a vascular cambium, palm stems or trunks are essentially devoid of secondary growth, meaning they produce no new vascular tissue and do not produce annual growth rings.

Therefore, palms cannot repair injuries to their stems, and diligent effort must be made to prevent injury. Wounds, such as those caused by nails or tree trimmer boot spikes, fail to heal and gradually enlarge. These wounds provide an opportunity for insect or pathogen invasion of the trunk, invasions that can be fatal.

Palms have only one apical meristem per stem. The apical meristem of a palm is often referred to as the bud or palm heart. If the apical meristem is killed, the entire palm (if solitary) or an individual palm stem (if clustering) will die. Thus, actions that might damage the apical meristem must be avoided. These include transportation damage (bouncing), insect attacks, and animal infestation (e.g., rats).

A palm stem increases in diameter before elongating. Initially, the stem consists of little more than overlapping leaf

bases protecting the apical meristem. Some palm trunks swell at the base or further up the trunk, but this is not secondary growth. As the palm matures and then elongates, what appear to be growth rings are actually leaf scars remaining after leaf abscission. Many palm stems have stemata, which are cells in the stem that contain a single silica body that provides strength or hardness to the trunk.

The leaves of palms are the largest known in the plant kingdom and are greatly admired by palm enthusiasts. Leaf production is slow, averaging about one new leaf per month. This slow

Fig. 1. The parts of a representative single-stemmed (solitary) palm. (Reprinted, by permission, from T. K. Broschat and A. W. Meerow, 2000)

Fig. 2. Three types of palm leaves: A, palmate; B, costapalmate; and C, pinnate. (Reprinted, by permission, from T. K. Broschat and A. W. Meerow, 2000)

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TABLE 1. Scientific and Common Names of Selected Palm Species Scientific Name Acoelorrhaphe wrightii (Griseb. & H. A. Wendl.) H. A. Wendl. ex Becc.

Synonyms Paurotis wrightii (Griseb. & H. A. Wendl.)

Britton Veitchia merrillii (Becc.) H. E. Moore

Adonidia merrillii (Becc.) Becc.

Archontophoenix alexandrae (F. v. Muell.) H. A. Wendl. & Drude Archontophoenix cunninghamiana (H. A. Wendl.) H. A. Wendl. & Drude Areca catechu L. Arenga pinnata (Wurmb) Merr.

Bactris gasipaes Kunth Bismarckia nobilis Hildebr. & H. A. Wendl.

Borassus flabellifer L. Brahea armata S. Watson Erythea armata S. Watson Brahea edulis H. A. Wendl. ex S. Watson Butia capitata (Mart.) Becc.

Carpentaria acuminata (H. A. Wendl. & Drude) Becc.

Caryota mitis Lour.

Caryota urens L. Chamaedorea cataractarum Mart.

Chamaedorea elegans Mart.

Neanthe bella (Mart.) O. F. Cook Collinia elegans (Mart.) Liebm. ex Oerst.

Chamaedorea seifrizii Burret Chamaedorea erumpens H. E. Moore Chamaerops humilis L. Coccothrinax argentata (Jacq.) L. H. Bailey Coccothrinax crinita (Griseb. & H. A. Wendl. ex C. H. Wright) Becc.

Cocos nucifera L. Copernicia prunifera (Mill.) H. E. Moore Cyrtostachys renda Blume Dictyosperma album (Bory) Scheff.

Dypsis cabadae (H. E. Moore) Beentje & J. Dransf.

Chrysalidocarpus cabadae H. E. Moore Dypsis decaryi (Jum.) Beentje & J. Dransf.

Neodypsis decaryi Jum. Dypsis lastelliana (Baill.) Beentje & J. Dransf.

Neodypsis lastelliana Baill.

Dypsis lutescens (H. A. Wendl.) Beentje & J. Dransf.

Chrysalidocarpus lutescens H. A. Wendl.

Elaeis guineensis Jacq.

Euterpe edulis Mart.

Euterpe oleracea Mart.

Gaussia attenuata (O. F. Cook) Becc.

Howea belmoreana (C. Moore & F. v. Muell.) Becc.

Howea forsteriana (C. Moore & F. v. Muell.) Becc.

Hyophorbe lagenicaulis (L. H. Bailey) H. E. Moore Mascarena lagenicaulis L. H. Bailey Hyophorbe verschaffeltii H. A. Wendl.

Mascarena verschaffeltii (H. A. Wendl.) L. H. Bailey Hyphaene Gaertn. spp.

Jubaea chilensis (Molina) Baill.

Latania Comm. ex Juss. spp.

Licuala grandis H. A. Wendl.

Licuala spinosa Thunb.

Livistona chinensis (Jacq.) R. Br. ex Mart.

Phoenix canariensis Chabaud Phoenix dactylifera L. Phoenix reclinata Jacq.

Phoenix roebelenii O’Brien Phoenix rupicola T. Anderson Phoenix sylvestris (L.) Roxb.

Pritchardia pacifica Seem. & H. A. Wendl.

Ptychosperma elegans (R. Br.) Blume Seaforthia elegans R. Br. Ptychosperma macarthurii (H. A. Wendl. ex H. J. Veitch) H. A. Actinophloeus macarthurii (H. A. Wendl. ex Wendl. ex Hook. f.

H. J. Veitch) Becc. ex Raderm.

Ravenea rivularis Jum. & H. Perrier Rhapidophyllum hystrix (Frazer ex Thouin) H. A. Wendl. & Drude Rhapis excelsa (Thunb.) Henry ex Rehder Roystonea oleracea (Jacq.) O. F. Cook Roystonea regia (Kunth) O. F. Cook Roystonea elata (W. Bartram) F. Harper Sabal mexicana Mart.

Sabal palmetto (Walter) Lodd. ex Schult. & Schult. f.

Serenoa repens (W. Bartram) Small Serenoa serrulata (Michx.) Nichols.

Syagrus romanzoffiana (Cham.) Glassman Arecastrum romanzoffianum (Cham.) Becc.

Cocos plumosa Hook. f.

Thrinax morrisii H. A. Wendl.

Thrinax radiata Lodd. ex Schult. & Schult. f.

Trachycarpus fortunei (Hook.) H. A. Wendl.

Veitchia arecina Becc.

Veitchia montgomeryana H. E. Moore Washingtonia filifera (Linden ex AndrĂŠ) H. A. Wendl. ex de Bary Washingtonia robusta H. A. Wendl.

Wodyetia bifurcata A. K. Irvine

Common Names Paurotis or Everglades palm Christmas or Manila palm Alexandra or King palm Piccabeen or Bangalow palm Betel nut palm Sugar or Black-fiber palm Peach or Pejibaye palm Bismarck palm Palmyra palm Blue hesper palm Guadalupe palm Pindo or Jelly palm Carpentaria palm Clustering fishtail palm Fishtail or Toddy palm Cat palm Parlor palm Bamboo or Reed palm European fan palm Silver palm Old man palm Coconut palm Carnauba wax palm Red sealing wax palm Hurricane or Princess palm Cabada palm Triangle palm Teddy bear palm Areca or Golden cane palm African oil palm Assai palm Assai palm Llume palm Belmore sentry palm Kentia or Sentry palm Bottle palm Spindle palm Gingerbread or Doum palm Chilean wine palm Latan palms Licuala palm Spiny licuala palm Chinese fan palm Canary Island date palm Date palm Senegal date palm Pygmy date palm Cliff date palm Wild date palm Fiji Island fan palm Solitaire palm Macarthur palm Majesty palm Needle palm Lady palm Caribbean royal palm Cuban royal palm Texas palm Cabbage or Sabal palm Saw palmetto Queen palm Keys thatch palm Florida thatch palm Windmill palm Montgomery palm California or Desert fan palm Mexican fan palm Foxtail palm

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rate of leaf production often confuses the diagnostician, especially as it relates to palm disorders. It is important to remember that there are several developing leaves in the apical meristem (bud) in addition to those that are visible. Damage to developing leaves in the bud may take several months to a year or longer before showing up on an emerging leaf. Likewise, the entire apical meristem may be killed, but it may take a year or longer for this to become apparent, since undamaged, alreadyformed leaves continue to emerge from the bud.

Palm leaves or fronds are produced in a multitude of shapes and sizes. However, all palm leaves consist of a blade, a petiole, and a leaf base. There are three primary classes of palm leaf blades: fan (either palmate or costapalmate), feather (pinnate or bipinnate), and entire. Leaflets on a pinnate or bipinnate palm leaf are also referred to as pinnae. Figure 2 illustrates the more common palm leaf types. While not technically correct, the petiole is often referred to as the rachis. However, “rachis” is a term reserved for the extension of the petiole into the leaf blade of feather palm leaves. In costapalmate leaves, the extension of the petiole into the blade is called the costa.

The youngest, unopened palm leaf is the spear leaf, and the entire leaf canopy is the crown.

The leaf base is also called the leaf sheath, since this petiole section sheaths or covers the stem. Senescence is the natural decline of the leaf. Some feather-leafed palms have long, tubular leaf bases that tightly wrap around each other on the stem, creating a distinctive area between the trunk and leaf canopy.

This area is called the crownshaft. Palms with crownshafts (e.g., Roystonea spp.) are “self-cleaning” palms, since the entire leaf (base, petiole, and frond) naturally drops off the trunk after senescence. Other palms naturally drop the petiole and frond, but the leaf base remains on the trunk (e.g., Sabal palmetto). Still other palms (e.g., Washingtonia spp.) require mechanical cleaning or pruning to rid the palm of naturally senesced leaves. These prunings are primarily for aesthetic reasons and management practices, such as moisture reduction, dust reduction, and removal of habitat for rodents. Occasionally, leaf prunings are required to reduce pathogen inoculum levels on host tissue. However, when the palm is also suffering from a nutrient deficiency, the reduction in inoculum level and potential disease severity must be balanced against nutrients that could be utilized by the plant from the pruned leaves.

A palm is mature when it produces its first inflorescence or flower. Depending on the species, palm inflorescences are produced either above the canopy (e.g., Brahea spp.), within the canopy (e.g., Cocos nucifera and Phoenix spp.), or below the canopy (e.g., Dypsis and Roystonea spp.). The main axis of an inflorescence is the peduncle, and the “branches” are rachillae (singular = rachilla). While palm flowers are relatively small, the fruits and seeds are usually large and, for some palms, quite conspicuous. Cocos nucifera planted along beaches and golf courses in Florida, Hawaii, and other tourist-dominated locales are routinely pruned to remove older leaves, flowers, and fruits.

This prevents injury from falling nuts or old leaves and illustrates that the precious coconut fruit of the plantation is a hazard in the urban landscape.

Palm Species There are approximately 2,700 species of palms in 200 genera that are currently placed in six subfamilies of the Palmae family. Because of the distribution of palms around the world by horticulturists (professionals and hobbyists), there are numerous common names for many palms. To avoid confusion, this compendium refers to palms by their scientific names, either genus and species or genus only. Table 1 is a list of some common palms used in the landscape. This table provides the scientific name of the palm, any known synonym scientific 4

names, and common names. For a complete classification of palms, refer to the book Genera Palmarum. For a complete list of palm genera and species, synonymous genera, and Latin naming authorities, refer to the Monocot Checklist, published by the Royal Botanic Gardens, Kew, U.K., on the Internet at http://www.rbgkew.org.uk/data/monocots.

Diseases and Disorders of Palms As monocots, palms are relatively easy to grow. However, single-stemmed (solitary-trunk) palms are also easy to kill, since these palms have a single apical meristem (bud). Once that tissue is damaged, whether from a pathogen, herbicide, nutritional deficiency, or environmental or mechanical factors, death is likely. Trunk injuries provide an entrance for insects and pathogens. Graceful, beautiful palm leaves quickly become unattractive when nutritional disorders or leaf diseases occur. Many diseases and disorders may look very similar in palms. Unlike many other plants, palms can be killed by nutritional disorders.

This includes large specimen palms such as Roystonea regia.

All pesticides must be used with great care on palms. See Physiological Disorders for information regarding potential herbicide damage, in both nursery and landscape settings. Not all fungicides have been evaluated for phytotoxicity on all species of palms. Phytotoxicity should be tested for in each situation by spraying the fungicide on a few plants of each species, rather than on the entire landscape or nursery, to ensure that the material is safe to use. Juvenile palms and young leaves are more susceptible to injury and should be included in preliminary phytotoxicity tests. The pesticide label should always be read completely and carefully. Only general fungicide recommendations are provided in the compendium, since regulations vary among countries and among states in the United States.

Palms used as ornamental plants are transported throughout the world. Interstate movement of palms across the southern third of the continental United States is common, as is movement among countries bordering the Caribbean Sea and the Gulf of Mexico. This is probably the primary method of palm pest and pathogen movement into new areas. Palm pests are transported great distances, as are other plant pests that may happen to be associated with the palm material. Thus, the quest to obtain an “instant” landscape may have long-term detrimental effects on the landscape, crop production, and environment.

Selected References Broschat, T. K., and Meerow, A. W. 2000. Ornamental Palm Horticulture. University Press of Florida, Gainesville, FL. Carpenter, J. B., and Elmer, H. S. 1978. Pests and Diseases of the Date Palm. U.S. Dep. Agric. Handb. 527.

Hartley, C. W. S. 1988. The Oil Palm, 3rd ed. Longman Scientific & Technical, Harlow, U.K. Howard, F. W., Moore, D., Giblin-Davis, R. M., and Abad, R. G. 2001. Insects on Palms. CABI Publishing, Wallingford, U.K. Neal, M. C. 1965. In Gardens of Hawaii. Bishop Museum Press, Honolulu, HI. Ohler, J. G., ed. 1999. Modern Coconut Management: Palm Cultivation and Products. Intermediate Technology Publications, London.

Tomlinson, P. B. 1990. The Structural Biology of Palms. Clarendon Press, Oxford, U.K. Turner, P. D. 1981. Oil Palm Diseases and Disorders. Oxford University Press, Oxford, U.K. Uhl, N. A., and Dransfield, J. 1987. Genera Palmarum. Allen Press, Lawrence, KS. Zaid, A., de Wet, P. F., Djerbi, M., and Oihabi, A. 1999. Diseases and pests on date palms. Pages 223-287 in: Date Palm Cultivation. A. Zaid, ed. FAO Plant Prod. Prot. Pap. 156.

(Prepared by M. L. Elliott, T. K. Broschat, J. Y. Uchida, and G. W. Simone)

Part I. Infectious Diseases Disease Caused by Algae Algae are ubiquitous in both aquatic and terrestrial environments. Some, such as blue-green algae, are classified as prokaryotes and are similar to bacteria. Others, such as green or brown algae, are eukaryotes and are similar to fungi and plants.

Algae are an important component of soils and considered a primary component in stabilizing soil aggregates. While they are often associated with water and primarily spread by water movement, algae may also be spread by air currents.

Selected References Bold, H. C., and Wynne, M. J. 1985. Introduction to the Algae: Structure and Reproduction, 2nd ed. Prentice-Hall, Inc., Englewood Cliffs, NJ. Metting, B. 1981. The systematics and ecology of soil algae. Bot. Rev. 47:195-312.

Algal Leaf Spot Algal leaf spot is commonly known as red rust because of the alga’s orange-colored reproductive structures. However, the name “red rust” is a misnomer because there is no rust fungus involved in this disease. Distribution of the disease is worldwide between latitudes 32°N and 32°S, and it occurs on many hosts outside the Palmae family. It is reported in association with various palm species from Australia, Brazil, China, the Congo region, Costa Rica, Honduras, India, Indonesia, Japan, Malaysia, Nicaragua, Nigeria, the United States (Florida, Hawaii, and Puerto Rico), and the various islands of the West Indies, but it is also likely to be present in many other countries.

Not all algae associated with palms are parasitic. Some are epiphytic, resulting in no physical damage but considered aesthetically unacceptable.

Symptoms Algal infection first appears as yellow-green pinpoint spots, primarily on the upper leaf surface and on the rachis. The algal thalli expand to 1–3 mm in diameter and coalesce, resulting in algal patches greater than 1 cm in diameter. Tissue necrosis is observed beneath the algal thallus. Premature senescence of leaves and a reduction in vigor of young palms are the observed symptoms on susceptible palm hosts, but these effects have not been quantified.

Causal Organisms Cephaleuros virescens Kunze is a green alga in the family Chroolepidaceae (synonym = Trentepohliaceae) and the primary alga species causing plant disease. This species is subaerial in growth habit. The thallus consists of a disk with a dichotomously lobed, radiating margin and may be one cell to several cells

thick. Part of the C. virescens colony grows horizontally between the leaf cuticle and epidermal cell layers. Under low magnification, the thallus appears velvety because of the presence of sterile hairs or setae and fertile branches that terminate in one to eight pedicels, each bearing one globose zoosporangium (Plate 1). Reddish orange, quadriflagellate zoospores are released when water is present (Plate 2). At reproductive maturity, the thallus appears orange because of the formation of a hematochrome pigment. Less obvious gametangia form in the subcuticular portion of the thallus. These emerge through the host cuticle and bear a single exit pore. Gametes are motile and biflagellate.

While C. virescens is the dominant pathogenic alga, other green alga species may also be involved in this disease. For example, in Malaysia, the related alga Trentepohlia sp. has been identified on palms with algal leaf spot. Cephaleuros spp. also form lichenus (symbiotic) relationships with various fungi. The lichenized forms of C. virescens are Strigula spp. (Plate 3).

Host Range and Epidemiology The pathogenic role of C. virescens has been widely documented on many crops. On palm species, however, the role of C. virescens as a pathogen or epiphyte has not always been clearly distinguished. Reports exist of the pathogenic role of C. virescens on Elaeis guineensis in the Congo region and Nigeria. Although this alga is reported on such palms as Arenga spp., Bactris gasipaes, Butia capitata, Caryota spp., Cocos nucifera, Phoenix dactylifera, Sabal palmetto, and Trachycarpus fortunei, this simply reflects the presence of the thallus, since pathogenicity has not necessarily been demonstrated.

C. virescens infection is initiated by zoospore germination and direct mechanical penetration of the leaf cuticle. Zoospore release is correlated with the humid, rainy season. Wind may disseminate sporangia for some distance. Short-distance spread is attributed to water splash of sporangia or zoospores. Vegetative growth is slow and may span 8–9 months until reproductive maturity is reached. The thallus likely remains fertile for the life of the palm leaf or rachis.

The incidence of algal leaf spot is higher on less vigorous, older tissue, particularly on palms in border rows or planted along roadsides. Suboptimal light, poor water drainage, inadequate air circulation, and other foliar diseases can predispose palms to algal leaf spot. Algal spots may exceed 20 per cm of leaf or leaflet length, and algal leaf spot is associated with up to 20% defoliation in plantation-grown E. guineensis.

Diagnostic Techniques The thallus is a reliable sign of C. virescens on palms. The role of the alga as a pathogen or epiphyte is determined by microscopic examination of freehand sections through the thallus on the host. Pathogenicity is indicated by the presence of subcuticular cells of the alga and the death of cells beneath the algal thallus. C. virescens can be cultured on modified Chu’s 5

No. 10 medium or potato dextrose agar, but cultures may take 14–21 days to become established. The alga grows as vegetative filaments and does not form reproductive structures in culture. Culturing efforts can also result in numerous isolations of such secondary fungi as Colletotrichum spp., which colonize empty algal reproductive structures.

Various saprophytic green and blue-green algae growing epiphytically on palms can be distinguished visually by their superficial, filmlike growth over leaves (Plate 4). A moist tissue easily removes this growth and exposes healthy leaf epidermis beneath the algal growth.

Management C. virescens management may be desirable at times. Cultural efforts include improved drainage and increased airflow by better plant spacing and selective pruning of overstory plants and older leaves in the palm canopy. Although copper-based fungicides have been used for C. virescens management on other crops, efficacy information for palms is unavailable.

Selected References Brunel, J., Prescott, G. W., and Tiffany, L. H., eds. 1950. The Culturing of Algae: A Symposium. Charles F. Kettering Foundation, Yellow Springs, OH.

Chapman, R. L. 1976. Ultrastructure of Cephaleuros virescens (Chroolepidaceae; Chlorophyta). I. Scanning electron microscopy of zoosporangia. Am. J. Bot. 63:1060-1070.

Chapman, R. L. 1980. Ultrastructure of Cephaleuros virescens (Chroolepidaceae; Chlorophyta). II. Gametes. Am. J. Bot. 67:1017.

Holcomb, G. E. 1986. Hosts of the parasitic alga Cephaleuros virescens in Louisiana and new host records for the continental United States. Plant Dis. 70:1080-1083.

Hsieh, H.-J. 1983. Notes on host plants of Cephaleuros virescens new for Taiwan. Bot. Bull. Acad. Sin. 24:89-96.

Joubert, J. J., and Rijkenberg, F. H. J. 1971. Parasitic green algae.

Annu. Rev. Phytopathol. 9:45-64.

Marlatt, R. B., and Alfieri, S. A., Jr. 1981. Hosts of a parasitic alga, Cephaleuros Kunze, in Florida. Plant Dis. 65:520-522.

Thompson, R. H., and Wujek, D. E. 1997. Trentepohliales: Cephaleuros, Phycopeltis, and Stomatochroon. Morphology, Taxonomy and Ecology. Science Publishers, Inc., Enfield, NH. Turner, P. D. 1976. Oil palm diseases—Introduction. Pages 421-466 in: Oil Palm Research. R. H. V. Corley, J. J. Hardon, and R. J. Wood, eds. Elsevier Scientific Publishing Co., New York.

Wolf, F. A. 1930. A parasitic alga, Cephaleuros virescens Kunze, on citrus and certain other plants. J. Elisha Mitchell Sci. Soc. 45:187205.

(Prepared by G. W. Simone)

Diseases Caused by Bacteria Bacteria are microscopic, prokaryotic organisms that are common throughout nature, including the landscape environment. Prokaryotic bacteria have no nuclei and no nuclear membranes, but a single, circular DNA molecule is present in each bacterial cell. Bacteria do have cytoplasmic membranes and cell walls. The majority of bacteria are involved in processes vital for the earth’s inhabitants. Some that benefit the landscape and field nurseries are those involved in nitrogen fixation, degradation of organic matter, nutrient recycling, and soil formation.

Of the large number of bacterial genera, only a few genera are pathogenic to plants. Although the number of bacterial genera that cause plant diseases expands, this is primarily because of their taxonomic reclassification. One of the important differences between bacteria and fungi as plant pathogens is that bacteria cannot enter the plant directly. Bacteria require a natural opening, such as stomata or hydathodes, or a wound to enter the plant and begin the infection process. Wounds may be natural, such as insect damage, or man-made, such as mechanical damage from pruning or using tree-climbing spikes.

There are only three bacterial diseases of palms, and only one is common throughout palm-growing areas. The bacterium Bacillus circulans Jordan has been shown to be pathogenic to callus tissue, meristematic tissue, and seedlings of greenhousegrown Phoenix dactylifera originating from artificially infested seeds. Thus far, a disease caused by this bacterium has not been observed in a natural field setting.

Selected References Goto, M. 1992. Fundamentals of Bacterial Plant Pathology. Academic Press, San Diego, CA. Leary, J. V., and Chun, W. W. C. 1989. Pathogenicity of Bacillus circulans to seedlings of date palm (Phoenix dactylifera). Plant Dis. 73:353-354.

Schaad, N. W., Jones, J. B., and Chun, W., eds. 2001. Laboratory Guide for Identification of Plant Pathogenic Bacteria, 3rd ed.

American Phytopathological Society, St. Paul, MN. Sylvia, D. M., Fuhrmann, J. J., Hartel, P. G., and Zuberer, D. A., eds.

1998. Principles and Applications of Soil Microbiology. Prentice Hall, Upper Saddle River, NJ.

6

Bacterial Blight Bacterial blight is a palm leaf disease with very limited host and geographic ranges. Since it occurs very infrequently, it is not a major disease of palms.

Symptoms The first symptom observed is small, water-soaked, translucent areas running along leaf veins. Mature lesions are brown to black, have a chlorotic halo, and are 1–2 mm wide and up to 50 mm long (Plate 5). The initial infection often occurs at the leaf margins through hydathodes. Leaves of all ages are apparently susceptible to infection, but leaves that are unfolding and not fully mature are more severely affected.

Causal Organism When bacterial blight was first described in 1978, the bacterium was identified as Pseudomonas alboprecipitans Rosen, but it has since been renamed Acidovorax avenae subsp. avenae (Manns) Willems et al. This bacterium is a gram-negative rod that is motile, with a single polar flagellum.

Host Range and Epidemiology Bacterial blight has been identified only on Caryota mitis in Florida (United States), and reports of its occurrence are very limited. Thus, very little is actually known about the disease.

Diagnostic Techniques After thoroughly washing leaf tissue, water-soaked lesions should be excised and crushed in sterile water or phosphate buffer. The resulting suspension is then streaked onto beef– yeast extract agar or yeast extract–dextrose–calcium carbonate (YDC) agar and incubated until discrete colonies appear. Alternatively, lesions may be directly plated onto the media. Colonies of A. avenae subsp. avenae on YDC agar are convex and dark beige and become very sticky after 3–4 days at 30–32°C. Colonies do not produce a yellow, insoluble pigment on nutri-

ent agar and are nonfluorescent on King’s medium B. Differentiation of subspecies within A. avenae is possible with diagnostic media or tests. Identification to subspecies can also be determined with commercial tests using carbon source utilization or fatty acid methyl esterase (FAME) profiles.

Management Since bacterial blight is limited in occurrence, no chemical controls have been identified. Cultural controls include removal of symptomatic leaves or the entire plant, if the palm is small. Ensuring good air circulation and eliminating overhead irrigation are also recommended to facilitate rapid drying of the leaves to limit infection and spread of the pathogen.

Selected References Knauss, J. F., Miller, J. W., and Virgona, R. J. 1978. Bacterial blight of fishtail palm, a new disease. Proc. Fla. State Hortic. Soc. 91:245-247.

Miller, J. W. 1992. Bacterial blight of fishtail palm caused by Pseudomonas avenae. Florida Dep. Agric. and Consumer Service, Div. Plant Industry. Plant Pathol. Circ. 355.

Saddler, G. S. 1994. Acidovorax avenae subsp. avenae. IMI Description No. 1211. Mycopathologia 128:41-43.

Schaad, N. W., Jones, J. B., and Chun, W., eds. 2001. Laboratory Guide for Identification of Plant Pathogenic Bacteria, 3rd ed.

American Phytopathological Society, St. Paul, MN. Willems, A., Goor, M., Thielemans, S., Gillis, M., Kersters, K., and De Ley, J. 1992. Transfer of several phytopathogenic Pseudomonas species to Acidovorax as Acidovorax avenae subsp. avenae subsp. nov., com. nov., Acidovorax avenae subsp. citrulli, Acidovorax avenae subsp. cattleyae, and Acidovorax konjaci. Int. J. Syst. Bacteriol. 42:107-119.

(Prepared by M. L. Elliott)

Bacterial Bud Rot Other common names for bacterial bud rot in the earlier Elaeis guineensis literature are lethal spear rot and little leaf.

Symptoms Initial symptoms are associated with the leaves. All or some portion of the petiole or rachis of the spear leaf exhibits brown discoloration, wet rot, or both. The spear leaf may appear wilted, chlorotic, or both. Leaves may collapse and hang from the crown or are easily pulled from the crown. If the disease progresses down into the bud, a rotted, putrid mass forms and death of the palm follows (Plates 6 and 7). If the disease does not kill the bud, the palm may recover.

Symptoms may take as long as 12 weeks to develop. The palm may recover or appear to be recovering by the emergence of abnormally short leaves (Plate 8), sometimes without leaflets. This symptom is referred to as little leaf. For palms that do recover, indicating that the bud was not affected, each new leaf that emerges is more normal in appearance.

Causal Organisms When bacterial bud rot was first described in the Republic of the Congo, the bacterium was identified as being similar to Erwinia lathryi Manns & Taubenhaus. This bacterium was renamed E. herbicola (LĂśhnis) Dye and is now combined into Pantoea agglomerans (Ewing & Fife) Gavini et al. Bacteria of the E. carotovora (Jones) Bergey et al. group have also been implicated as the cause of bacterial bud rot. However, the etiology of bacterial bud rot is confusing, since numerous bacteria, including genera normally considered saprobic, have been isolated from diseased tissue without confirmation of their pathogenicity. Furthermore, in South America, it is not clear that the disease is caused by a bacterium.

Host Range and Epidemiology A presumed bacterial bud rot has been associated with numerous palms in the United States (Florida) and with Elaeis guineensis in Colombia, Costa Rica, Democratic Republic of the Congo, Ecuador, Nicaragua, Nigeria, Panama, Republic of the Congo, and Southeast Asia. In Florida, occurrence of bacterial bud rot is normally preceded by cold weather, usually a freeze. When the disease was first described in the Republic of the Congo, it was noted that weakened palms and Elaeis guineensis cultivars not adapted to the area were most likely to succumb to the disease. In South American Elaeis guineensis plantations, poor drainage, compacted soils, and unbalanced nutrition have been suggested as predisposing factors, as has attack by insects. It is quite likely that the bacterium acts as an opportunistic pathogen.

Diagnostic Techniques Since there may be more than one causal organism involved with bacterial bud rot, tissue from the edge of the advancing rot (not the middle of the rotted tissue) should be sampled and then processed to obtain either bacterial or fungal pathogens. A microscopic mount should be made of the advancing edge of the lesion to determine whether masses of bacteria are present.

For bacterial isolation, the infected tissue should be washed well. Small pieces of tissue should then be cut, crushed in sterile water or phosphate buffer, and soaked in the diluent for up to 30 min. The resulting suspension is then streaked onto standard nutrient media (e.g., nutrient agar, tryptic soy agar, or LuriaBertani agar) and selective media. Crystal violet pectate medium is a selective medium for E. chrysanthemi Burkholder et al. and E. carotovora. The P. agglomerans group normally produces a bright yellow pigment on Luria-Bertani agar. Other diagnostic tests and media might be used depending on the bacterial genus and species of the suspected pathogen.

For fungal isolation, the infected tissue should be washed well.

Small pieces of tissue should then be cut, surface-disinfested, blotted dry, and placed onto a nonselective medium (e.g., potato dextrose agar, acidified potato dextrose agar, or water agar) and selective media for Phytophthora spp. See Phytophthora Diseases for cultivation protocols that optimize fungal spore formation and identification.

Management Resistant cultivars of Elaeis guineensis and hybrids thereof have been documented. For all other palms normally used as ornamentals, no systematic resistance studies have been conducted. In the landscape, two approaches may be appropriate.

First, when the rot is found on the exposed spear leaf, the tissue can be excised before the disease spreads into the bud tissue. Second, bud drenches with a copper-based fungicide are commonly used to protect palm buds in the southeastern United States after presumed cold damage has occurred. However, since the role of bacteria in bud rot is still unknown, the benefit of the drenches is also unknown. It is important to remember that damage to the bud may not be apparent until weeks or even months after the damage has occurred. Therefore, once the damage has become apparent, chemical treatments may be too late to be effective.

Selected References Duff, A. D. S. 1962. Bud rot disease of the oil palm. Nature 195:918919.

Gibson, I. A. S. 1979. Two important disorders of oil palm in Latin America. PANS (Pest Artic. News Summ.) 25:270-274.

Hartley, C. W. S. 1988. The Oil Palm, 3rd ed. Longman Scientific & Technical, Harlow, U.K. Schaad, N. W., Jones, J. B., and Chun, W., eds. 2001. Laboratory Guide for Identification of Plant Pathogenic Bacteria, 3rd ed.

American Phytopathological Society, St. Paul, MN.

7

Turner, P. D. 1981. Oil Palm Diseases and Disorders. Oxford University Press, Oxford, U.K.

(Prepared by M. L. Elliott)

Host Range and Epidemiology Sudden decline has only been identified on P. dactylifera in central Saudi Arabia. No information is known about the epidemiology of the disease.

Diagnostic Techniques

Sudden decline is a lethal disease of Phoenix dactylifera and has only recently been identified. The disease name reflects the short time period from initial symptom development to palm death.

Plant materials should be thoroughly washed prior to isolation. Infected tissue is then surface-disinfested and placed onto a nonselective medium (e.g., potato dextrose agar or nutrient agar). Alternatively, the tissue can be ground in buffer and the resulting suspension streaked onto a nonselective medium. The bacterium is identified with diagnostic tests and media used for identifying Erwinia spp.

Symptoms

Management

Sudden Decline

Initially, the youngest leaves of P. dactylifera appear pale green, with the discoloration moving from the tip of the leaf downward. The bases of these leaves have a brown discoloration, followed by a rotting of the tissue. Such leaves can be easily pulled from the bud. Infected trees also have roots with discolored vascular tissue. In addition, the unopened spear leaf appears wilted. The palm dies approximately 2 weeks after the spear leaf wilts, and the suckers die 1 week later.

Causal Organism Erwinia chrysanthemi Burkholder et al. has been identified as the causal organism of sudden decline.

Evaluation of local P. dactylifera cultivars in Saudi Arabia demonstrated that ‘Helwa’ and ‘Roshody’ were relatively resistant to sudden decline and that ‘Succary’ was the most susceptible.

Other management techniques have not been determined thus far.

Selected References Abdalla, M. Y. 2001. Sudden decline of date palm trees caused by Erwinia chrysanthemi. Plant Dis. 85:24-26.

Schaad, N. W., Jones, J. B., and Chun, W., eds. 2001. Laboratory Guide for Identification of Plant Pathogenic Bacteria, 3rd ed.

American Phytopathological Society, St. Paul, MN.

(Prepared by M. L. Elliott)

Diseases Caused by “fungi” The use of the small “f” in the title of this section is deliberate. The taxonomic classification of fungi has changed significantly in recent years as scientists expand their knowledge regarding the phylogenetic relationships among organisms. Previously, eukaryotic organisms composed of threadlike filaments (hyphae) that reproduced by spores and obtained nutrition by absorption were all classified within the kingdom Fungi. Today, the number of kingdoms containing fungi has expanded.

The kingdom Fungi includes the phyla Ascomycota, Basidiomycota, Chytridiomycota, and Zygomycota. The phylum Oomycota, which includes the genera Pythium and Phytophthora, is included in either the kingdom Stramenopila or the kingdom Chromista, depending on your taxonomic viewpoint. Despite these taxonomic changes, the word fungi (with a small “f”) is still used in the generic sense to refer to these organisms.

The fungi, as a generic group, are very diverse, eukaryotic microorganisms. Eukaryotes have a nucleus in the cytoplasm, which is bound by a nuclear membrane. While some fungi are single-celled organisms, most are multicellular with filamentous vegetative bodies or mycelia. The individual filaments are called hyphae, and the composition of the hyphal cell wall is a distinguishing characteristic among the fungi. Cell walls of fungi in the kingdom Fungi contain chitin, whereas cell walls of Oomycota fungi do not.

Fungi reproduce both sexually and asexually via spores.

These reproductive structures are the primary means of identifying fungi taxonomically. The nuclei of fungi in the kingdom Fungi are haploid during vegetative growth and asexual reproduction. In contrast, nuclei of the vegetative hyphae and biflagellate zoospores (asexual spores) of the Oomycota fungi are diploid, with the gametes as the only haploid cells.

Fungi acquire nutrition by absorption from external sources, primarily through hyphal cell walls. A fungus that is decomposing nonliving organic matter, usually via fungal enzymes, 8

as its source of nutrition is called a saprobe or saprotroph. The term “saprophyte” is also used to refer to saprobes, but the latter is the preferred term. When a fungus forms an association with a plant that results in a negative influence on that plant, the fungus is referred to as a plant pathogen. The majority of fungal species are saprobes, but a relatively few fungi are pathogens. As a plant pathogen, the fungus is acquiring its nutrition from the plant either by decomposing plant tissue directly or by forming special fungal organs to absorb plant cell contents.

Plant-pathogenic fungi are either facultative or obligate parasites. In general, obligately parasitic fungi do not grow and reproduce except in the presence of their plant hosts. Facultatively parasitic fungi, however, can live as saprobes for part of their life cycle.

Selected References Alexopoulos, C. J., Mims, C. W., and Blackwell, M. 1996. Introductory Mycology, 4th ed. John Wiley & Sons, Inc., New York.

Barnett, H. L., and Hunter, B. B. 1998. Illustrated Genera of Imperfect Fungi, 4th ed. American Phytopathological Society, St. Paul, MN. Dick, M. W. 2001. Straminipilous Fungi. Kluwer Academic Publishers, Dordrecht, The Netherlands.

Hanlin, R. T. 1990. Illustrated Genera of Ascomycetes. American Phytopathological Society, St. Paul, MN. Hanlin, R. T. 1998. Illustrated Genera of Ascomycetes, Vol. II. American Phytopathological Society, St. Paul, MN. Hanlin, R. T. 1998. Combined Keys to Illustrated Genera of Ascomycetes, Vol. I & II. American Phytopathological Society, St. Paul, MN. Hawksworth, D. L., Kirk, P. M., Sutton, B. C., and Pegler, D. N. 1995. Ainsworth & Bisby’s Dictionary of the Fungi, 8th ed. CAB International, Wallingford, U.K.

Sylvia, D. M., Fuhrmann, J. J., Hartel, P. G., and Zuberer, D. A., eds.

1999. Principles and Applications of Soil Microbiology. Prentice Hall, Upper Saddle River, NJ.

(Prepared by M. L. Elliott and J. Y. Uchida)

borne from the last conidiogenous cell and is pale brown and straight to slightly curved, with 7–13 septations. Conidia are 11–13 (base) to 2–3 (apex) × 50–70 µm. At the release of each conidium, the next conidiogenous cell is produced by percurrent proliferation or growth through the preexisting pore.

Host Range and Epidemiology

Annellophora Leaf Spot Annellophora leaf spot is a foliar disease limited to several important palm genera. It is reported from Malaysia, Papua New Guinea, Sierra Leone, and the United States (Texas).

Symptoms Young lesions are round to oval and brown. A narrow, yellow halo and a dark brown to black center develop as lesions age (Fig. 3). These symptoms are not distinctive from symptoms caused by Cercospora spp. or Stigmina palmivora (Sacc.)

Hughes. Numerous leaf infections cause distortion or death of palm leaves and leaflets.

Causal Organism Annellophora phoenicis M. B. Ellis is a dematiaceous hyphomycete characterized by the development of solitary or clustered (two to three) annellophores produced terminally or laterally from hyphal strands. The annellophore is brown, swollen at the base, and septate, with as many as seven successive conidiogenous cells (3–5 × 30–70 µm) (Fig. 4). A conidium is

The recorded hosts for A. phoenicis include Cocos nucifera, Phoenix canariensis, and other Phoenix spp. Disease severity is highest under conditions of high leaf wetness or foliar wounding caused by other diseases, insect pests, or environmental factors.

Diagnostic Techniques A. phoenicis reproduces from either leaf surface and readily sporulates in a moisture chamber. The pathogen can be isolated from infected tissue following standard surface disinfestation methods (e.g., 0.5% sodium hypochlorite solution) followed by culturing on potato dextrose agar. A. phoenicis colonies grow slowly and are dark brown.

Management Cultural manipulations and fungicide applications reduce leaf spot incidence and severity. Palms should be established in sites with adequate air circulation and plant spacing to minimize periods of leaf wetness. Irrigation spray should be directed away from the palm canopy in both landscape and nursery settings. During wet seasons, fungicides may be needed to minimize disease severity. Although specific efficacy data are unavailable, fungicides containing chlorothalonil, iprodione, or mancozeb may be effective. A formulation that includes a surfactant or a wettable formulation product with a surfactant added should be chosen to improve fungicide coverage and adherence to the palm canopy.

Selected References Ellis, M. B. 1958. Clasterosporium and some allied Dematiaceae— Phragmosporae. Mycol. Pap. 70, pp. 83-89. Commonwealth Mycological Institute, Kew, Surrey, U.K. Farr, D. F., Rossman, A. Y., Palm, M. E., and McCray, E. B. Fungal Databases, Systematic Botany & Mycology Laboratory, ARS, USDA, at http://nt.ars-grin.gov/FungalDatabases/DatabaseFrame.cfm.

Vann, S. R., and Taber, R. A. 1985. Annellophora leaf spot of date palm in Texas. Plant Dis. 69:903-904.

(Prepared by G. W. Simone) Fig. 3. Symptoms of Annellophora leaf spot on a Phoenix sp.

(Courtesy S. R. Vann)

Fig. 4. Conidia of Annellophora phoenicis. (Courtesy S. R. Vann)

Bipolaris and Exserohilum Leaf Spots The fungal pathogen genus Helminthosporium has been reported on many plants, and controversy over the nomenclature developed because some of the pathogens were identified as Bipolaris or Drechslera spp. by different authors. The taxonomy of this group of fungi has improved recently, and isolates can be placed in the genera Bipolaris, Drechslera, or Exserohilum, with characteristics of the anamorphic or asexual state clearly delineated for each genus. These genera also have distinct teleomorphic or sexual states associated with each anamorph. In current usage, the genus Helminthosporium is applied to relatively few fungi that are mostly associated with diseases of dicotyledons. Thus, old records of Helminthosporium spp. on palm reflect the occurrence of Bipolaris, Exserohilum, and other genera. These pathogens commonly cause leaf flecks, spots, and blights of several species of palms. Moderate numbers of spots and blights reduce marketability, decrease quality, and increase crop production time.

9

Symptoms Leaf spots caused by Bipolaris spp. are initially small, watersoaked lesions that develop within 24 hr following inoculation with a spore suspension. Brown, pinpoint lesions develop in 2– 4 days. After 1 week, spots on young leaves are brown, elliptical (oval) to circular, 1 mm in diameter, and surrounded by a chlorotic band (Plate 9), while spots on mature, expanded leaves are brown, circular to irregularly elliptical, up to 5 mm long, and surrounded by a diffuse chlorotic zone (Plate 10). The oldest leaves have spots that are brown and 1 mm in diameter with a water-soaked zone about 1.5 mm wide. After 1 month, spots on the youngest leaves can develop into blights (Plate 11). Spots on the mature leaf are multi-ringed with a brown center surrounded by greenish yellow tissue, which is followed by a reddish brown ring, and at times, by an asymmetric, chlorotic zone that may be 20 mm long or longer (Plate 12).

Inoculation with these pathogens on various palm species demonstrates that lesion type is of limited value for identifying some pathogens. Inoculation of B. incurvata (C. Bernard) Alcorn on Howea forsteriana resulted in varying symptoms based on leaf age. The youngest leaves, which were expanded but still tender and very flexible at inoculation, were the most susceptible with 50 to more than 100 spots per leaflet, while older, dark green, firm leaves had 10 or fewer spots per leaf.

Leaf spots produced by an Exserohilum sp. on a Dypsis sp. were also variable. However, B. hawaiiensis (M. B. Ellis) J. Y. Uchida & Aragaki, isolated from a Chamaedorea sp. and inoculated on a Dypsis sp., caused hundreds of flecks, but no spot was larger than 1 mm. Leaf spots are also caused by Phaeotrichoconis crotalariae (M. A. Salam & P. N. Rao) Subramanian on several palm species.

Causal Organisms Several Bipolaris and Exserohilum spp. are pathogenic to palms. Both genera are hyphomycetes that produce darkcolored asexual spores or conidia. These spores are useful for accurate identification of these fungi and are relatively easy to obtain. All descriptions provided below are from cultures grown on V8 juice agar under approximately 2,500-lx continuous light from cool-white fluorescent lamps.

E. rostratum (Drechs.) K. J. Leonard & E. G. Suggs produces long (up to 200 µm), obclavate, rostrate, light gray-black to olivaceous spores with up to 18 pseudosepta in the light at 20°C. Spore diameter is about 13 µm at the widest point (Plate 13). Spores formed in the dark average 100 µm long and are elongate-fusoid with rounded ends and darker in color. Regardless of light conditions for conidia formation, spores have a protuberant hilum, are pseudoseptate, and appear to be divided into several cells. Intermediate or mixed spore morphologies are produced with diurnal light changes. Maintaining cultures in continuous light reduces variation in spore morphology and aids accurate identification.

B. incurvata produces numerous conidiophores at 24°C in the light, and spores are induced to form by transferring cultures to 20°C in the light for 12 hr. Conidia are curved and fusoid and taper to a rounded apex. Spores are gray to dark olivebrown and average 17 × 130 µm with 6–14 pseudosepta (Plate 14).

At 24°C in the light, B. setariae (Sawada) Shoemaker produces conidia that are mostly curved, fusoid to elongateellipsoid with a rounded apex and base, mostly 10–13 × 50–70 µm with five to nine pseudosepta, and dark olive-brown at maturity. Conidia formed in the dark are elongate-ellipsoid, 10–14 × 40–60 µm with four to nine pseudosepta, and grayish olivebrown at maturity.

B. cynodontis (Marignoni) Shoemaker produces slightly curved, fusoid conidia with tapered blunt ends. Conidia have seven to eight pseudosepta, are pale to mid-golden brown with thin smooth walls, and average 30 × 50 µm for those formed in the light.

B. hawaiiensis produces conidia that are straight, ellipsoidal, oblong or cylindrical, rounded at the ends, and brown to dark brown. Conidia have mostly three to six pseudosepta and average 8 × 24 µm for spores formed in the light at 24°C. It is the smallest Bipolaris species commonly found on palms.

P. crotalariae produces unbranched, straight or flexuous, brown, smooth conidiophores with solitary, obclavate, rostrate, septate conidia that are 14–22 × 50–85 µm. Each conidium has a golden brown body with a large brown scar and a long, narrow, hyaline to pale brown, thin-walled beak (Plate 15).

Host Range and Epidemiology TABLE 2. Palm Genera Reported as Hosts of Bipolaris, Exserohilum, or Phaeotrichoconis spp.

Palm Genus Adonidia Archontophoenix Areca Carpentaria Caryota Chamaedorea Chamaerops Cocos Dypsis Elaeis Geonoma Gronophyllum Howea Latania Licuala Livistona Phoenix Pritchardia Ptychosperma Ravenea Rhapis Roystonea Syagrus Veitchia Washingtonia a The

Bipolaris

Exserohilum

Phaeotrichoconis

+ + – – + + – + + + – – + – + + + + + + + + + – +

±a + + + + + – – + – + + + + – – – – + – + + + + +

– – – – + + + – + – – – – – – + + – – – + – + – +

host was reported as a Veitchia sp., which may have included Veitchia merrillii, renamed as Adonidia merrillii.

10

Bipolaris, Exserohilum, and Phaeotrichoconis spp. have been associated with a number of palm genera (Table 2). Bipolaris spp. are common on palms throughout the world. B. incurvata has been reported as the causal organism of Cocos nucifera leaf spots and blights from Fiji, French Polynesia, India, Jamaica, Malaysia, the Philippines, Seychelles, Thailand, Vietnam, and many other countries. In the United States, B. incurvata is a pathogen of Cocos nucifera, Dypsis lutescens, and Howea forsteriana. B. setariae has been reported on Caryota mitis, Chamaedorea elegans, Dypsis lutescens, Phoenix roebelenii, Phoenix canariensis, Cocos nucifera, Howea forsteriana, Livistona chinensis, Ravenea rivularis, a Rhapis sp., and a Roystonea sp., with pathogenicity confirmed on the first four palms. B. cynodontis has been reported on Adonidia merrillii and Dypsis lutescens. B. hawaiiensis has been isolated from Dypsis lutescens and from Elaeis guineensis in India. Bipolaris spp. have been reported on Licuala ramsayi (F. v. Muell.)

Domin., Livistona chinensis, a Pritchardia sp., Ptychosperma elegans, Ravenea rivularis, Syagrus romanzoffiana, and Washingtonia robusta. Worldwide, other Bipolaris spp. attacking the Palmae family are B. australiensis (M. B. Ellis) Tsuda & Ueyama, B. maydis (Nisikado & Miyake) Shoemaker, B. melinidis Alcorn (synonym = B. curvispora (El Shafie) Sivanesan), and B. zeicola (G. L. Stout) Shoemaker.

E. rostratum is reported on most of the same palms as B. setariae, but it is also reported on Archontophoenix alexandrae, Areca catechu, Carpentaria acuminata, Caryota mitis, Chamaedorea seifrizii, Geonoma interrupta (Ruiz & Pav.) Mart., a

Latania sp., Ptychosperma macarthurii, Rhapis excelsa, Roystonea regia, Syagrus romanzoffiana, a Veitchia sp., and a Washingtonia sp. Exserohilum spp. have been reported on Areca catechu, a Howea sp., Roystonea elata, an unidentified Roystonea sp., and Syagrus romanzoffiana. Pathogenicity of E. rostratum has been confirmed on Dypsis lutescens, where foliar blights and spots were produced in 2 weeks, while flecks and spots were produced on a Latania sp. For the three isolates obtained from Dypsis lutescens, a Latania sp., and a Dendrobium orchid, pathogenicity on Dypsis lutescens and the Latania sp. was similar. In greenhouse tests in Florida and Australia, leaf spots were produced by E. rostratum on Archontophoenix cunninghamiana, Caryota mitis, Chamaedorea elegans, Chamaedorea seifrizii, Dypsis lutescens, a Gronophyllum sp., Ptychosperma elegans, Rhapis excelsa, Roystonea regia, and Syagrus romanzoffiana, while a Bipolaris sp. (species not given) caused spots on Archontophoenix alexandrae, Archontophoenix cunninghamiana, Caryota mitis, Chamaedorea seifrizii, Cocos nucifera, Dypsis lutescens, Howea belmoreana, Howea forsteriana, Licuala ramsayi, Ptychosperma elegans, a Rhapis sp., and Washingtonia robusta.

In Hawaii, isolations from nearly 250 flecks and spots on Dypsis lutescens showed that 48% were aborted infections (no organism grew from these spots), 24% yielded Colletotrichum gloeosporioides (Penz.) Penz. & Sacc. in Penz., 23% yielded B. incurvata, and 6% yielded other fungi (Alternaria alternata (Fr.:Fr.) Keissl., a Nigrospora sp., a Phoma sp., etc). In the literature, both C. gloeosporioides and B. incurvata are viewed as secondary invaders. When isolates of both species were tested, only B. incurvata consistently caused leaf spots and blights, while isolates of C. gloeosporioides were not pathogenic. Only one isolate of C. gloeosporioides produced a few flecks. Similar results have been obtained on Cocos nucifera for B. incurvata and Pestalotiopsis spp., where B. incurvata caused leaf spots that were secondarily invaded by Pestalotiopsis spp.

The genus Helminthosporium has been reported on more than 40 palm species, which, as noted previously, probably represents the presence of Bipolaris or Exserohilum spp. The genus Drechslera is also reported on a few palm species, but since these reports are on palms for which Bipolaris or Exserohilum spp. have been reported, the Drechslera sp. listed may belong to either genus. Except for one listing of D. setariae (Sawada) Shoemaker, all others are listed as Drechslera species.

P. crotalariae and an unidentified Phaeotrichoconis sp. have been associated with leaf spots of Caryota mitis, Caryota urens, Chamaedorea elegans, Chamaerops humilis, Dypsis lutescens, Livistona chinensis, Phoenix canariensis, Rhapis excelsa, Syagrus romanzoffiana, Washingtonia robusta, and an unidentified Washingtonia sp. Pathogenicity has been confirmed for Caryota urens, Chamaedorea elegans, Dypsis lutescens, and Rhapis excelsa.

The ascospores of the sexual stage of Bipolaris and Exserohilum spp. are easily blown by air currents but are almost never observed in nature. However, asexual spores (conidia) are formed on blighted tissue and rotted stems in humid environments.

These pathogens depend on rapid conidia formation on the surface of diseased plants to produce huge numbers of spores that spread disease. Fungal growth and sporulation, spore germination, and leaf penetration are all dependent on high humidity or leaf wetness levels. Small spots expand into blights that produce spores that repeat the cycle. Many small spots fail to expand and other fungi, such as A. alternata, C. gloeosporioides, a Nigrospora sp., and a Phoma sp., colonize these spots. Many saprobic fungi commonly invade the dead tissue.

Diagnostic Techniques This group of fungi causes a range of leaf and petiole symptoms that includes light to dark brown spots, spots with chlorotic zones, spots with reddish brown zones, and spots

with long chlorotic streaks. The type of spot depends on leaf maturity at infection and on environmental conditions, with infections of young leaves and at high humidity resulting in the most severe blights and leaf destruction. Both Bipolaris and Exserohilum spp. commonly sporulate on the surface of blighted tissue. If spores are absent, these fungi can be induced to sporulate by incubating the specimen in a moist chamber (dish or plastic bag with moist paper towel) in the light at 20– 24°C for 1–2 days.

Isolation is relatively simple when using the following procedures. Leaf specimens should be washed well by rubbing them under running tap water. A small amount of liquid detergent should be used for washing flecks and spots. Washed specimens are placed on a paper towel. Each leaf spot is cut into 2 × 5-mm sections containing some healthy tissue and part of the spot. Cut specimens are immersed in a 0.5% sodium hypochlorite solution amended with a small amount of detergent, which serves as a wetting agent. Immersion time should be no more than 30–60 sec. The tissue is blotted dry and placed on the surface of water agar plates. The plates are incubated at or below 24°C in the light.

Mycelia of this group of fungi develop rapidly and generally emerge by the second day. Growth is vigorous and hyphal tip transfers can be made. After 4–7 days, conidia are formed on the water agar surface and on the specimen. Cultures initiated with single spores are highly recommended. Spores can be transferred with a sterile glass needle or scalpel tip to water agar, spread apart, allowed to germinate for 3 hr, and then transferred individually to dilute potato dextrose agar or V8 juice agar.

Most species grow well and produce abundant spores on V8 juice agar. All cultures should be grown between 20 and 24°C under continuous fluorescent light. Spores should form in 5–7 days. For isolates that do not form spores readily, 5-day-old cultures should be placed at 20°C in the light for 24 hr. If V8 juice agar is not available, water agar on which autoclaved palm leaf pieces have been placed should be used. These fungi readily produce spores on the host tissue and surrounding agar.

Management Restricting the entry of causal organisms into production greenhouses and nurseries can prevent diseases on potted palms. All plants should be disease free as they enter the production area. If plants with spots are purchased or if disease is already present, all large spots or blights must be removed and destroyed. Diseased plant tissue should not be placed in a compost pile, because the spores will blow to healthy plants. If only a few plants are diseased, all infected plants should be isolated from other clean, healthy plants. Bipolaris and Exserohilum spp. are excellent pathogens of many other nonpalm hosts. Since high inoculum levels can be produced on weeds surrounding the greenhouse or nursery, elimination of weeds in general, but especially grasses, is recommended.

Use of general purpose contact fungicides such as maneb or mancozeb reduces disease levels. Good coverage is important when contact fungicides are used. These fungicides prevent pathogen spore germination and growth; thus, incomplete coverage allows some of the fungal spores to penetrate the leaf. It is the most cost effective to use fungicides in combination with effective sanitation practices that reduce inoculum levels.

Efforts should be made to reduce humidity, because this will decrease the number of spores produced, the number of spores that germinate, and the number of spores that penetrate the host. Spacing between plants should be increased to promote rapid drying of the foliage and to attain good fungicide coverage. Overhead irrigation should be avoided. Drip irrigation or low, below-canopy (i.e., below the palm leaves) sprinklers should be used for large plants. Plants should be irrigated in the morning to avoid leaf wetness at night. Solid-covered greenhouses are recommended for high-rainfall environments.

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As with any disease, plants should be protected from abiotic damages that create a weakened host. Fertilization programs that are imbalanced, damage from pesticides and heat or cold injury, and leaf necrosis caused by water stress should be avoided. Wounded or weakened tissue allows these pathogens to form a colony in the canopy, which serves as the focal point of disease spread.

Selected References Alcorn, J. L. 1983. Generic concepts in Drechslera, Bipolaris and Exserohilum. Mycotaxon 17:1-86.

Alcorn, J. L. 1988. The taxonomy of “Helminthosporium” species.

Annu. Rev. Phytopathol. 26:37-56.

Alfieri, S. A., Langdon, K. R., Kimbrough, J. W., El-Gholl, N. E., and Wehlburg, C. 1994. Diseases and Disorders of Plants in Florida.

Florida Dep. Agric. and Consumer Service, Div. Plant Industry.

Bull. 14.

Chase, A. R. 1982. Leaf spot disease of Areca and other palms. Foliage Dig. 5:4-5.

Chase, A. R. 1982. Dematiaceous leaf spots of Chrysalidocarpus lutescens and other palms in Florida. Plant Dis. 66:697-699.

Forsberg, L. I. 1987. Diseases of ornamental palms. Qld. Agric. J. 113:279-286.

Management Since this disease is new, management techniques are not fully developed. Protective fungicides may be useful when the disease is most prevalent. Wound prevention of the leaf tissue may be useful.

Selected References Barnett, H. L., and Hunter, B. B. 1998. Illustrated Genera of Imperfect Fungi, 4th ed. American Phytopathological Society, St. Paul, MN. Polizza, G. 2002. Severe outbreak of leaf spot and blight caused by Botrytis cinerea on majesty palm in southern Italy. Plant Dis. 86: 815.

Polizza, G., and Vitale, A. 2003. First report of leaf spot and blight caused by Botrytis cinerea on pygmy date palm in Italy. Plant Dis. 87:1398.

(Prepared by M. L. Elliott)

Calonectria Leaf Spot (Cylindrocladium Leaf Spot)

(Prepared by J. Y. Uchida)

Botrytis Leaf Spot and Blight Botrytis leaf spot and blight is a new disease that has been observed on a limited number of palm species in Italy.

Several species of the ascomycete fungal genus Calonectria cause leaf spots on palms. For some Calonectria species, the anamorphic or imperfect state, which are all Cylindrocladium spp., is more likely to be observed causing leaf spots.

Symptoms

Botrytis cinerea Pers.:Fr. is the causal organism of Botrytis leaf spot and blight. Conidia formed on potato dextrose agar are one-celled, ovoid, and 5.2–10 × 7–14 µm. Sclerotia are black and irregular in shape and size.

Leaf spots caused by this group of fungi are generally grayish brown, dark brown, or nearly black (Plate 16). They are often gray in the center and may have a brownish black edge.

Young spots are brown, circular (0.5 mm in diameter), and frequently surrounded by a chlorotic zone about 1 mm wide. As spots expand, the typical gray centers with brown edges develop (Plate 17). Spots are nearly circular to elliptical, up to 3 × 5 mm. With time, the edges become more irregular. Sporulation of the fungus on older lesions and reinfection of the leaf form a few larger spots with small spots and flecks nearby. The rachis and petiole are also infected with similar dark lesions.

Advanced stages of the disease are characterized by coalescing spots and chlorosis and necrosis of broad bands of the leaf, especially the margins and tips. Entire leaflets or leaves may be lost as they desiccate from lesion expansion.

Host Range and Epidemiology

Causal Organisms

This pathogen has been reported only from Italy on three palm species, Phoenix canariensis, P. roebelenii, and Ravenea rivularis. Because the disease is new, little information is known about its epidemiology. The disease has been observed on palms growing in both greenhouses and field nurseries, but only during the winter months of December through March during cool, humid weather. For R. rivularis, only palms that are 3 years old or younger develop symptoms. The disease appears to be more severe on wounded palms.

Calonectria colhounii Peerally, C. ilicicola Boedijn & Reitsma (synonym = C. crotalariae (C. A. Loos) D. K. Bell & Sobers), and C. theae C. A. Loos cause leaf spotting and blights as described above. The anamorphs for all three species are a Cylindrocladium sp. Leaf spots caused by these fungi are nearly indistinguishable. The conidiophores or spore-bearing structures are distinctive for Calonectria spp., with spore-producing phialides formed on penicillate branches that arise laterally from the main axis of the conidiophore (Plate 18). The conidiophore axis is frequently very long and terminates in a vesicle. The shape and size of this vesicle, which varies with the species, are narrow and clavate (clublike) for C. theae (Plate 19) and C. colhounii, while those of C. ilicicola (Plate 20) are sphaeropedunculate in shape (spherical and tapering toward the base). Conidia of all three species primarily have three septa (Plate 21).

Most isolates of C. theae also produce distinctive macroconidia that are large, angular spores with 4–15 cells (Plate 22).

Macroconidia have a right-angle bend near the center of the spore. These macroconidia are formed in water agar isolation plates after a few days. The teleomorphic or sexual stage is characterized by red to reddish brown perithecia for C. theae and C. ilicicola and by yellow to yellowish brown perithecia

Symptoms The disease initially appears as small, chlorotic spots on leaf tissue. Depending on the host, these expand into necrotic spots with a gray mold or into spots with brown or gray, wrinkled, sunken centers and yellow or reddish brown margins. Spots may enlarge and coalesce to cause blights of young expanding leaves. Death of apical buds is possible.

Causal Organism

Diagnostic Techniques Conidiophores and conidia can be directly observed by examining leaf tissue exhibiting gray mold symptoms. Alternatively, the conidia can be scraped from the leaf tissue into sterile water and streaked onto potato dextrose agar. Tissue isolations are made by surface-sterilizing small pieces of necrotic leaf tissue in 1% HgCl2 for 20 sec or 1% sodium hypochlorite for 60 sec, rinsing in sterile water, and then placing on potato dextrose agar or malt extract agar. After 2 days, single hyphal tips can be transferred to potato dextrose agar. Colonies are initially hyaline on the medium but become gray to brown as conidia develop.

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for C. colhounii. Both C. theae and C. ilicicola have eight ascospores per ascus (Plate 23), while C. colhounii has four ascospores per ascus. All three species are homothallic, and C. ilicicola forms perithecia readily in culture and on diseased host tissue.

Host Range and Epidemiology There are only a few published reports of Calonectria or Cylindrocladium spp. as pathogens of palms worldwide. In Australia, naturally infected Howea forsteriana, Ptychosperma elegans, and Washingtonia robusta have been reported, and pathogenicity of Cylindrocladium floridanum Sobers & C. P. Seym. to H. forsteriana and W. robusta has been confirmed.

Disease also developed on Caryota mitis, Chamaedorea elegans, Dypsis lutescens, H. belmoreana, and P. elegans inoculated with Cylindrocladium floridanum.

The disease is also reported from the United States. In Tennessee, leaf spots of W. robusta were caused by Cylindrocladium pteridis F. A. Wolf. Isolates of Cylindrocladium pteridis from W. robusta, Eucalyptus cinerea F. v. Muell. ex Benth., and Rumohra adiantiformis (G. Forst.) Ching were equally pathogenic to Chamaedorea elegans, H. belmoreana, and H. forsteriana in Georgia. In Florida, Cylindrocladium pteridis was observed from leaf spots on Chamaedorea elegans, Cocos nucifera, H. belmoreana, H. forsteriana, W. robusta, and an unidentified Washingtonia sp. In Hawaii, C. theae, C. colhounii, and C. ilicicola have been confirmed as pathogens of H. forsteriana without wounding. Chamaedorea elegans, Dypsis lutescens, and Laccospadix australasica H. A. Wendl. & Drude were less susceptible than H. forsteriana to C. theae and C. colhounii. Both a seedling blight of Laccospadix australasica, characterized by severe root and petiole rots, and moderate root rots of H. forsteriana have been associated with C. ilicicola in Hawaii. Cylindrocladium scoparium Morg. was associated with leaf spots on Dictyosperma album, H. forsteriana, Livistona chinensis, and W. robusta in Florida. Also from Florida, Cylindrocladium spp. were associated with leaf spots of Dypsis lutescens, H. belmoreana, H. forsteriana, and Syagrus romanzoffiana and root rots of Chamaedorea elegans, Chamaerops humilis, and Mauritia flexuosa L. f.

Disease cycles generally involve the anamorphic or conidial stages of the pathogens, but several species produce perithecia and ascospores readily. Conidia are dispersed by insects, tools, gloves, plant handling, air movement, and splashing water. Ascospores are discharged from the perithecia and spread by air currents. The perithecia of these pathogens are commonly formed on diseased tissue. Compared with conidial distribution, ascospores travel much farther and can spread disease throughout the entire greenhouse. Ascospores also gather at the perithecial opening or ostiole and are splash-distributed. Conidia and ascospores land on host tissue and germinate with available moisture. Wet leaves or high relative humidity are conducive to spore germination and subsequent penetration of the host. The pathogen grows within the host and, after 1 week, produces conidiophores on the leaf or petiole surface and forms conidia.

Some species, such as C. ilicicola, produce microsclerotia that survive in host tissue, soil, or potting mix for long periods of time. These germinate as host roots penetrate infested potting mix or soil. Movement of the pathogen from roots to the collar allows production of conidia or ascospores on aerial parts of the palm, thus initiating the aerial disease cycle. Longdistance spread occurs as diseased plants or seeds are transported between states or countries.

Alternative nonpalm hosts are common and, since ornamental nurseries grow a mixture of crops, other diseased plants may serve as sources of inoculum. For example, C. ilicicola is a pathogen of Anthurium andraeanum Linden, Cissus rhombifolia Vahl, Dracaena marginata Lam., Leea coccinea Planch., and Schefflera actinophylla (Endl.) Harms.

Diagnostic Techniques Leaf spots and blights caused by Calonectria spp. are similar to those caused by Bipolaris and Exserohilum spp. Examination of blights and older lesions often reveals spores of either pathogen. If spores are not present, diseased leaves should be cleaned and placed in a moisture chamber, such as a petri dish with moist tissue paper, and sporulation should occur in 1 or 2 days.

Calonectria spp. can be easily isolated from leaf spots. Leaf sections with spots must be washed well with soap and rinsed in running tap water to reduce surface contaminants and saprobes that make isolation difficult. Leaf spots are cut into small sections (2 × 5 mm) and disinfested by placing them in a 0.25– 0.5% sodium hypochlorite solution with a small amount of detergent as a wetting agent. After a few seconds, the specimen pieces are removed, blotted dry, and transferred to water agar or acidified potato dextrose agar. Plates should be maintained at 24–26°C in the light and checked daily for fungal growth.

Mycelial growth of Calonectria spp. emerges within the first 3 days. It is septate and usually grows faster than other fungi, such as Colletotrichum spp. Colony growth on acidified potato dextrose agar is cream to white on the upperside and brown on the underside. If a Calonectria sp. is suspected, different types of mycelia should be selected on the first or second day following plating of the specimen and hyphal tips should be aseptically transferred to potato dextrose agar or V8 juice agar. If individual hyphal tips cannot be selected and dissected out, several of the same type should be cut and placed on V8 juice agar or potato dextrose agar. These plates should be incubated at 24–26°C in the light, and conidia should form in 4–6 days. Germinating conidia on water agar allows for establishment of single-spore cultures. After several days, conidia sometimes form on the water agar used for isolation purposes, either from mycelial growth on the water agar or directly on the leaf tissue. Large, angular macroconidia indicate the presence of C. theae.

For macroconidia formation, agar plugs from a pure culture can be placed on water agar and incubated at 24°C in the light for 2 weeks. C. theae and C. colhounii produce perithecia on autoclaved palm leaves placed on the surface of water agar and incubated at 24–25°C for at least 1 month.

Management As with many diseases caused by hyphomycetes, distribution of the fungus is dependent on the production and distribution of conidia of the pathogen. Hyphomycetes that have sexual stages that readily produce ascocarps also release ascospores that are airborne. Potential for spread is much greater when the sexual stage is produced. Conidia or ascospore germination and penetration of the pathogen into the plant require moisture. Thus, moisture control is crucial to management of disease development and spread. Efforts should be made to keep leaves dry.

These should include depending less on overhead irrigation, properly timing irrigation to avoid wet foliage at night, using solid-covered greenhouses for nurseries in high-rainfall environments, increasing space between plants and trimming surrounding vegetation to promote air movement, properly distributing plant types within the greenhouse (i.e., avoiding placement of large plants with thick growth at the site where air enters the greenhouse), and implementing any other cultural practices that will reduce humidity.

Reducing the inoculum is also crucial. Sanitation practices that remove diseased plants should be continually practiced.

The crop should be examined every few days for leaf spots. All severely diseased leaves and plants should be removed. Diseased leaves should be trimmed and leaf bases (sheaths) removed if possible. If only a few plants are diseased, they should be gathered and isolated from healthy plants. Weeds should be removed from pots and the greenhouse. If possible, palms should not be grown with crops known to be susceptible to Calonectria diseases.

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The crop can be protected from pests by controlling insects, snails, and slugs that spread disease or that injure plants, providing avenues for pathogen entry. Damage to host plants from water stress, sunburn, fertilizer burn, herbicide phytotoxicity, and heat or cold injury should be avoided, since damaged tissue can be invaded by pathogens.

Disease-preventive fungicides may be applied, but all label directions should be followed. Thiophanate methyl and maneb are effective against the listed Calonectria spp.

Selected References Alfieri, S. A., Langdon, K. R., Kimbrough, J. W., El-Gholl, N. E., and Wehlburg, C. 1994. Diseases and Disorders of Plants in Florida.

Florida Dep. Agric. and Consumer Service, Div. Plant Industry.

Bull. 14.

Crous, P. W. 2002. Taxonomy and Pathology of Cylindrocladium (Calonectria) and Allied Genera. American Phytopathological Society, St. Paul, MN. Forsberg, L. I. 1987. Diseases of ornamental palms. Qld. Agric. J. 113:279-286.

Uchida, J. Y., and Aragaki, M. 1992. Calonectria leaf spot of Howeia forsterana in Hawaii. Plant Dis. 76:853-856.

Uchida, J. Y., and Aragaki, M. 1997. Comparative morphology and pathology of Calonectria theae and C. colhounii in Hawaii. Plant Dis. 81:298-300.

(Prepared by J. Y. Uchida)

Colletotrichum Leaf and Fruit Spot The genus Colletotrichum has been associated with more than 45 palm species, with 31 of these also reported in association with the teleomorphic or sexual stage, Glomerella cingulata (Stoneman) Spauld. & H. Schrenk. Compared with the large number of palm species that is associated with Colletotrichum spp., confirmations of pathogenicity are few. Disease fails to develop following inoculation with several Colletotrichum isolates collected from leaf spots of palm. Therefore, the role of Colletotrichum spp. as the causal organisms of leaf diseases on a large number of palm species needs to be reevaluated.

Symptoms A serious leaf spot of Cyrtostachys renda was discovered in the United States (Hawaii) in 1994. Leaf spots begin as small, water-soaked, green areas about 1–2 mm wide, following inoculation of nonwounded leaves (Plate 24). These diseased areas expand into circular spots with tan to light brown centers, bordered by water-soaked tissue. As spots expand, lesion centers become very light tan to cream, with a brown margin around some spots (Plate 25). Circular spots are mostly 3–7 mm wide, and the central, necrotic areas enlarge as spots coalesce (Plate 26). Young leaves are very susceptible and develop large spots, while mature leaves develop fewer and smaller spots.

With adequate moisture, the fungus forms conidia on older lesions, resulting in new, small flecks and spots developing near the black edges of the larger, older spots. On mature leaves, these tiny spots do not expand and the leaf can be covered with hundreds of tiny flecks and spots. With heavy infection, there is general chlorosis of the entire leaf. Chlorosis may also occur around individual spots. Petioles and leaf bases (sheaths) are also infected, and typical spots are 5–10 mm long and brown to gray with dark brown to black edges (Plate 27). Heavy spotting and blighting cause leaf death, and the plant appears thin and unthrifty. Moderate disease levels greatly reduce the quality of the plant.

Colletotrichum gloeosporioides (Penz.) Penz. & Sacc. in Penz. is known to infect immature Serenoa repens fruits in the 14

United States (Florida). In the field, the fungus caused lesions on flowers and fruits and abscission of fruits, which caused 100% fruit loss in central and southern Florida in 1997. Small, black lesions formed on Serenoa repens fruits 21 days after inoculation, and the fruits were completely blackened after 35 days, followed by fruit abscission (Plate 28). Acervuli were present in fruit lesions.

Colletotrichum spp. are also associated with leaf spots of many other palms. This group of fungi has been associated with tiny black flecks to brown spots of various sizes to large chlorotic or necrotic lesions of various shapes. Lesion centers are generally tan to dark with chlorotic halos. Young seedlings are considered especially susceptible.

Causal Organisms In general, Colletotrichum spp. produce subepidermal acervuli with simple conidiophores and hyaline, single-celled conidia. The conidia of the Cyrtostachys renda pathogen are slightly smaller than other C. gloeosporioides isolates collected from many tropical hosts. However, given the broad species description of J. A. von Arx, this Cyrtostachys renda isolate fits within C. gloeosporioides. The genus Glomerella Spauld. & H. Schrenk., the teleomorphic state, has not been observed on the host or in culture. The isolates pathogenic to Serenoa repens were also identified as C. gloeosporioides. C. dematium (Pers.) Grove also has been associated with palm leaf spots in Florida.

More than 60 species of palm have had leaf spots associated with the genus Gloeosporium Desmaz. & Mont. Colletotrichumlike fungi that have no setae were often identified as Gloeosporium spp. A taxonomic review has placed most of these Gloeosporium spp. into Colletotrichum and other genera. There are cultures of Colletotrichum spp. that do and do not produce setae, but this characteristic is not used to separate genera. Environmental factors such as nutrients and humidity can affect setae production.

Host Range and Epidemiology C. gloeosporioides was confirmed as the causal organism of a severe flower and fruit drop of Serenoa repens in the United States (Florida), a fruit loss of Areca catechu in India, a foliar disease of Cyrtostachys renda in the United States (Hawaii), and leaf spots on Phoenix roebelenii in the United States (Florida). Given the clear pathogenicity of the isolates from Cyrtostachys renda, Phoenix roebelenii, and Serenoa repens, Colletotrichum spp. isolated at high frequencies from diseased palm tissue should be tested for pathogenicity, especially if other common leaf pathogens such as Bipolaris, Calonectria, or Phytophthora spp. are not present. Inoculation of Dypsis lutescens, Chamaedorea seifrizii, Howea forsteriana, and Rhapis excelsa with the Cyrtostachys renda isolates of C. gloeosporioides did not cause significant spotting on these hosts. Inoculated leaves remained clean or a few small spots were formed by one isolate. However, tomato and eggplant fruits developed water-soaked lesions in 3 days when inoculated with the Serenoa repens isolate of C. gloeosporioides.

The ascospores of Glomerella spp. are discharged into the air and disseminate the fungi over great distances. Colletotrichum spp. are excellent saprobes, growing well on necrotic tissue and rapidly producing conidia. Thus, these ubiquitous fungi spread easily. This may explain why nearly all Colletotrichum spp. isolated from palm leaf spots in Hawaii have been nonpathogenic. C. gloeosporioides was isolated from 24% of about 250 flecks and spots of Dypsis lutescens; 23% of the spots produced Bipolaris incurvata (C. Bernard) Alcorn. Except for one isolate that caused a few flecks, all Colletotrichum isolates tested were not pathogenic, while Bipolaris isolates were consistently and highly pathogenic. It is likely that, for some leaf spots, Colletotrichum spp. are invading spots caused by other pathogens. In general, Colletotrichum isolates forming

Selected References

the Glomerella (teleomorphic) state have not been pathogenic on numerous other hosts.

Diagnostic Techniques Colletotrichum spp. are easily isolated on water agar. The diseased leaf area should be thoroughly washed, sectioned into smaller pieces that have been surface-disinfested with 0.5% sodium hypochlorite for a few seconds, blotted dry, and placed on the agar surface. Plates are incubated at 24–26°C in the light. Mycelia of Colletotrichum spp. grow from the specimen in 1 or 2 days. After 3–4 days, spores form on individual conidiophores on the agar surface. Pure cultures can be established by transferring hyphal tips to V8 juice agar or potato dextrose agar. Some isolates form groups of conidiophores resembling sporodochia after 1 week, and setae may or may not form. Sporulation is common on the host tissue in the isolation plate, but it is very difficult to obtain a pure culture from the specimen piece itself.

To increase the amount of conidia, which is needed for pathogenicity studies, a loopful of spores should be streaked onto a new plate of medium. Large masses of conidia form, especially on potato dextrose agar. Some isolates form perithecia on V8 juice agar about 10–14 days after cultures are initiated.

These isolates generally produce fewer conidia and should be grown on potato dextrose agar for conidial formation. Perithecia also form on the plant specimen or on water agar isolation plates after a few weeks.

Management Efforts to control this disease must begin with sanitation. All dead and severely diseased leaves, petioles, flowers, and fruits should be removed from the plant. For leaves with only a few spots, infected leaflets or leaf sections with disease can be pruned. Infected plant debris should be gathered and removed from the site, since it is a source of inoculum. Potting soil from infected plants should not be reused unless it is steam-sterilized or pasteurized to eliminate pathogens. All pots, trays, and tags must be washed and surface-disinfested with 20% household bleach.

Moisture control is extremely important for disease control.

Without moisture control, the pathogen will splash to new leaves, germinate, penetrate the host, and produce more spores in high humidity, resulting in a continuation of the disease cycle. Overhead irrigation should be limited and rapid drying of the foliage is encouraged. Practices to encourage air movement include increasing plant spacing, trimming dead growth, eliminating all weeds in the pots, and trimming vegetation surrounding the nursery. Dry weather improves crop quality; thus, efforts should be made to remove older leaves during this time.

When wet weather returns, existing spots form conidia, which splash to healthy leaves and spread the disease. Moving the crop to a solid-covered greenhouse helps break the disease cycle.

Many diseases caused by Colletotrichum spp. have been treated with fungicides to reduce disease levels. Mancozeb or similar compounds reduce the number of spores released and the number of spores germinating. Thiophanate methyl fungicides are highly effective for disease prevention but should be rotated with other nonbenzimidazole fungicides to prevent rapid selection of pathogens resistant to fungicides that contain a benzimidazole as the active ingredient. The most effective use of fungicides is in an integrated disease management program that sanitizes the environment by reducing spore levels and controls moisture to prevent high levels of sporulation and new disease initiation.

A suggested control of premature fruit drop of Serenoa repens is burning of the rangelands before flower production.

Fungicides are not considered very cost effective because of the large number of wild Serenoa repens, the asynchronous flowering, and the possibility of other inoculum sources on nonpalm plants.

Alfieri, S. A., Langdon, K. R., Kimbrough, J. W., El-Gholl, N. E., and Wehlburg, C. 1994. Diseases and Disorders of Plants in Florida.

Florida Dep. Agric. and Consumer Service, Div. Plant Industry.

Bull. 14.

Carrington, M. E., Roberts, P. D., Urs, N. V. R. R., McGovern, R. J., Seijo, T. E., and Mullahey, J. J. 2001. Premature fruit drop in saw palmettos caused by Colletotrichum gloeosporioides. Plant Dis. 85:122-125.

Sutton, B. C. 1992. The genus Glomerella and its anamorph Colletotrichum. Pages 1-26 in: Colletotrichum: Biology, Pathology and Control. J. A. Bailey and M. J. Jeger, eds. CAB International, Melksham, U.K. Uchida, J. Y., and Kadooka, C. Y. 1997. Colletotrichum leaf spot of red sealing wax palm. CTAHR Fact Sheet, Plant Dis. PD-10. University of Hawaii at Manoa, College of Tropical Agriculture and Human Resources, Publications and Information Office, Honolulu.

(Prepared by J. Y. Uchida)

Damping-Off While the seedlings of many palms are attacked by various pathogens that kill plants soon after germination or emergence, there have not been many research studies of damping-off on palms.

Symptoms The first symptom of damping-off is the late and irregular emergence of seedlings (Plate 29). Preemergence damping-off refers to seedlings that are killed before emerging from the growing medium. Seedlings that emerge and then rot are referred to as having postemergence damping-off or simply damping-off.

Depending on the palm species, the seed itself may be soft and entirely rotted, along with the seedling. Seedlings exhibit radicle or hypocotyl rots. As apical meristem tips are killed, seedlings are lost (Plate 30). Green leaves may protrude from diseased stems but become brown and dry in a few days.

Causal Organisms Damping-off and seedling blights of palms have been associated with Phytophthora nicotianae Breda de Haan, Phytophthora palmivora (E. J. Butler) E. J. Butler, Sclerotium rolfsii Sacc., and species of the genera Calonectria, Fusarium, Pythium, Rhizoctonia, and Thielaviopsis (synonym = Chalara).

Host Range and Epidemiology Damping-off has been reported for Acoelorrhaphe wrightii, Arenga spp., Carpentaria acuminata, Chamaedorea elegans, Chamaedorea seifrizii, an unidentified Chamaedorea sp., Dypsis lutescens, Elaeis spp., Nannorrhops spp., Physokentia spp., Syagrus spp., Washingtonia robusta, and Wettinia spp. Seedling blights of an Areca sp., a Pinanga sp., Pritchardia thurstonii F. v. Muell. & Drude, Ptychosperma macarthurii, and a Thrinax sp. occurred when inoculated with Phytophthora nicotianae.

Damping-off reflects poor seed quality (e.g., seeds contaminated with a pathogen), contaminated medium that was used to grow seeds, medium that was contaminated after seeds were planted, seeds that were planted excessively deep in the medium, insect damage followed by fungal invasion, or poor environmental conditions for seed germination and seedling growth, such as excessive moisture or improper temperatures.

Each disease cycle begins with the source of the inoculum or contamination. Palm seeds can be attacked by pathogens that subsequently invade seedlings as they grow. If germination trays are placed on the ground rather than on raised benches, 15

there is the real danger that pathogens in the soil will splash or grow into the trays. Deep planting exposes the shoot to a long period of contact with organisms in the media. Some pathogens, especially weak pathogens, invade the buried shoot. Insects that feed on seeds, seedlings, or both provide damaged tissue for opportunistic pathogens in the medium.

Diseased or dead seedlings should be discarded and removed from the nursery to prevent movement of fungal spores or propagules to other crops. Seedlings that survive a damping-off problem are often weakened plants that maintain the pathogen in the nursery in infected roots. Rhizoctonia or Pythium rots on adult plants are examples.

Diagnostic Techniques Germination trays need to be closely monitored for palm seeds that germinate more slowly than normal for the particular palm species being grown. Seeds should be checked immediately to determine depth of planting, presence of insects, condition of the medium (e.g., soggy), temperature range, and watering consistency. Any diseased seedling should be washed well and sections of dead roots, stem, leaf petiole, or leaf blade cut off. Specimen pieces with part of the healthy tissue still present are preferable. Specimen pieces should be dipped in 0.5% sodium hypochlorite for a few seconds, blotted dry, and placed on the surface of water agar. If Pythium or Phytophthora spp. are suspected, some of the specimens should be plated after dipping in 0.25% sodium hypochlorite, since these pathogens are not as tolerant of sodium hypochlorite. Plates should be maintained at 24–26°C in the light.

Isolation plates should be checked daily for mycelial growth of potential pathogens. Pythium and Phytophthora spp. generally grow within 24 hr and produce rapidly expanding coenocytic mycelia. Hyphal strands of these pathogens have no or few cross walls, and the cytoplasm flows within the hyphae.

The agar and hyphae should be carefully cut and transferred to V8 juice agar. Phytophthora spp. may begin to form sporangia within the first 3 days on the water agar isolation plates. Since Phytophthora spp. compete poorly with many saprobes and other pathogens, suspected Phytophthora mycelia should be transferred as soon as possible. Pythium spp. grow rapidly, with sporangia and oospores commonly forming on the isolation plate.

Fusarium spp. produce mycelia with many cross walls and often form canoe-shaped spores and other smaller one- or twocelled spores on water agar after a few days. Calonectria spp. produce septate mycelia after 2–3 days. Hyphal sections of these fungi should be transferred to V8 juice agar. Within 1 week, the anamorphs of Calonectria spp., Cylindrocladium spp., produce conidia. Depending on the Calonectria species, tiny red perithecia may be present on diseased plant tissue and on the V8 juice agar. However, certain nonpathogenic Fusarium spp. also form red perithecia. Thus, the type of ascospore formed must be determined.

Rhizoctonia spp. grow rapidly and are present on the first or second day. This group of fungi produces wide mycelia with right-angle branching and becomes some shade of brown on V8 juice agar and potato dextrose agar. Hyphae of Sclerotium rolfsii are characterized by the presence of clamp connections.

Sclerotium rolfsii cultures produce tan to brown, spherical sclerotia that look like mustard seeds after 7–14 days. These tan to dark brown sclerotia also form on diseased tissue and can be a clue as to the presence of this fungus. Some Rhizoctonia isolates also form sclerotia in culture. Sclerotium and Rhizoctonia spp. do not form asexual spores.

Thielaviopsis spp. are generally slow growing on water agar, producing distinctive conidiophores with chains of endoconidia. A sterile glass needle should be used to touch the spore chain and transfer it to V8 juice agar or potato dextrose agar to obtain a pure culture. A second type of spore that forms on these media is single-celled, dark, and ellipsoidal to obpyri16

form with thickened walls. All isolates should be maintained at 24–26°C in the light.

Although unreported, Colletotrichum spp., confirmed pathogens of Areca catechu, Cyrtostachys renda, Phoenix roebelenii, and Serenoa repens, may contaminate seedling trays and infect young plants. These organisms should be considered potential damping-off pathogens for these palm species. See Colletotrichum Leaf and Fruit Spot for more information on these pathogens.

Management There have not been many research studies of damping-off on palms, but studies of damping-off on other plants are applicable. In addition, knowledge regarding the germination of the various palm species is vital. Estimates have been made that more than 25% of all palm species require more than 100 days to germinate. Furthermore, the percent germination for many species is often less than 20%. Moisture, temperature, planting depth, and use of fresh, disease-free seeds are the most critical factors for germination. Germination procedures vary for different palms but most require high temperatures for germination. The time required for germination varies from less than 2 weeks for Washingtonia spp. to 1 year or longer for other palm species. Patience is encouraged, as well as caution so that ungerminated seeds that may still be viable are not thrown out.

The use of clean seeds is important to prevent seedling disease and subsequent root rots of older plants. It is preferable that seeds be picked from the trees when they are fully ripe.

Seeds should not be collected from the ground unless absolutely necessary. The soft outer flesh should be removed and the seeds rinsed thoroughly in clean water. It is best to plant palm seeds shortly after cleaning, preferably within 24 hr.

While not necessary for seeds picked from trees, seeds collected from the ground can be disinfested by dipping them in a 20% household bleach solution, containing a small amount of detergent, for approximately 2–3 min. They should then be rinsed in running water.

For small-sized seeds, many seeds can be spread on the surface of a light plant growth medium, such as moderate- or course-grade vermiculite, covered very lightly with more medium, watered to keep moist but not soaking wet, and grown at 30–35°C. If lower temperatures are used, germination time will be longer. Some growers place seeds in trays and then in a moist environment, such as a partially closed plastic bag or small plastic tent on the germination bench. For seeds collected from the ground, fewer should be planted per pot and they should be monitored closely. In this way, if disease develops, fewer seeds will be lost.

All media, trays, benches, etc. must be clean. Solid-covered structures are best for germinating and growing seedlings.

Without a solid-covered structure, moisture control is impossible in wet environments. Trays should not be placed on the ground if possible. A relatively inexpensive method can be devised to keep seedling trays off the ground. The ground under the trays can be covered with a thick layer of gravel and sand.

The entire area (ground, benches, etc.) should be routinely sprayed with insecticides to keep insect populations small.

Ants and other insects that crawl into trays can carry pathogen spores into the medium. All slugs or snails must be eliminated for similar reasons. There should be no moss or algal growth in the seed germination area, since heavy algal growth increases the population of shoreflies and fungus gnats. These insects are also attracted to the decomposing pulp on fresh palm seeds and can move pathogens into seed trays. Pulp should be removed before planting or insecticides should be used to reduce insect populations.

Planted seeds should be monitored frequently. Any seedling that begins to wilt or die should be removed immediately. Seed germination media must be kept uniformly moist, but never soaking wet. Damping-off is usually an indication that the me-

dium has been kept too wet. The causal organism should be determined for dead seedlings and appropriate fungicides should be applied. Labels should be checked for the appropriate chemicals to use and all directions should be followed.

Selected References Broschat, T. K., and Meerow, A. W. 2000. Ornamental Palm Horticulture. University Press of Florida, Gainesville, FL. Singleton, L. L., Mihail, J. D., and Rush, C. M., eds. 1992. Methods for Research on Soilborne Phytopathogenic Fungi. American Phytopathological Society, St. Paul, MN.

(Prepared by J. Y. Uchida)

Diamond Scale Diamond scale is a disease caused by a single pathogen on a single host in the southwestern United States. See Rachis Blight and Tar Spot for diseases caused by similar pathogens.

Symptoms The pathogen is present on leaf blades and petioles as black, diamond-shaped fruiting structures (stroma) throughout the leaf canopy (Plates 31 and 32). Numbers of stroma on plant tissue can range from a few to thousands. Eventually, leaves turn chlorotic (Plate 33), senesce prematurely, and die (Plate 34).

Causal Organism Phaeochoropsis neowashingtoniae (Shear) K. D. Hyde and P. F. Cannon (synonyms = Sphaerodothis neowashingtoniae Shear and Phaeochora neowashingtoniae (Shear) Theiss. & Syd.) is the causal organism of diamond scale. Stromata form on both sides of the leaf tissue. They range in size (1.5–3.5 × 3–8 mm) and are strongly domed (Plates 31 and 32). While initially light brown, melanization of fungal cells results in uniform, shiny black stromata. Cross sections reveal individual chambers separated by dark-colored columns of cells (Plate 35). Asci are 80–117 × 92–132 µm, initially with very long (up to 80 µm) stalks. Ascospores are 20–33 × 55–75 µm, hyaline, and enveloped in mucilage when young, becoming dark brown as they mature. They are oblong-ellipsoid, aseptate, and flattened on one side, sometimes forming a shallow depression.

Host Range and Epidemiology This pathogen has only been reported on Washingtonia filifera and only in California and Arizona (United States). Ascospores are released by weathering of the stroma and are produced in such numbers that the leaf may appear black. Germination of spores around the stroma rings it with new lesions and eventually new stroma.

Diagnostic Techniques The black, diamond-shaped stroma are diagnostic, with the long axis parallel to the leaf veins.

Management Removal of infected foliage from the site (nursery or landscape) and use of protective fungicides may be helpful, although appropriate timing of the fungicide treatment is unknown.

Selected References Hyde, K. D., and Cannon, P. F. 1999. Fungi causing tar spots on palms. Mycol. Pap. 175. CABI Publishing, Wallingford, U.K. Ohr, H. D. 1991. Diamond scale. Pages 10-11 in: Diseases and Disorders of Ornamental Palms, A. R. Chase and T. K. Broschat, eds.

American Phytopathological Society, St. Paul, MN.

(Prepared by M. L. Elliott)

Fusarium Wilt Fusarium wilt of palms is a vascular wilt disease caused by a complex of formae specialis of Fusarium oxysporum Schlechtend.:Fr. This complex currently consists of three distinct diseases: Canary Island date palm wilt, date palm wilt or Bayoud disease, and oil palm wilt. The latter is also known as Fusariose or Tracheomycose. Canary Island date palm wilt, primarily infecting Phoenix canariensis, is the most recent Fusarium wilt disease of palms to be described and is reported from Australia, the Canary Islands, France, Italy, Japan, and the United States (California, Florida, and Nevada). Date palm wilt of Phoenix dactylifera is reported from Algeria and Morocco. Oil palm wilt of Elaeis guineensis occurs in Benin, Brazil, Cameroon, Colombia, Democratic Republic of the Congo, Ecuador, Ghana, Ivory Coast, Nigeria, Republic of the Congo, and Surinam. Since all three primary palm hosts have aesthetic value as ornamentals, they are profiled separately.

Canary Island Date Palm Wilt Symptoms Canary Island date palm wilt of Phoenix canariensis exhibits decline symptoms similar to those caused by other root and stem diseases. Affected palms decline from the lower canopy upward to the meristem (Plate 36), resulting in a cessation of new growth prior to palm death (Plate 37). During the period of infection, palm vigor is reduced as new fronds fail to emerge.

The fronds on infected trees exhibit a subtle wilt as the angle between the frond and the trunk increases by 10–30 degrees.

Frond strength lessens, and affected fronds can be snapped along the rachis because of increased brittleness.

Initial symptoms generally develop on the lower fronds and progress upward on the trunk. Pinnae (palm leaflets) and spines begin to desiccate and die from the petiole base outward to the frond tip in a one-sided manner (Plate 38). This decline continues back to the petiole base on the opposite side. Pinnae whiten as they desiccate and die. Simultaneous to pinnae death on a frond, a dark brown stripe develops on the petiole surface, beginning at the petiole base and progressing outward to the frond tip (Plates 39 and 40). Some pinnae and spines may exhibit a similar discoloration on their abaxial surface. Discrete areas of vascular discoloration are evident in both cross and longitudinal sections of the petiole and the trunk (Plate 41). This discoloration may range from salmonpink to reddish brown. Much of the trunk discoloration is limited to the outer perimeter of the bole. Mature trees die in a few months.

Symptoms may vary in affected palms. A palm may exhibit the first symptom on a frond in the midcanopy (Plate 42).

Symptom progression continues in the midcanopy to produce a one-sided death. Lower-frond symptoms eventually develop in this symptom scenario. This pattern of disease development may span 2 years. Additionally, frond death may initiate at the frond tip, producing pinnae desiccation on one or both sides of the petiole and progressing back to the rachis base (Plate 43). In some trees, frond dieback may follow both tip and one-sided dieback patterns. In some areas, other diseases or pest problems may alter the sequence of symptoms. Incidence of pink rot (Gliocladium vermoeseni (Biourge) Thom) in the southwestern United States may hasten tree death and confuse petiole symptoms as a result of the external hyphae of G. vermoeseni. Similarly, the incidence of palm weevils may hasten frond wilt and tree death without the consistent development of symptoms associated with Canary Island date palm wilt.

17

Causal Organism The causal organism is F. oxysporum Schlechtend.:Fr. f. sp. canariensis Mercier & Louvet, a moniliaceous member of the subdivision Deuteromycotina. Septate hyphae produce aseptate, short conidiophores that give rise to both microconidia and macroconidia. Microconidia are hyaline, primarily one-celled (few with one septum), oval to elliptical, 3–5 × 3–15 µm, and clustered at the tip of the conidiophore. Macroconidia are 3–5 × 10–35 µm, two- to five-celled, falcate-fusoid, and produced less frequently from conidiophores but do form abundantly off sporodochia in culture. Chlamydospores are both terminal and intercalary, are 2–11 µm in diameter, and may develop from macroconidia. Violet-purple sclerotia can form in culture.

Host Range and Epidemiology This pathogen has the broadest host range of the three F. oxysporum formae specialis affecting the palm family. F. oxysporum f. sp. canariensis has been reported on both P. canariensis and P. dactylifera. Additional hosts in the genus Phoenix include P. reclinata and also P. sylvestris, as diagnosed by polymerase chain reaction (PCR) assay (G. W. Simone, unpublished data). Washingtonia filifera is also reported to be susceptible to this pathogen (H. D. Ohr, unpublished data). Symptom expression in P. dactylifera is variable.

Trees may not be killed, although the offshoots do decline to death. P. dactylifera seedlings inoculated with F. oxysporum f. sp. canariensis exhibit the same symptoms as do seedlings of P. canariensis.

Little information is available on the epidemiology of Canary Island date palm wilt. Palm loss occurs in landscape sites, street borders, and field nurseries in many diverse areas of the world. Primary infection is believed to occur through invasion of the pneumathodes of aerenchymatous roots. These roots appear both on the lower trunk of the palm and in the upper soil profile. The pathogen progresses through aerenchymatous roots to the trunk cortex and then into discrete vascular traces and upward into the canopy of the palm.

Introduction of F. oxysporum f. sp. canariensis into a region has two likely paths. P. canariensis is a highly prized landscape tree. The horticultural industry has long imported both plants and seeds to meet the growing needs of the landscape industry.

It is conceivable that importation of infected plants in the past accounts for some of the distribution of this wilt pathogen.

With the development of the specific primer set for F. oxysporum f. sp. canariensis, use of a PCR assay has revealed this pathogen on seed coats. Additionally, F. oxysporum f. sp. canariensis was detected by PCR assay from seedlings grown from wilt-infected plants. These palms had husks removed before planting and were grown in isolation for 2 years prior to destructive sampling (G. W. Simone, unpublished data). PCR assays have also provided evidence that multiple introductions have occurred in Florida, since F. oxysporum f. sp. canariensis isolates from Florida have DNA fingerprints more closely related to those from Sicily, France, and Japan than to others in Florida.

Short- and long-distance spread have other possible causes.

One aspect of the nursery industry in the United States focuses upon the rescue of specimen trees (including palms) from old landscapes or areas faced with significant redevelopment. Often these nurseries buy, excavate, and field plant these trees for periods of 6 months to several years while awaiting sale. This is a situation that can both introduce the pathogen from an infested landscape site into a holding nursery and then serve to spread the pathogen through the newly infested nursery soil to a series of root-wounded new trees destined for still other landscape sites.

The landscape maintenance industry may have a critical role in local spread of this wilt disease as well. In new plantings, 50–75% of existing canopy may be pruned to enhance tree establishment. Low-hanging, viable fronds of Phoenix spp. are 18

often removed on a seasonal or annual basis because of liability concerns. Additionally, palms are often sheared severely in a “hurricane cut” on a yearly basis, in which more than 50% of the tree canopy is removed. These frequent periods of green frond removal provide opportunity for the spread of F. oxysporum f. sp. canariensis on chain saws or other pruning tools.

Transmission of this pathogen by chain saws during pruning of young P. canariensis has been demonstrated (T. V. Feather, unpublished data).

One final disease-spread scenario to consider is the role of municipal yard waste recycling programs in areas where the disease exists. The majority of these programs collect, chip, pile, and redistribute this infested material to countless other landscapes to be used as mulch around ornamental plantings, which include palms. Since this material is not composted, it is unlikely that dormant chlamydospores are eradicated in the palm debris.

Diagnostic Techniques The diagnostic process begins with interpretation of symptoms on affected palms. It is important that all the key symptoms of Canary Island date palm wilt be present, not just one symptom. Lower-frond death may reflect either Fusarium wilt or Ganoderma butt rot. Similarly, tip dieback or onesided death of pinnae on a frond could be caused by diamond scale, rachis blight, or Fusarium wilt. When all the symptoms of wilt exist, any portion of the plant can be sampled for potential recovery of F. oxysporum f. sp. canariensis. The most reliable areas to sample are areas of vascular discoloration in the fronds or trunk, while root tissue has a lower frequency of recovery.

Vascular tissue pieces should be surface-disinfested in a solution of 0.5% sodium hypochlorite or similar disinfestant for several minutes. Standard growth media, such as potato dextrose agar, support isolation of F. oxysporum f. sp. canariensis.

This pathogen covers a 100-mm petri plate in 5–6 days, producing an appressed colony with an orange-pink to salmonpink color if incubated in the light. In the dark, the colony has a fluffy growth habit, is white, and develops a purple stain in the medium. Microconidia develop first, but both conidial stages should be evident in 7–10 days. Sporodochia develop in culture and produce abundant macroconidia. Chlamydospores and sclerotia require 2–4 weeks in the light to develop. This pathogen grows optimally between 24 and 30°C, with little or no growth at 5 or 40°C. For the most rapid and accurate identification of this pathogen, colonies should be transferred to carnation leaf agar.

Determining mycological characteristics as described above for cultures obtained from diseased plant tissue only confirms the presence of F. oxysporum and not the presence of any specific formae specialis. There are many palm tissues in which saprophytic strains of F. oxysporum exist. Validation of F. oxysporum f. sp. canariensis requires use of the oligonucleotide primer set in a PCR assay. This technology identified 98% of the 71 F. oxysporum f. sp. canariensis isolates examined, including isolates from around the world and isolates of the oil palm wilt and date palm wilt pathogens. The only false positives in the research detected by this primer set were isolates of F. oxysporum f. sp. cubense (E. F. Sm.) W. C. Snyder & H. N. Hans. Recognition of the banana wilt pathogen indicates a common ancestor but should not pose a source of confusion when dealing with palm samples.

Management An integrated program for Canary Island date palm wilt (Fusarium wilt) management in the landscape and nursery is based more on common sense than on data. In view of the host range of F. oxysporum f. sp. canariensis and its suspected epidemiology, management suggestions are provided for both nursery and landscape settings.

For nursery production sites, the seedborne aspect of the pathogen places new emphasis on the procurement of pathogen-free seed sources. Seed lots should be purchased from areas of the world without Canary Island date palm wilt when possible. The technology exists to certify seed-bearing trees in areas where the disease is endemic. For nurseries specializing in the procurement of specimen Phoenix palms, field collection of these trees poses a risk of moving the pathogen into a holding nursery and subsequently passing infected stock into the trade. Some landscape architects in Florida have required successful PCR assay certification of specimen Phoenix palms as part of the bid process. Field production nurseries of susceptible Phoenix spp. should be routinely inspected for symptoms of wilt prior to any pruning operations. Suspect palms should be assayed by PCR for final determination. Symptomatic trees should be pruned separately, immediately followed by the use of an appropriate disinfestant for pruning tools.

In the landscape setting, only adapted palms from reputable sources should be purchased. Phoenix spp. should not be placed into sites with excessive irrigation, poor drainage, or a high water table. These conditions stress the palms and encourage more aerenchymatous root development and, hence, higher risk for invasion by F. oxysporum f. sp. canariensis. Other plant material should not be placed over the immediate palm root system, since installation damages the aerenchymatous roots. In areas where this disease is known, municipal mulch sources should not be used around susceptible palms if possible. When the disease is diagnosed, total removal of the palm should follow. The diseased palm should be sent to a landfill or incinerator whenever possible and not to a municipal plant material recycling center. The dead palm should be replaced with a nonpalm species if possible. Reinstallation of Phoenix or Washingtonia spp. should be avoided. In areas with this disease, landscape maintenance companies should become familiar with wilt symptoms. Palms should not be pruned with chain saws. Alternative pruning tools should be used, one per tree.

They should be brushed free of wood dust and disinfested in a 1:3 dilution of a pine oil product in water for 7–10 min. This disinfestant treatment is superior to household bleach since it is not corrosive and is as efficacious as denatured ethanol, without the risk of flammability (G. W. Simone, unpublished data).

Trunk-injected systemic fungicides have had no benefit in disease management (T. V. Feather, unpublished data).

Selected References Arai, K., and Yamamoto, A. 1974. New Fusarium disease of Canary Island date palm in Japan. Bull. Fac. Agric. Kagoshima Univ. 27: 31-37.

Burgess, L. W., Summerell, B. A., Bullock, S., Gott, K. P., and Backhouse, D. 1994. Laboratory Manual for Fusarium Research, 3rd ed. University of Sydney, Sydney, Australia.

Feather, T. V. 1982. Occurrence, etiology and control of wilt and dieback of Phoenix canariensis in California. Ph.D. diss. University of California, Riverside, CA. Feather, T. V., Ohr, H. D., and Munnecke, D. E. 1979. Occurrence of Fusarium oxysporum and Gliocladium vermoeseni on Phoenix canariensis in California, and their effects on seedlings of Phoenix species. Date Grow. Inst. Rep. 54:17-18.

Feather, T. V., Ohr, H. D., Munnecke, D. E., and Carpenter, J. B. 1989. The occurrence of Fusarium oxysporum on Phoenix canariensis, a potential danger to date production in California. Plant Dis. 73:78-80.

Mercier, S., and Louvet, J. 1973. Recherches sur les Fusarioses. X.— Une Fusariose vasculaire (Fusarium oxysporum) du palmier des canaries (Phoenix canariensis). Ann. Phytopathol. 5:203-211.

Pfalzgraf, K. 2002. Loss of a legacy—Fusarium oxysporum and ornamental Phoenix canariensis. Palms 46:161-166.

Plyler, T. R., Simone, G. W., Fernandez, D., and Kistler, H. C. 1999.

Rapid detection of the Fusarium oxysporum lineage containing the Canary Island date palm wilt pathogen. Phytopathology 89:407413.

Priest, M. J., and Letham, D. B. 1996. Vascular wilt of Phoenix canariensis in New South Wales caused by Fusarium oxysporum.

Australas. Plant Pathol. 25:110-113.

Simone, G. W., and Cashion, G. 1996. Fusarium wilt of Canary Island date palms in Florida. Plant Pathol. Fact Sheet PP-44. Florida Extension Service, Institute of Food and Agricultural Sciences, University of Florida, Gainesville, FL.

Date Palm Wilt (Bayoud Disease) Symptoms The Draa Valley in Morocco is believed to be the first observed site of date palm wilt of Phoenix dactylifera prior to the 1870s. Initial symptoms appear in one or more mature leaves.

Spines or pinnae turn white on one side of the leaf base, and the discoloration progresses outward to the apex of the leaf before progressing down the other side to the leaf base. As foliar symptoms develop, the petiole base develops a brown, slightly sunken zone of discoloration. A cross section of the petiole reveals a reddish brown discoloration. Symptom development may span days to weeks. Additional leaves in the affected whorl or whorls develop symptoms until one or more whorls of leaves are dead. During disease development, new leaves cease to emerge. The canopy symptoms continue until the bud dies.

Tree death may span a period of 6–24 months or longer. If palm trunks are split longitudinally, they exhibit reddish brown strands of vascular tissue that can extend from roots to the meristem. Foliar symptoms can vary. Disease can cause leaf dieback from apex to base. After palm death, it is not unusual for offshoots to remain unaffected for long periods of time (up to 10 years).

Causal Organism The date palm wilt pathogen is Fusarium oxysporum Schlechtend.:Fr. f. sp. albedinis (Killian & Maire) Gordon.

This fungus produces microconidia from simple phialides that arise laterally from hyphae. Microconidia are generally onecelled, ellipsoid to variable, and 3–5 × 3–15 µm. Macroconidia are borne on phialides, have one to three septa, and are 3–5 × 20–35 µm. Chlamydospores are produced singly or in small groups in an intercalary or terminal manner and are 5–12 µm in diameter. Sclerotia are dark blue to black, but they rarely occur.

The teleomorph is believed to be a Gibberella sp., but the teleomorph has not been associated with this disease. This pathogen can grow at temperatures as low as 18°C and as high as 32°C, with an optimum of 21–27.5°C. Based upon evaluation of more than 40 isolates from Morocco and Algeria, all isolates appear to share a single clonal lineage. All isolates fell within one vegetative compatibility group and exhibited no polymorphisms by either restriction fragment length polymorphism (RFLP) or random amplified polymorphic DNA (RAPD) analysis.

Host Range and Epidemiology As with many formae specialis, F. oxysporum f. sp. albedinis has a very narrow host range, invading only P. dactylifera. Although several reports exist in the literature suggesting that this pathogen also invades P. canariensis, these reports coincide in time with the first reports of Canary Island date palm wilt. In view of the broader host range for F. oxysporum f. sp. canariensis Mercier & Louvet, these reports may actually reflect Canary Island date palm wilt. Field investigations on the survival of F. oxysporum f. sp. albedinis in common field flora revealed that henna (Lawsonia inermis L.) and alfalfa (Medicago sativa L.) were symptomless hosts of this pathogen.

Primary infection is believed to proceed from chlamydospores lying dormant in the soil or in plant debris to depths of 5–30 cm in the soil. Root pneumathodes are invaded by F. oxy19

sporum f. sp. albedinis. The fungus initially grows inside the aerenchyma and then grows inter- and intracellularly through the cortex until it penetrates the vascular tissues. The pathogen ascends to the meristem by microconidial movement through vessels and direct germ tube penetration of transverse vessel walls. During the ascent, the pathogen colonizes the adjacent parenchyma and sclerenchyma cells, producing vascular discoloration and abundant chlamydospores. Evidence for the infection of the female flower exists, but this is not the primary path of infection. This pathogen is not borne in seeds or pollen.

Epidemic outbreaks of date palm wilt have been correlated to weather patterns, irrigation frequency and quantity, and P. dactylifera variety. In general, disease spread slows during dry seasons with minimal irrigation, minimal pruning of the canopy, or both. Disease spread increases after wet seasons, increased irrigation schedules, or severe pruning. Fields with a predominance of susceptible cultivars sustain higher rates of disease spread than do fields planted to resistant cultivars. Directional movement of soil water does not appear to correlate to pathogen spread. Tree-to-tree spread appears dependent upon root contact. The rate of disease spread is unaffected by soil or water salinity levels up to 5 g/liter. Long-distance spread of the disease appears due to movement of infected offshoots or movement of infected palm fragments or products.

Diagnostic Techniques The symptoms of date palm wilt of P. dactylifera have been quite distinctive for purposes of diagnosis over the years. The recent report of a second wilt/decline disease of P. dactylifera from Saudi Arabia caused by F. proliferatum (T. Matsushima) Nirenberg now lowers the accuracy of field diagnosis based upon symptomatology. Mixed populations of F. proliferatum and F. oxysporum from tissues of palms affected with this new disease mandate careful diagnosis and use of new technologies.

The pathogen can be isolated from almost any symptomatic plant part. More F. oxysporum f. sp. albedinis colonies are recovered from symptomatic canopy or trunk tissue than from root tissue. Palm tissue exhibiting vascular discoloration should be sampled. Excised pieces should be treated with a general disinfestant solution (e.g., 0.5% sodium hypochlorite) for several minutes. General isolation media such as potato dextrose agar are adequate for pathogen isolation, but carnation leaf agar should be used as the medium for final identification. Recovery of Fusarium spp. that possess the mycological characteristics listed earlier offers reasonable validation for the presence of F. oxysporum f. sp. albedinis.

Absolute confirmation of F. oxysporum f. sp. albedinis can be obtained with a proof-of-pathogenicity trial using a susceptible P. dactylifera cultivar seedling (e.g., Medjool). Suspect isolates should be inoculated at a rate of 2 ml of 106 spores per ml onto seedlings with one or two leaves. The assay should result in seedling mortality in less than 120 days. In situations requiring faster diagnosis to the formae specialis level, two newer methods exist. A distinct polymorphism has been reported in culture from F. oxysporum f. sp. albedinis isolates, since the wild type degenerates from prolonged maintenance on synthetic media by mass transfer. Also, Fernandez et al. have designed a set of primer pairs for polymerase chain reaction (PCR) assays that recognizes 95% of the F. oxysporum f. sp. albedinis isolates tested. Use of these primer pairs in PCR assays allows for presymptomatic detection of wilt-infected plants in the field. Use of this assay will allow faster implementation of eradication programs, accelerated screening of progeny from breeding programs, and use of critical regulatory functions such as palm indexing for international movement of germ plasm.

Management Date palm wilt (Fusarium wilt) is such an economically devastating disease that all strategies for disease management have 20

been and continue to be examined for utility. During the first 80 years of documented disease spread in Morocco alone, more than 10 million date palm trees have been destroyed.

In nations where date palm wilt exists, regional exclusionary measures have not been very successful in limiting in-country spread of this disease. Strict quarantine regulations dealing with the movement of infected palm products or offshoots between nations have been effective in preventing the introduction of F. oxysporum f. sp. albedinis to adjacent countries.

Physical exclusion measures can also prevent short-distance spread of F. oxysporum f. sp. albedinis. Early field research in Morocco demonstrated the effectiveness of 2-m-deep trench barriers in preventing pathogen spread from soil/root contact between infested and healthy P. dactylifera production plots.

Use of this physical method of exclusion may extend the productive life of some palm plantings.

Much effort has gone into the eradication of infected trees in primary infection foci by means of control. Uprooting and burning infected trees has proven labor and cost intensive. Following these actions with soil fumigation with chloropicrin and a suitable fallow period have often been cost prohibitive. Limitations to effective eradication action have included the absence of both a diagnostic method for early disease detection and a uniform and fair program of compensation for the citizen or grower involved in the eradication program.

Cultural investigations demonstrated that disease spread could be slowed by a reduction in fertility and irrigation quantity. Unfortunately, as disease spread decreased so did yield.

Managing irrigation timing has allowed the use of susceptible cultivars in an active disease focus, if the irrigation was suspended between May and October. Although these cultivars survived, yield was too low to be economical. Suspension of intercropping with high-irrigation-demand species such as vegetables and forages also served to slow disease spread and development, but not without an impact on productivity. Introducing resistant palm cultivars improves yield potential under conditions of controlled irrigation timing or with the use of intercropping with supplemental irrigation.

Fungicide protection and therapy actions have not been efficacious or cost effective for the field production of P. dactylifera. Development of antagonistic biological species has seemed promising in vitro but has not reached field application status. Some of these microbes may prove useful as inoculants in seed production nurseries or in the replanting of fallowed fields.

Resistance breeding is believed to be the best management tool for long-range control of date palm wilt. Resistance has been sought from established varieties, from natural populations of seed-grown palms, and through hybridization programs.

The cultivars Black Bou-Sthammi, Bou-Feggous ou-Moussa, Bou Ijjou, Bou-Sthammi, Bou Zeggar, Iklane, Outoukdim, Sairs-Layalet, Sthammi, Tadment, Takerboucht, and White Bou-Sthammi have been documented with resistance to F. oxysporum f. sp. albedinis and have served as the basis for the early breeding efforts in North Africa. This effort has been doubly challenging since few of these selections offered acceptable fruit quality.

Selected References Abdalla, M. Y., Al-Rokibah, A., Moretti, A., and Mulè, G. 2000.

Pathogenicity of toxigenic Fusarium proliferatum from date palm in Saudi Arabia. Plant Dis. 84:321-324.

Belarbi-Halli, R., and Mangenot, F. 1986. Bayoud disease of date palm: Ultrastructure of root infection through pneumatodes. Can. J. Bot. 64:1703-1711.

Bulit, J., Louvet, J., Bouhot, D., and Toutain, G. 1967. Recherches sur les Fusarioses. I. Travaux sur le Bayoud, Fusariose du palmier dattier en Afrique du Nord. Ann. Epiphyt. 18:213-239.

Carpenter, J. B., and Elmer, H. S. 1978. Pests and Diseases of the Date Palm. U.S. Dep. Agric. Handb. 527.

Corte, A. 1973. La tracheomicosi da Fusarium oxysporum f. sp. albedinis della Phoenix canariensis. Not. Mal. Piante (Genoa) 88/89:107-117.

Djerbi, M. 1982. Bayoud disease in North Africa: History, distribution, diagnosis, and control. Date Palm J. 1(2):153-197.

Fernandez, D., Quinten, M., Tantaoui, A., Geiger, J.-P., Daboussi, M.J., and Langin, T. 1998. Fot 1 insertions in the Fusarium oxysporum f. sp. albedinis genome provide diagnostic PCR targets for detection of the date palm pathogen. Appl. Environ. Microbiol.

64:633-636.

Killian, C., and Maire, R. 1930. Le Bayoud, maladie du dattier. Bull.

Soc. Hist. Nat. Afr. Nord 21:89-101.

Malencon, G. 1936. Les palmeraires du Draa et le Bayoud. Rev. Mycol. 1:191-206.

Pereau-Leroy, P. 1954. Varietes de dattier resistantes a la fusariose.

Fruits 9:450-451.

Pereau-Leroy, P. 1958. Le palmier dattier au Maroc. Min. Agric. Maroc, Serv. Rech. Agron. et Inst. Francais Rech. Fruit, Outre-mer, France. Rabat, Morocco.

Sabaou, N., Bounaga, N., and Bounaga, D. 1983. Actions antibiotique, mycolytique et parasitaire de deux actinomycetes envers Fusarium oxysporum f. sp. albedinis et autres formae speciales.

Can. J. Microbiol. 29:194-199.

Sedra, M. H., and Djerbi, M. 1985. Mise au point d’une methode rapide et precise d’identification in vitro du Fusarium oxysporum f. sp. albedinis, agent causal du Bayoud. Ann. Inst. Natl. Rech.

Agron. Tunis. 58:1-12.

Tantaoui, A., Ouinten, M., Geiger, J.-P., and Fernandez, D. 1996.

Characterization of a single clonal lineage of Fusarium oxysporum f. sp. albedinis causing Bayoud disease of date palm in Morocco.

Phytopathology 86:787-792.

Oil Palm Wilt Symptoms The symptomatology of Fusarium oxysporum Schlechtend.:Fr. f. sp. elaeidis Toovey in Elaeis guineensis is quite complex depending upon host age and whether the disease progression is acute or chronic. The pathogen can infect and kill seedlings as well as mature trees.

Seedling infection is not common but can occur in nurseries.

Initial symptoms are growth retardation, few or no pinnate leaves, narrowed and shortened leaf size, and chlorosis of older leaves. Vascular discoloration occurs in the bole as well as in roots. Tip necrosis may occur in some infected seedlings.

Field palms can show two patterns of disease expression— chronic and acute. In palms of pre-fruit-bearing age, a lemonyellow chlorosis develops in the midcanopy. Pinnae discolor from the tip back to the petiole base, while chlorosis spreads to adjacent fronds before moving to older canopy. Vascular discoloration can be observed in the petiole. Early chlorosis is followed by rapid death of old canopy (held erect) and finally spear collapse. Roots appear predominantly healthy. Disease development on young trees proceeds rapidly to death in 2–3 months. Infrequently, there is a remission of symptoms, which may indicate a degree of resistance.

Mature palms also express a variety of symptoms. Lower leaves wilt and desiccate, usually without chlorosis. Affected fronds break near the base and skirt the trunk. Growth slows as new fronds express chlorosis and stunting. There is premature fruit fall and withering and death of young bunches. Occasionally, trees exhibit chlorosis, wilt, and desiccation of two to three younger, midcanopy leaves. A one-sided decline develops before lower-canopy symptoms develop. Foliar symptom progression ascends the tree rapidly until the bud dies.

Some roots are brownish black with brownish red streaks in the vascular traces close to the trunk. A grayish black vascular discoloration is evident in the periphery of the trunk and may extend 1 m above the soil line. Fungal hyphae are evident in the vessels. This acute disease progression may

take only 3–4 months before tree death. If disease progression is chronic in a mature tree, affected leaves dry and fall.

New growth slows, and the stem shrivels or narrows to a pencillike point. Vascular discoloration is evident in the trunk but rarely in the petioles. The root system declines, with most roots exhibiting a brown vascular core. Affected trees may survive several years.

Causal Organism The oil palm wilt pathogen is F. oxysporum f. sp. elaeidis and was first described in 1946, with pathogenicity proven in 1951. The fungus forms microconidia, macroconidia, and chlamydospores in vivo. As with the other Fusarium wilt pathogens, no teleomorph has been observed. Microconidia are hyaline, oval to ellipsoid, 2.5–3.4 × 4.2–6.8 µm, and borne on short, aseptate conidiophores. Macroconidia are straight to slightly curved with pointed ends, have three to five septa, and are 3–4.5 × 13.5–33 µm. Chlamydospores are mostly smooth walled, borne singly or in pairs, intercalary or terminal, and 4.5–9 µm in diameter. Dimensions for Brazilian isolates of F. oxysporum f. sp. elaeidis are somewhat larger than the original report. In some wilt situations in Africa, a second pathogen (F. oxysporum var. redolens Gordon) has been identified as causing a wilt disease.

Host Range and Epidemiology The host range of F. oxysporum f. sp. elaeidis in the palm family is restricted to the genus Elaeis, consisting of E. guineensis and E. oleifera (Kunth) Cortes. Symptomless weed hosts for F. oxysporum f. sp. elaeidis include Amaranthus spinosus L., Eupatorium odoratum L., and Cyperus (Mariscus) alternifolius L. The pathogen was also isolated from Imperata cylindrica (L.) P. Beauv., but back-inoculation to Elaeis spp. was not successful.

Infection of Elaeis spp. occurs in both wounded and unwounded adventitious and underground roots through pneumathodes. The fungus progresses through the cortex, reaching the xylem, and then proceeds into the lower trunk and peripheral leaves through the invaded vascular tissues. Chlamydospores form in infected tissue directly and from macroconidia and serve as survival structures. The pathogen is externally borne on the seed coat, in the kernel, and in pollen.

Diagnostic Techniques Discolored vascular tissue should be sampled from symptomatic trees. Isolation of the pathogen has greater success from trunk or rachis tissue than from root tissue. A standard disinfestant solution (e.g., 0.5% sodium hypochlorite) should be used for 2–3 min, and the tissue should then be placed onto potato dextrose agar. F. oxysporum f. sp. elaeidis produces aerial mycelium on potato dextrose agar at 25°C and a blue stain in the medium in 2 weeks. Colonies should be transferred to carnation leaf agar for accurate identification. The most abundant spore stage is the microconidium, with macroconidia forming as the culture ages. Chlamydospores form in 3–7 days, salmonrose sporodochia form within 1 week, and bluish green to purple sclerotia begin to appear at about 2 weeks. Conformance to the above mycological characteristics combined with plant symptoms provide high confidence of diagnosis. Final confirmation requires a test of pathogenicity. Seedling bioassays utilize oil palm seedlings (two leaves) and require 4–5 months before symptoms are expressed.

A comparison of 76 strains of F. oxysporum f. sp. elaeidis and 21 strains of soilborne F. oxysporum has been performed with a random DNA probe from F. oxysporum f. sp. elaeidis.

The saprobic strains exhibited a simple restriction pattern and were distinct from the F. oxysporum f. sp. elaeidis strains.

A rapid diagnostic tool may become available to assist in the detection of new disease sites and the evaluation of germ plasm.

21

Management The importance of E. guineensis as a crop species cannot be overstated, since it is the largest single source of the world’s supply of vegetable oil. As the incidence of F. oxysporum f. sp. elaeidis continues to spread (23% in some Nigerian estates), any and every management tool is being examined. International quarantines on movement of Elaeis plants, seeds, and pollen from infested areas have certainly limited the movement of F. oxysporum f. sp. elaeidis among the major palm oilproducing areas of the world. Unfortunately, as the pathogen is slowed by regulation, so are efforts toward plant resistance breeding for long-range control of this disease. Brazilian isolates of F. oxysporum f. sp. elaeidis were demonstrated to be virulent against all African sources of resistance. It seems increasingly critical to broadly evaluate sources of tolerance and resistance against this genetically variable pathogen. Notable sources of resistance are known in Dura, Pisifera, and Tenera types.

Eradication measures have been a standard component of wilt management. Dead or dying trees are felled and burned on site, if possible. Soil in a 3-m radius of the diseased palm is treated with dazomet and covered for 30 days. Replanting plants 3.9 m off-center from infested stumps rather than 2.25 m has reduced wilt incidence by 50%. Heat and chemical treatments have not been effective for seed eradication of F. oxysporum f. sp. elaeidis, except for the use of the fungicide captafol as a 7-day seed soak.

The impact of oil palm production practices on F. oxysporum f. sp. elaeidis has been examined. Increased levels of potassium and magnesium disfavor the pathogen, while calcium levels have had no effect. The use of Brachiaria spp. as a cover crop has reduced wilt incidence compared with the use of Pueraria spp. The use of bare soil has reduced F. oxysporum f. sp. elaeidis incidence, while the use of the cover crop Calopogonium coeruleum (Benth.) Sauvalle has increased the pathogen’s incidence. The use of bunch straw mulch has increased wilt disease incidence, while the use of farmyard manure has reduced soil populations of the pathogen.

New technologies offer promise in support of traditional strategies of disease control. Palm tolerance to F. oxysporum f. sp. elaeidis has been correlated with the synthesis of fungal inhibitors in the tissue. These can be triggered by either inoculation with avirulent strains of F. oxysporum or by external application of analogues of fungal elicitors such as arachidonic acid. These may be useful to boost resistance expression. Similarly, chloroform/ethanol-soluble secondary metabolites from oil palm seed husks are antifungal toward F. oxysporum f. sp. elaeidis. These compounds offer potential use as seed treatments in nursery production, as a dip treatment at palm planting, or as a soil drench for infested sites.

Selected References Coremans-Pelseneer, J., Vanhaelen, M., and Vanhaelen-Fastre, R. 1991. Chemical resistance of Elaeis guineensis Jacq. seeds to Fusarium oxysporum Schl. elaeidis. Meded. Fac. Landbouwwet.

Rijksuniv. Gent 56(2a):185-187.

Flood, J., Mepsted, R., Valez, A., and Cooper, R. M. 1993. Comparison of Fusarium oxysporum f. sp. elaeidis from Africa and South America. Plant Pathol. 42:168-171.

Flood, J., Mepsted, R., and Cooper, R. M. 1994. Population dynamics of Fusarium species on oil palm seeds following chemical and heat treatments. Plant Pathol. 43:177-182.

Fraselle, J. V. 1951. Experimental evidence of the pathogenicity of Fusarium oxysporum Schl. to the oil palm (Elaeis guineensis J.).

Nature 167:447.

Ho, Y. W., Varghese, G., and Taylor, G. S. 1985. Fusarium oxysporum var. redolens from Africa as a cause of vascular wilt disease of oil palm. Phytopathol. Z. 113:373-376.

Holliday, P. 1970. Fusarium oxysporum f. sp. elaeidis. C.M.I. Description of Pathogenic Fungi and Bacteria 216. Commonwealth Mycological Institute, Kew, U.K.

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Locke, T., and Colhoun, J. 1973. Fusarium oxysporum f. sp. elaeidis as a seed-borne pathogen. Trans. Br. Mycol. Soc. 60:594-595.

Mouyna, I., Renard, J. L., and Brygoo, Y. 1996. DNA polymorphism among Fusarium oxysporum f. sp. elaeidis populations from oil palm, using a repeated and dispersed sequence ‘Palm’. Curr.

Genet. 30:174-180.

Prendergast, A. G. 1957. Observations on the epidemiology of vascular wilt disease of the oil palm (Elaeis guineensis Jacq.). J. West Afr. Inst. Oil Palm Res. 2:148-175.

Prendergast, A. G. 1963. A method of testing oil palm progenies at the nursery stage for resistance to vascular wilt disease caused by Fusarium oxysporum Schl. J. West Afr. Inst. Oil Palm Res. 4:156175.

Renard, J. L., and de Franqueville, H. 1991. Effectiveness of crop techniques in the integrated control of oil palm vascular wilt. Oléagineux 46:255-265.

Renard, J. L., Noiret, J. M., and Meunier, J. 1980. Sources and ranges of resistance to Fusarium wilt in the oil palms Elaeis guineensis and Elaeis melanococca. Oléagineux 35:387-393.

Taquet, B., Ravise, A., Renard, J. L., and Kunesch, G. 1985. Modulation des reactions de defense du palmier a huile contre le Fusarium oxysporum f. sp. elaeidis (Schlecht) Toovey. Phytopathol.

Z. 112:298-314.

Van de Lande, H. L. 1984. Vascular wilt disease of oil palm (Elaeis guineensis Jacq.) in Para, Brazil. Oil Palm News 28:6-10.

Wardlaw, C. W. 1946. Fusarium oxysporum on the oil palm. Nature 158:712.

(Prepared by G. W. Simone)

Ganoderma Butt Rot (Basal Stem Rot) This disease has many names, but it is always lethal to palms. In the continental United States, the disease is referred to as Ganoderma butt rot, and thus far it is limited to the southeastern region of the country. This includes all of Florida and the coastal areas of those states bordering the Atlantic Ocean and the Gulf of Mexico. It has not been documented in Mexico or the islands of the West Indies. In general, the natural geographic range of the disease in the United States matches that of Sabal palmetto. However, as infected, but nonsymptomatic, mature palms are moved out of this region for horticultural purposes, the disease has the potential to spread.

In tropical regions that grow Elaeis guineensis as a plantation crop or in wild groves, the disease is referred to as basal stem rot. It is considered the most serious disease of E. guineensis in Southeast Asia, especially in Indonesia and Malaysia, but also Papua New Guinea and Thailand. Other countries with records of basal stem rot on E. guineensis include Angola, Benin, Cameroon, Colombia, Democratic Republic of the Congo, Ghana, Honduras, India, Ivory Coast, Nigeria, Principé, Sao Tomé, Tanzania, Zambia, and Zimbabwe.

In India and Sri Lanka, the disease is a serious problem of Cocos nucifera plantations, where the disease is now known as basal stem rot but was previously referred to as Thanjavur wilt.

On Areca catechu plantations in India, the disease is known as anabe disease.

Symptoms Overall, symptoms are similar throughout the world, with slight variations based on the region where the disease is observed. This may be a function of the Ganoderma species associated with the disease in each region. In the United States, the primary foliar symptoms, either alone or in combination, are mild to severe wilt (Plate 44), reduced growth, overall off-color foliage (not chlorotic, just paler green than normal), and older fronds that are chlorotic or necrotic (Plate 45) and eventually droop (Plate 46). Similar symptoms are observed on E. guineen-

sis, but an excessive number of unopened spear leaves are also observed on mature palms, and young palms often have a onesided chlorosis and necrosis of the lower fronds.

By the time foliar symptoms develop, usually more than half of the lower internal stem tissue (trunk) has been killed by the fungus. Since a portion of the rotted trunk tissue includes xylem tissue, this accounts for the wilt symptoms observed. The formation of basidiocarps externally on the trunk tissue may or may not be associated with symptoms. It is common for basidiocarps to form prior to observable symptoms.

Cross sections of the lower trunk show an infected central area that is usually soft and distinct in color from the healthy tissue surrounding it (Plate 47). There is often a very distinct boundary between infected and healthy tissue, termed a reaction zone or black line. Serial cross sections of infected palm stem tissue show a disease progression pattern that is cone shaped, with the greatest portion of diseased trunk at the soil line and the disease progressing upward in the center of the trunk (Plates 48 and 49). The fungus does not normally extend more than 1.5–2 m up into the palm trunk.

Once the fungus has moved to the outside edge of the palm stem, basidiocarps may form external to the trunk, releasing basidiospores (Plate 50). In general, a palm stem dies within 1 year after basidiocarps begin to form on the trunk base. For palms with multiple stems (e.g., Dypsis lutescens), death of the palm takes longer than 1 year, and it is not uncommon to see new growth develop. Eventually, however, the palm dies.

Minimal root symptoms are normally associated with the disease in the continental United States while the palms are living, but roots of E. guineensis and C. nucifera in tropical areas are affected by the fungus. The roots are very friable, and their internal tissues become very dry and powdery, with the stele becoming black. However, this is observed on older roots that have been infected for a considerable time prior to the appearance of foliar symptoms.

C. nucifera in India and Sri Lanka often exude a reddish brown, viscous fluid from longitudinal cracks in the base of the palm trunk. This symptom of stem bleeding has also been observed on affected palms in the United States, but it is not a definitive diagnostic characteristic, since other diseases and disorders may cause the same symptom.

Causal Organisms The taxonomy and identification of Ganoderma spp. can best be described as chaotic over the past century. For example, 15 species of the genus Ganoderma have been recorded worldwide as probable causal organisms of basal stem rot of E. guineensis. One reason for this chaos is that morphological characteristics, especially of basidiocarps and mycelia, are not uniform and appear to vary with the environment. However, recent molecular systematic studies indicate that there may be some uniformity concerning the two primary species pathogenic on palms, G. zonatum Murr. and G. boninense Pat. While other Ganoderma spp. may be pathogenic to palms, these are the two most important economic species that are relatively host specific to palms.

In the United States, Ganoderma butt rot is caused by G. zonatum. Earlier literature lists G. sulcatum Murr. as a causal organism of this disease. The two species are now grouped together as G. zonatum. This species has also been implicated as the cause of basal stem rot of E. guineensis in Africa (Democratic Republic of the Congo, Ghana, Nigeria, Sao Tomé, and Tanzania), and its presence has been confirmed in southern South America.

In Southeast Asia and associated Pacific Island regions, the disease on E. guineensis is primarily caused by G. boninense.

G. zonatum and G. boninense differ molecularly and morphologically, and both species are distinct from G. lucidum (W. Curt.:Fr.) P. Karst., which is considered the pathogen of basal stem rot of C. nucifera in India. However, G. boninense is con-

sidered the pathogen of this disease on C. nucifera in Sri Lanka, despite the fact that symptoms are similar in both countries. The safest statement is simply that basal stem rot of palms outside the continental United States is caused by Ganoderma spp.

The basidiocarp, also referred to as a conk, sporophore, or basidiomata, of both G. zonatum and G. boninense initially begins as a white flat mass that is irregular to circular in shape.

As it develops, it begins to expand outward from the trunk into a ball shape, but it remains white and is relatively soft when touched (Plate 50). As the basidiocarp matures, it protrudes from the trunk, eventually forming a distinct, shelflike structure that is quite hard, with a glazed reddish brown top surface and a white undersurface that often swells at the edge (Plate 50). The top surface of G. zonatum has distinct zones of growth, hence the species name (Plate 51). The basidiocarp of G. zonatum is directly attached to the trunk (sessile) as opposed to the basidiocarp of G. boninense, which is formed on a stipe (stalk) protruding from the trunk. The size of the G. zonatum basidiocarp varies, but it can be upward of 20 cm at its widest point and 5 cm thick; the G. boninense basidiocarp is smaller in diameter (up to 11 cm). Also, the basidiocarp upper surface of G. zonatum is lighter in color with a thinner crust than that of G. boninense.

The undersurface of the basidiocarp is composed of microscopic pores or tubes where the basidiospores are formed and released. The basidiospores are ellipsoid, with a size range of 5–7 × 11–14 µm for G. zonatum and of 5–7 × 9–13 µm for G. boninense. The white undersurface of the basidiocarp becomes brownish red once the basidiospores are released from a mature basidiocarp because of the color of the spores en masse.

As with all Ganoderma spp., the basidiospore is double walled.

The basidiocarp is annual; once it matures and releases spores, it does not begin growing again. The old basidiocarp may remain on the trunk for another year or longer, but it is no longer fertile.

In southern, subtropical Florida, basidiocarps of G. zonatum are produced throughout the year. In more temperate areas, they are often only produced during the warmer months.

Basidiocarps are normally observed growing on the lower portion of a palm trunk or on a cut stump. However, the basidiocarps also protrude from the soil as the fungus begins to rot the palm roots of a dead or nearly dead palm; this is more common with G. boninense.

Host Range and Epidemiology Both G. zonatum and G. boninense appear to be fairly specific pathogens of palms. Other Ganoderma spp. cited as pathogens of palms have wider host ranges that include woody dicots. To date, the death of 58 palm species caused by G. zonatum has been documented in Florida (Table 3). While the majority of these palms were planted as ornamentals in the landscape, even naturalized Sabal palmetto has been killed by the fungus. For G. zonatum, all palm species having woody trunks should be considered potential hosts of this fungus.

There are many more palm species than those listed in Table 3, but they are not common in the continental United States and so may simply have escaped infection or have not been reported thus far. While G. boninense is considered a pathogen on E. guineensis and a saprophyte on C. nucifera in Malaysia, it is a pathogen on C. nucifera in Sri Lanka. Other palm hosts of G. boninense are not well documented.

Ganoderma spp. are wood-rotting fungi that decompose lignin, cellulose, and related polysaccharides. The exact infection point is unknown, but the trunk tissue is rotted internally. The fungus appears to colonize and degrade the palm trunk tissue closest to the soil line first and then expands in diameter at the base and moves up the center or near-center of the trunk. Once the fungus has moved to the outside edge of the palm trunk, basidiocarps may form external to the trunk, releasing basidiospores.

23

Although research on Ganoderma butt rot is still quite limited, the diversity and isolation of locations in Florida where the disease occurs (northwest to northeast to the most southern tip of the state) support the theory of spores as a primary method of spread. Spores had been discounted as an inoculum source in the E. guineensis plantations of Malaysia, but recent research suggests otherwise. However, infected palm materials (roots, stumps, boles, etc.) should not be overlooked as important methods of disease spread via mycelia, especially in a landscape dominated by palms.

In Florida, no common environmental conditions, soil types, or landscape management practices have been observed that favor the development of Ganoderma butt rot. The disease has

TABLE 3. Palm Hosts of Ganoderma zonatum in Florida Acoelorrhaphe wrightii (Griseb. & H. A. Wendl.) H. A. Wendl. ex Becc.

Acrocomia aculeata (Jacq.) Lodd. ex Mart.

Adonidia merrillii (Becc.) Becc.

Aiphanes sp., unidentified Arenga engleri Becc.

Arenga tremula (Blanco) Becc.

Arenga undulatifolia Becc.

Attalea sp., unidentified Bactris major Jacq.

Brahea berlandieri Bartlett Brahea brandegeei (Purpus) H. E. Moore Brahea dulcis (Kunth) Mart.

Brahea edulis H. A. Wendl. ex S. Watson × Brahea brandegeei (Purpus) H. E. Moore Butia capitata (Mart.) Becc.

Butia eriospatha (Mart. ex Drude) Becc.

Carpentaria acuminata (H. A. Wendl. & Drude) Becc.

Caryota mitis Lour.

Chamaerops humilis L. Coccothrinax sp., unidentified Cocos nucifera L. Copernicia curtisii Becc.

Dictyosperma album (Bory) Scheff.

Dypsis cabadae (H. E. Moore) Beentje & J. Dransf.

Dypsis lutescens (H. A. Wendl.) Beentje & J. Dransf.

Elaeis guineensis Jacq.

Euterpe edulis Mart.

Gastrococos crispa (Kunth) H. E. Moore Hyophorbe indica Gaertn.

Livistona benthamii F. M. Bailey Livistona chinensis (Jacq.) R. Br. ex Mart.

Livistona merrillii Becc.

Livistona muelleri F. M. Bailey Livistona saribus (Lour.) Merr. ex A. Chev.

Nannorrhops ritchiana (Griff.) Aitch.

Phoenix canariensis Chabaud Phoenix dactylifera L. Phoenix reclinata Jacq.

Phoenix roebelenii O’Brien Phoenix sylvestris (L.) Roxb.

Ptychosperma elegans (R. Br.) Blume Ptychosperma macarthurii (H. A. Wendl. ex H. J. Veitch) H. A. Wendl. ex Hook. f.

Ptychosperma salomonense Burret Roystonea altissima (Mill.) H. E. Moore Roystonea oleracea (Jacq.) O. F. Cook Roystonea regia (Kunth) O. F. Cook Sabal causiarum (O. F. Cook) Becc.

Sabal mauritiiformis (H. Karst.) Griseb. & H. A. Wendl.

Sabal palmetto (Walter) Lodd. ex Schult. & Schult. f.

Sabal uresana Trel.

Satakentia liukiuensis (Hatusima) H. E. Moore Serenoa repens (W. Bartram) Small Syagrus oleracea (Mart.) Becc.

Syagrus picrophylla Barb. Rodr.

Syagrus romanzoffiana (Cham.) Glassman Syagrus sancona (Kunth) H. Karst.

Syagrus schizophylla (Mart.) Glassman Syagrus × costae Glassman Washingtonia robusta H. A. Wendl.

24

been observed in natural settings (palms never transplanted) and in highly maintained, transplanted landscapes. It has been observed on palms that have been maintained very well nutritionally (no nutrient deficiencies) and on palms that were severely stressed by deficiencies. The disease has been observed in well-drained settings and in swamps. The fungus has killed trees that had no apparent mechanical injuries and those that had been severely damaged by, for example, weed trimmers. Soil type also appears to have no relationship with the disease, since diseased palms have been observed on deep sands (both silica and calcareous), muck (peat), and limestone rock. There has been no discernable pattern to provide clues as to which palms will become infected and die from G. zonatum.

Similar observations have been made for basal stem rot on E. guineensis.

Diagnostic Techniques Confirmation of Ganoderma butt rot cannot be made until the basidiocarp forms on a standing palm or until an unhealthy tree is cut down and the symptomatic internal trunk rotting is observed. Many of the external symptoms described previously can also be attributed to other biotic or abiotic causes. It is important to remember that Ganoderma spp., including G. zonatum, are also saprobic fungi. Therefore, development of Ganoderma basidiocarps on stumps or roots in the months after a palm has died and been cut down is not diagnostic.

There are currently no reliable diagnostic methods for isolating and identifying G. zonatum from trunk tissue, roots, or soil.

The selective medium developed for G. boninense has not been effective in isolating G. zonatum. Molecular and immunological methods have been developed to aid in the diagnosis of basal stem rot of E. guineensis, presumably G. boninense, in the Pacific Island region (e.g., Indonesia and Papua New Guinea).

Management In general, the fungus is predominantly located in the lower 1.5–2 m of trunk. This has three implications for ornamental palms. First, the fungus is not spread with leaf-pruning tools, since it is not associated with leaf tissue. Second, the lower trunk portion should not be chipped and used for mulch. If possible, the diseased section should be placed in a legal landfill or incinerated; it should not be sent to a plant material recycling center. The remaining portion of the palm trunk can be chipped and used for mulch in the landscape.

Third, only the lower 1.5–2 m of trunk needs to be protected from the fungus. However, typical systemic fungicides will not be effective unless they are capable of spreading beyond the vascular tissue and protecting all the internal tissue in the lower portion of the trunk. Currently, there is no fungicide with this capability. With no means of predicting or determining which palms are infected with G. zonatum, the use of fungicides as a control method, either preventively or curatively, is effectively eliminated for the present time. Therefore, there are no fungicide recommendations for Ganoderma butt rot. Field testing of fungicides for control of G. boninense (basal stem rot) also has not been successful.

Since basidiospores are probably the primary method of spreading the fungus, palms should be monitored closely, especially after a palm has died or been removed for any reason.

The fungus readily colonizes and rots palm stumps or dead palm trunks and, in this process, likely produces basidiocarps that will release millions of basidiospores (Plate 52). Palms with multiple stems (e.g., Dypsis lutescens) should not be thinned since the woody “stumps” will be colonized. Therefore, palms and palm stumps should be monitored for basidiocarps. The basidiocarp should be removed early in its formation (i.e., prior to spore release) and placed in a trash receptacle that will be incinerated or delivered to a landfill. It should not be placed in trash that will be recycled in the landscape. Palms

in the immediate landscape and neighborhood should be monitored at least once a month. Since the spores easily move with wind or water, it should be a community effort to reduce the spread of the spores of this lethal fungus.

Once a basidiocarp is observed on a palm, the palm should be removed, primarily for safety reasons since a significant portion of the trunk is already rotted. This is especially important during the hurricane season. When the palm is removed, as much as possible of the stump and root system should be removed. Any palm material left behind acts as a host for G. zonatum.

Since it is highly probable that the fungus survives in the soil, it is not recommended that a palm be replanted in a location where a palm has previously died from G. zonatum. Since only palms are affected by G. zonatum, it is safe to plant other species such as pines, oaks, woody shrubs, and other dicots. It is unknown as to how long the fungus survives in the soil.

If the area must be replanted with a palm, the stump and all roots should be removed from the site. Prior to replanting, the soil should then be fumigated with a registered fumigant for the landscape. However, this does not guarantee that the newly planted palms will remain free of G. zonatum.

Selected References Elliott, M. L., and Broschat, T. K. 2001. Observations and pathogenicity experiments on Ganoderma zonatum in Florida. Palms 45:62-72.

Flood, J., Bridge, P. D., and Holderness, M., eds. 2000. Ganoderma Diseases of Perennial Crops. CABI Publishing, Wallingford, U.K. Gilbertson, R. L., and Ryvarden, L. 1986. North American Polypores.

FungiFlora A/S, Oslo, Norway.

Gottlieb, A. M., and Wright, J. E. 1999. Taxonomy of Ganoderma from southern South America: Subgenus Ganoderma. Mycol. Res. 103:661-673.

Miller, R. N. G., Holderness, M., Bridge, P. D., Chung, G. F., and Zakaria, M. H. 1999. Genetic diversity of Ganoderma in oil palm plantings. Plant Pathol. 48:595-603.

Moncalvo, J.-M., Wang, H.-H., and Hseu, R.-Y. 1995. Phylogenetic relationships in Ganoderma inferred from the internal transcribed spacers and 25S ribosomal DNA sequences. Mycologia 87:223238.

Steyaert, R. L. 1967. Les Ganoderma palmicoles. Bull. Jard. Bot. Natl. Belg. 37:465-492.

Turner, P. D. 1981. Oil Palm Diseases and Disorders. Oxford University Press, Oxford, U.K. Utomo, C., and Niepold, F. 2000. Development of diagnostic methods for detecting Ganoderma-infected oil palms. J. Phytopathol.

148:507-514.

(Prepared by M. L. Elliott)

Gliocladium Blight (Pink Rot) Gliocladium blight (pink rot) is caused by the imperfect (Deuteromycota) fungus Gliocladium vermoeseni (Biourge) Thom. This pathogen has been reported on several palms and is cosmopolitan in distribution. It is commonly reported on Chamaedorea spp. as a sheath (leaf base), collar, and stem rot.

Invasion of the apical tips of young plants destroys many seedlings.

Symptoms G. vermoeseni causes leaf spots, rachis spots and decay, sheath rot, bud rot, and stem and trunk rots. Potted palms with infected sheaths (leaf bases) (Plates 53 and 54) produce chlorotic leaves. Loss of infected leaves causes the lower section of the plant to appear thin and barren, and marketability is reduced (Plate 55). Following entry of the pathogen into the stem, G. vermoeseni colonizes the host and moves within the

plant. Stem lesions are initially dark brown and often begin near the soil line. On Syagrus spp., the trunk can be infected at any height. Under moist conditions, especially when palms are grown close together, the pathogen causes stem rots that defoliate, wilt, or kill small palms, such as Chamaedorea spp. Diseased plants that survive produce progressively smaller, unthrifty leaves. Stems of larger palms are girdled and plants slowly decline.

The most characteristic sign of this disease is the pinkish orange mass of spores produced by G. vermoeseni (Plates 56 and 57). These masses, produced on sporodochia, are common on dead sections of palm trunks, stems, and petioles. A clear or gummy exudate is produced on the stem of some palms.

Causal Organism Gliocladium blight of ornamental palms was first reported in 1938, with the causal organism identified as Penicillium vermoeseni Biourge, which is now G. vermoeseni. This fungus produces single-celled, hyaline conidia in chains from phialides (Plate 58). In culture and on diseased host tissue, the conidial masses are pink to salmon. Conidia are 3–4 × 4–6 µm and are ellipsoidal when mature.

Host Range and Epidemiology G. vermoeseni has been reported on Archontophoenix cunninghamiana, Chamaedorea elegans, Chamaedorea erumpens, Chamaedorea seifrizii, other Chamaedorea spp., Chamaerops humilis, Dypsis decaryi, Dypsis lutescens, other Dypsis spp., Howea belmoreana, Howea forsteriana, Phoenix dactylifera, Syagrus romanzoffiana, and Washingtonia robusta. In 1975, D. E. Bliss isolated this fungus from a Howea sp., Phoenix canariensis, Syagrus romanzoffiana, and Washingtonia filifera in California. Potted Chamaedorea and Dypsis spp. appear to be the most frequently diseased palms in greenhouse settings, while Syagrus and Washingtonia spp. are the most severely affected in landscape plantings.

Good reproduction of the disease has been achieved on Chamaedorea spp. when sheaths are wounded before inoculation and when incubation and disease development occur under humid conditions. In a dry environment, few lesions develop without wounds. Wounded plants of Chamaedorea cataractarum, Chamaedorea elegans, and Chamaedorea seifrizii were all diseased following inoculation, while none of the Chamaedorea cataractarum or Chamaedorea elegans developed disease without wounds. For Chamaedorea seifrizii, 50% of the plants inoculated without wounds developed disease. On a Dypsis sp., a spray application of spores on unwounded plants resulted in a few light brown spots, 1–3 mm in diameter, primarily on the younger leaves.

The fungus is believed to enter the plant via wounds or damaged tissue, such as those caused by pruning, insect injury, mollusk feeding, sunburn, freezing, fertilizer burn, or pesticide phytotoxicity. The pathogen is invasive and rapidly colonizes weakened host tissue or young bud tissue. Following entry of the fungus through wounds, the host is colonized and large numbers of conidia are subsequently formed on the external plant surface. These spore masses provide the inoculum needed to move the disease to other plants. Spores are readily blown or splashed to healthy plants. Injured plants and high humidity favor pathogen establishment and disease progress.

Growth of G. vermoeseni appears to be strongly temperature dependent. Radial growth of the fungus in culture at 18 and 30°C was reduced by half when compared with growth at 24 and 27°C. Growth of the fungus did not occur at 33°C.

Diagnostic Techniques Generally, this disease is diagnosed by the tremendous number of pink- to salmon-colored spore masses formed on the surface of diseased plants. Spore masses can be small to large, and when examined microscopically, single-cell, hyaline spores 25

produced in chains are observed on phialides. Isolation is accomplished by washing specimens well, cutting sections of stem or sheath rots, dipping the sections in 0.5% sodium hypochlorite solution, and placing the sections on water agar. G. vermoeseni grows well on potato dextrose agar, corn meal agar, V8 juice agar, and many other media. Cultures should be maintained at 24–26°C, and sections with high sporulation levels should be transferred to preserve the sporulating sectors.

Management For greenhouse-grown palms, all dead leaves should be removed and each pot should be cleaned. Cleaning of the plants should continue as infected leaves mature and abscise. Tearing sheaths off the stem must be avoided. Immediately following cleaning operations, a general contact fungicide, such as maneb or mancozeb, or systemic fungicides, such as thiophanate methyl, should be applied to protect immature tissue and tissue injured during the cleaning process. Fungicides also reduce the amount of spores produced on existing lesions and kill spores on plant surfaces.

The spacing should be increased between plants in the greenhouse to promote air movement, reduce humidity, and provide better coverage by fungicides. Plants should be irrigated in the morning to avoid prolonged periods of wet foliage, or drip irrigation should be used. Producing crops under solid cover is also recommended in rainy climates. A good pest control program should be maintained, since injury by insects, slugs, and other pests promotes infection by G. vermoeseni.

For landscape palms, severe trimming of mature trees should be avoided. Cutting green petioles exposes highly susceptible tissue to G. vermoeseni and other fungal pathogens. If possible, leaf removal should be restricted to dry periods when temperatures exceed 30°C, when the fungus may be less active. If palms are heavily pruned, fungicides should be applied to protect wounds and exposed immature tissue.

Selected References Alfieri, S. A., Langdon, K. R., Kimbrough, J. W., El-Gholl, N. E., and Wehlburg, C. 1994. Diseases and Disorders of Plants in Florida.

Florida Dep. Agric. and Consumer Service, Div. Plant Industry.

Bull. 14.

Atliano, R. A., Llewellyn, W. R., and Donselman, H. M. 1980. Control of Gliocladium in Chamaedorea palms. Proc. Fla. State Hortic.

Soc. 93:194-195.

Marziana, F., Aloj, B., and Noviello, C. 1980. Chamaedorea elegans host of Gliocladium vermoeseni (Biourge) Thom in Italy. Inst. Patol. Veg. Dell. Univ. Napoli, Portici. Ser. IV 14:108-115.

Reynolds, J. E. 1964. Gliocladium disease of palm in Dade County, Florida. Plant Dis. Rep. 48:718-720.

(Prepared by J. Y. Uchida)

Graphiola Leaf Spot (False Smut) Graphiola leaf spot is a foliar disease widely known in the literature as false smut. Since this pathogen was described by J. B. Mougeot in 1823 as an ascomycete and later by P. A. Poiteau in 1824 as a myxomycete, it has been the focus of almost 160 years of taxonomic investigation. During this period, the pathogen has been reclassified as a pyrenomycete, as two distinct species in the order Uredinales, in the form class Hypomyces, in the order Ustilaginales, and finally into the order Exobasidiales.

Graphiola phoenicis (Moug.) Poit. is worldwide in distribution in palm-growing areas and is also reported from greenhouse botanical collections. Disease reports exist from the following countries: Algeria, Argentina, Australia, Austria, Barbados, Belgium, Brazil, Brunei, the Canary Islands, China, Colombia, 26

the Congo region, Cuba, Cyprus, the Czech Republic, Denmark, Dominican Republic, Egypt, El Salvador, Fiji, France, French Guiana, Germany, Ghana, Great Britain, Greece, India, Italy, Jamaica, Japan, Libya, Malaysia, Malawi, Mali, Mauritania, Mexico, the Netherlands, New Caledonia, New Zealand, Niger, Pakistan, Peru, the Philippines, Senegal, South Africa, Taiwan, Trinidad & Tobago, the United States (Alabama, Arizona, California, Florida, Georgia, Hawaii, Mississippi, Texas, West Virginia, Puerto Rico, and the Virgin Islands), Uruguay, Venezuela, and Zimbabwe.

Symptoms Symptoms are relatively obscure in contrast to the fungal signs. Initially, very small, yellow to brown or black spots develop on both leaf surfaces and the rachis (Plate 59). More spots occur on the upper surface of the leaf closest to the frond base. These spots eventually swell and rupture as the reproductive sori of the pathogen emerge from below the leaf epidermis.

At this time, the sori obscure the foliar symptom of the disease (Plate 60). Symptoms are more commonly observed on 2-yearold and older fronds. High disease incidence can cause premature senescence of older fronds.

Phoenix palms are also very susceptible to nutritional deficiencies. This is especially acute in Florida (United States), where P. dactylifera is prone to potassium deficiency. Symptoms of potassium deficiency are observed on the older leaves and first appear as very small, yellow-green spots, giving a speckled appearance. These symptoms can be confused with those of Graphiola leaf spot. While this disease may contribute to the shortened life of a Phoenix palm leaf, potassium deficiency is the more likely cause of premature frond senescence. See Physiological Disorders for information on nutrient deficiencies.

Causal Organism G. phoenicis is currently in its own family, Graphiolaceae, based in part on ribosomal DNA sequence comparisons. The conspicuous feature of G. phoenicis is the black, cup-shaped fruiting body (sorus) that has a 0.5- to 1-mm diameter at maturity (Plate 61). The peridium partially encloses the sterile and generative hyphae and defines the ostiole through which the sterile hyphae emerge.

Fascicles of generative hyphae arise from basal cells in the sorus. Generative hyphae produce elongate cells prior to the formation of a chain of barrel-shaped cells that sequentially act as basidia. The base of the generative hyphae is dikaryotic, with karyogamy occurring in the basidia prior to the development of lateral primary spores. Meiosis is assumed to occur in each successive basidium prior to its separation from the generative hypha. The primary spores are 4.5–5.8 × 5.3–6.3 µm, appear yellowish brown in mass, and are dispersed through the sorus ostiole by the hygroscopic action of the sterile filaments.

These filaments may extend 2.5 mm beyond the sorus (Plate 61). Each spore is two-celled, is warty in texture, and readily disassociates at the septum. Each daughter cell is 2–2.5 × 3.8– 7.5 µm and may germinate by hypha or by budding in a yeastlike manner.

Host Range and Epidemiology G. phoenicis is reported on 28 palm species (Table 4). In addition, there are at least four other species of the genus Graphiola reported on palms worldwide: G. congesta Berk. & Rav. on Coccothrinax argentata, Coccothrinax argentea (Lodd. ex Schult. & Schult. f.) Sarg. ex Becc., Sabal adansonii Guers., Sabal mexicana, Sabal palmetto, and Serenoa repens in the southeastern United States; G. borassi Syd. & E. J. Butler on Borassus flabellifer in India; G. cylindrica Y. Kobayasi on Livistona chinensis and Trachycarpus fortunei in Japan; and G. thaxteri E. Fischer on Sabal megacarpa (Chapm.) Small and Sabal mexicana in Florida. Fischer has described six additional species, for which little information exists.

Hyphal germination and penetration of open stomata is the assumed infection route, since no yeast stage has been observed in vivo. Infection hyphae are slender (1–1.3 µm) and believed to rupture thin-walled hypodermal cells within the stomatal cavity. The fungus grows inter- and intracellularly in a limited area directly beneath the site of sorus development. The time span from infection to reproduction is 10–11 months. Sori emerge from interveinal tissue or along veins in leaflets and may be solitary or clustered, reaching densities of more than 20 per cm2 of leaflet. Although infection can occur on both leaf surfaces, sori development is greatest on the upper leaf surface. Disease occurrence is most often associated with humid palm production areas.

Diagnostic Techniques The black sori of G. phoenicis are diagnostic for this disease—especially if a yellowish fascicle of spore-bearing hyphae is visible. Immature sori can sometimes be induced to sporulate by treatment for 48 hr in a high-humidity chamber.

The pathogen can be cultured from the yellow secondary spores off the sterile hyphae on a variety of media, such as potato sucrose agar, oatmeal agar, or malt agar. Germination occurs in 6–8 hr, producing 1- to 2-mm-diameter, pink, yeastlike colonies with smooth margins. Spores are 1.5–2 × 5–7 µm. Hyphal growth initiates after 24 hr, but vegetative growth is generally slow (8–10 mm in 10 days).

Management Damage from G. phoenicis is documented in commercial P. dactylifera plantings. High densities of lesions can cause premature frond senescence, reducing normal frond life of 6–8 years to 3 years. Reducing planting density from 4 × 5 m to 8 × 8 or 10 × 10 m reduced disease incidence in commercial P. dactylifera plantings in Brazil. Palms grown in the landscape should be properly spaced to maximize air circulation. Judicious pruning and destruction of infected plant parts contributes to disease management. Irrigation systems should be directed below the palm canopy in the maintenance of other landscape materials.

Nursery use of irrigation should be timed for early morning hours to minimize the period of leaf wetness.

P. dactylifera cultivars have shown differences in disease reaction to G. phoenicis. Cultivars reported as resistant include

TABLE 4. Palm Hosts of Graphiola phoenicis Acoelorrhaphe wrightii (Griseb. & H. A. Wendl.) H. A. Wendl. ex Becc.

Arenga pinnata (Wurmb) Merr.

Butia capitata (Mart.) Becc.

Chamaerops humilis L. Coccothrinax argentata (Jacq.) L. H. Bailey Cocos nucifera L. Dypsis lutescens (H. A. Wendl.) Beentje & J. Dransf.

Livistona alfredii F. v. Muell.

Livistona chinensis (Jacq.) R. Br. ex Mart.

Phoenix canariensis Chabaud Phoenix dactylifera L. Phoenix loureirii Kunth Phoenix paludosa Roxb.

Phoenix reclinata Jacq.

Phoenix roebelenii O’Brien Phoenix sp., unidentified Phoenix sylvestris (L.) Roxb.

Phoenix theophrasti Greuter Prestoea acuminata (Willd.) H. E. Moore Roystonea regia (Kunth) O. F. Cook Roystonea sp., unidentified Sabal minor (Jacq.) Pers.

Sabal palmetto (Walter) Lodd. ex Schult. & Schult. f.

Sabal sp., unidentified Syagrus romanzoffiana (Cham.) Glassman Thrinax morrisii H. A. Wendl.

Washingtonia robusta H. A. Wendl.

Washingtonia sp., unidentified

Aglanee, Barakawy, Barhee, Jozee, Khadrawy (from northern Iraq), Kustawy, and Tadala. Intermediate reactions are expressed by the cultivars Amirhajj, Ammary, Amry, Fursi, Halawy, Medjool, Samain, Sayir, and Thoory. Many susceptible cultivars have been reported in the literature and include the following: Anhat, Bedraya, Braim, Dayri, Deglet Noor, Hallawi, Hayani, Koroch, Shamran, and Zahidi.

In the United States, mature P. dactylifera are transported from arid date production sites in California for landscape use in humid environments of Florida and other southeastern states.

Currently, 75% of California’s date crop is derived from the susceptible cultivar Deglet Noor. Another 20% of the California crop is derived from the cultivar Medjool. Zahidi is the third cultivar most often found in the date production areas.

Therefore, incidence and severity of Graphiola leaf spot can be expected to increase with the widespread use of susceptible cultivars under the humid environmental conditions of the southeastern United States.

Fungicides are also used for G. phoenicis control. Field tests in India demonstrated that products with carbendazim, mancozeb, cupric hydroxide, copper oxychloride, thiophanate methyl, or zineb were effective in disease management, when timed for the spore-release period. Greenhouse trials in Italy and the Czech Republic demonstrated that propineb, oxycarboxin, biteranol, and benomyl provided excellent control of G. phoenicis indoors. In nursery and landscape situations, fungicides may be needed for disease control. Properly labeled products should be applied as preventives during the reproductive period for this pathogen, usually during periods of moist conditions.

Selected References Begerow, D., Bauer, R., and Oberwinkler, F. 1997. Phylogenetic studies on nuclear large subunit ribosomal DNA sequences of smut fungi and related taxa. Can. J. Bot. 75:2045-2056.

Cole, G. T. 1983. Graphiola phoenicis: A taxonomic enigma. Mycologia 75:93-116.

Farr, D. F., Rossman, A. Y., Palm, M. E., and McCray, E. B. Fungal Databases, Systematic Botany & Mycology Laboratory, ARS, USDA, at http://nt.ars-grin.gov/FungalDatabases/DatabaseFrame.cfm.

Gafar, K., Monem, S. Abed El, Said, K., Necola, N., Badr, N., and Megid, K. Abd El. 1979. Studies on false smut of date palm trees and its control. Agric. Res. Rev. 57:1-8.

Lima, M. F. 1996. First report of Graphiola leaf spot caused by Graphiola phoenicis on date palm (Phoenix dactylifera) in the state of Pernambuco, in the northeast of Brazil. Plant Dis. 80:823.

Mehta, N., Gupta, P. C., Thareja, R. K., and Dang, J. K. 1989. Varietal behavior and efficacy of different fungicides for the control of date palm leaf spot caused by Graphiola phoenicis. Trop. Pest Manage. 35:117-119.

Nixon, R. W. 1957. Differences among varieties of the date palm in tolerance of Graphiola leaf spot. Plant Dis. Rep. 41:1026-1028.

Oberwinkler, F., Bandoni, R. J., Blanz, P., Deml, G., and KisimovaHorowitz, L. 1982. Graphiolales: Basidiomycetes parasitic on palms. Plant Syst. Evol. 140:251-277.

Perisic, M., Babovic, M., and Pantic, A. 1985. A study on Graphiola phoenicis (Moug.) Poit., a parasite of the palm Phoenix canariensis Chauband. Mikrobiologija (Belgr.) 22:165-169.

Polizzi, G., and Agosteo, G. E. 1995. Efficacy of natural fungicides and chemicals in the control of Graphiola phoenicis, causal agent of false smut of palms. Dif. Piante 18:122-126.

(Prepared by G. W. Simone)

Pestalotiopsis Diseases Pestalotiopsis Steyaert spp. can cause diseases of numerous palm tissues. When the pathogen causes a leaf spot, it is referred to as Pestalotiopsis leaf spot or, less commonly, gray leaf blight. Palm diseases reported to be associated with Pesta27

lotiopsis and Pestalotia De Not. spp. are common because dark, multicelled spores with appendages are common on many decaying or diseased plant tissues. While these genera are easily identifiable as a general group, the nomenclatural confusion between these genera and the superficial similarity of these genera make a review difficult.

Symptoms Leaf spots associated with Pestalotiopsis spp. begin as tiny, black spots that expand to about 2 mm long and are elongate, elliptical to circular lesions, depending on the host. These lesions may expand to form small blights that are bordered by black tissue, with some chlorosis between lesions (Plate 62).

Lesions also occur on the rachis and petioles (Plate 63). Dead tissue is often tan to light gray and very thin. In addition to leaf spots, blights on a Chamaerops sp., Roystonea regia, and Syagrus romanzoffiana; a crown rot of Phoenix roebelenii (Plate 64) and a Syagrus sp.; and a general decline of Butia capitata have been associated with Pestalotiopsis spp. Pestalotiopsis spp. invade the trunks, sheaths, and petioles of Areca catechu (betel nut palm) by infecting wounds created by insects (Plates 65 and 66).

Causal Organism Fungi casually identified as Pestalotia spp. may be Pestalotiopsis spp. Fungi identified as Pestalotiopsis spp. may have two or more, unbranched, apical, conidial appendages and four conidial septations (five-celled) with a hyaline basal cell. Conidia are distinctive with three dark cells in the middle of the spore and two to four appendages at the apical end. Conidia are produced on groups of conidiophores. Pestalotia spp. are taxonomically described as having three to nine, simple or branched, apical appendages; five septations (six-celled); and branched appendages on the basal cell. There are other important differences between these two genera, such as conidial wall structure, that are used less frequently.

While there are numerous Pestalotiopsis spp. associated with palms, Pestalotiopsis palmarum (Cooke) Steyaert is the most common. Conidia are fusiform, are usually straight, have four septations with slight constrictions at the septa, and are 4.5–7.5 × 17–25 µm. The apical appendages are 5–25 µm long.

Host Range and Epidemiology Pestalotiopsis palmarum has been recovered from numerous palms in the United States and from Cocos nucifera and Elaeis guineensis worldwide. Records of Pestalotia spp. can also be found, and many of these may have been of the genus Pestalotiopsis. Pestalotia spp. have been reported on more than 28 palms. Pestalotiopsis spp. have been reported on 39 palm species in Florida, with Pestalotia palmarum listed as synonymous to Pestalotiopsis palmarum in 35 of the 39 listed species. Leaf spots associated with Pestalotiopsis palmarum have been reported on Adonidia merrillii, Arenga spp., Bismarckia nobilis, Butia capitata, Caryota mitis, an unidentified Caryota sp., Chamaedorea elegans, Chamaedorea erumpens, Chamaerops humilis, Coccothrinax spp., Cocos nucifera, Dictyosperma album, Dypsis decaryi, Dypsis lutescens, a Hyophorbe sp., Livistona chinensis, Phoenix canariensis, Phoenix dactylifera, Phoenix reclinata, Phoenix roebelenii, Pinanga spp., Pseudophoenix spp., Ptychosperma elegans, Ptychosperma macarthurii, Rhapidophyllum hystrix, Rhapis excelsa, an unidentified Rhapis sp., Roystonea regia, Sabal palmetto, Serenoa repens, Syagrus romanzoffiana, Thrinax spp., Trachycarpus fortunei, and Washingtonia robusta. Additionally, an unidentified species of the genus Pestalotiopsis has been reported on Areca catechu and on Carpentaria, Howea, Ravenea, and Syagrus spp.

Pathogenicity was confirmed for a Pestalotiopsis sp. on wounded Areca catechu and Howea forsteriana. The fungus did not penetrate nonwounded tissue. Secondary Pestalotiopsis invasion of leaf spots caused by Bipolaris incurvata (C. 28

Bernard) Alcorn was also confirmed on Cocos nucifera. In this study, the Pestalotiopsis sp. did not invade Cocos nucifera leaves without existing lesions. In the Philippines, Pestalotiopsis spp. invaded wounds caused by a leaf miner on Cocos nucifera.

Based on the known disease cycles of other imperfect fungi and the presence of spores borne on acervuli on older lesions, these spores are likely to splash to wounded leaves or other damaged tissue and invade palm hosts. Since Pestalotiopsis spp. are associated with insect wounds, spores are likely to be dispersed by insects also. High humidity is expected to favor invasion by Pestalotiopsis spp. In general, Pestalotiopsis spp. do not penetrate leaves without wounds in pathogenicity studies. B. incurvata lesions are secondarily invaded by Pestalotiopsis spp. and became enlarged, changing from a dark brown spot to a straw-colored lesion with a broad, dark brown margin.

Thus, health of the foliage and protection from other leaf pathogens must be a priority.

Diagnostic Techniques Pestalotiopsis spp. are readily isolated from diseased tissue.

Infected leaves should be washed, sectioned into pieces about 2–4 mm long, disinfested in 0.5% sodium hypochlorite solution, blotted dry, and placed on the surface of water agar or acidified water agar. Single hyphal tips can be transferred from water agar plates to various nutrient agars, such as V8 juice agar and potato dextrose agar. Colonies are white. Sporulation begins after 1 week or longer, and spores are produced on groups of conidiophores.

If isolation media are not available, the diseased tissue should be washed well, placed in a moist chamber (glass dish or plastic bag with a moist tissue), and incubated at 24–26°C in the light. Conidia form in 1–3 days on older lesions.

Management Severely diseased leaves should be removed and destroyed on palms grown in greenhouses for the ornamental trade. Cultural controls such as increasing air movement to reduce humidity, increasing plant spacing, removing weeds or trimming surrounding vegetation, using solid-covered greenhouses, and timing water application to avoid prolonged periods of leaf wetness should be used. Application of protective, broad-spectrum fungicides should help to keep young leaves healthy. Given the known invasion by Pestalotiopsis spp. of leaf spots caused by B. incurvata, a disease management program that controls B. incurvata and other known fungal leaf pathogens should be used.

For palms in the landscape, the cultural controls suggested above should be used to reduce disease levels. Palms should be kept nutritionally healthy and insect pests should be controlled to avoid invasion of damaged tissue by Pestalotiopsis spp. Insecticides and fungicides should be applied to wounds invaded

by Pestalotiopsis spp. The time for trimming operations should be selected to allow wounded tissue to dry as rapidly as possible, e.g., trimming mature trees during periods of wet conditions should be avoided. If wet weather cannot be avoided, fungicide should be applied immediately on trimmed petioles or trunks.

Selected References Alfieri, S. A., Langdon, K. R., Kimbrough, J. W., El-Gholl, N. E., and Wehlburg, C. 1994. Diseases and Disorders of Plants in Florida.

Florida Dep. Agric. and Consumer Service, Div. Plant Industry.

Bull. 14.

Brown, J. S. 1975. Investigation of some coconut leaf spots in Papua New Guinea. Papua New Guinea Agric. J. 26:31-42.

Sutton, B. C. 1980. Pages 263-265 in: The Coelomycetes: Fungi Imperfecti with Pycnidia, Acervuli and Stroma. Commonwealth Mycological Institute, Kew, U.K.

(Prepared by J. Y. Uchida)