Corn Integrated Pest Management
Integrated Pest Management (IPM) is a holistic approach to monitoring pest lifecycles and prevalence in farmer’s fields to determine the best pest control methods to protect crop yield potential in the current and following growing seasons. IPM is a continuum of pest management evaluations, decisions, and controls that encompass pest lifecycles and prevalence, agronomic practices such as crop rotation, seed selection, soil management, and timely use of pesticides to minimize pest damage and protect environmental resources.

Red Root Rot a Late Season Disease of Corn * Roots and basal stalk tissue infected with red root rot characteristically have reddish-pink, rotted roots. * Stalks are weakened and susceptible to lodging. * Premature plant death is common and can occur quickly and yield losses can be as high as 15-20%. * Genetic resistance to red root rot is uncommon and breeding for disease resistance is difficult.
Management

Management options are limited. Crop rotation with a non-host such as soybean can provide some control.1 Genetic resistance has been difficult to incorporate into corn products, although the rate of disease development varies greatly between corn products. Research on inheritance of disease resistance indicates that it is a polygenic trait with additive gene action, which has complicated breeding efforts.1 Environmental stress during the season may contribute to disease infection and severity. Sap Beetles in Corn

Sap beetles are considered minor pests of corn. Adults prefer to feed on corn kernels, ear tips, and stalks that have previously been injured by other insects such as corn earworm or corn borer larvae. Controlling corn ear pests should prevent sap beetles from becoming a problem.
Damage
Sap beetles are secondary pests of corn and usually are attracted to corn ear tips or stalks that are damaged by corn earworm or corn borer feeding.2 The beetles may also enter into undamaged ears anytime from early silk to maturity. Both adults and larvae can damage ears. Larvae may hollow out kernels of the upper half of the ear.3
Scouting and Management
Feeding from sap beetles can be confused with other insects. When scouting for other insects, look for sap beetles. If sap beetles are numerous and other insects are not found, feeding damage may be from sap beetles. In addition, look for sap beetle larvae on corn ears (Figure 2). If needed, your local area agronomist can help assist in identifying what type of feeding damage is present in a particular field.
Sap beetle control is rarely justified in field corn because the injury caused by these beetles is usually negligible and of little economic significance. Controlling corn ear pests such as corn earworm and corn borer should prevent sap beetles from becoming a problem.4
Herbicide Rotation Best Management Practices * Excess rains have led to changes in planting decisions. * Some fields may have burndown, corn, or soybean herbicides already applied. * Labels need to be reviewed for rotational restrictions before planting crop products. * Active ingredients with rotational restrictions may also be in pre-packed herbicide mixes.

Fallow

Some fallow fields may have received excess rains after an applied burndown treatment and the fields remained unplanted. The unplanted ground may now need another burndown treatment, and a common tank mix partner for burndown ahead of corn and soybean is 2,4-D. 2,4-D labels should be read as they vary in several ways, including plant back restrictions. In general, 2,4-D at 1 pt./acre needs to be applied 7 days prior to planting corn or soybean.

Crop Replant

Replanted acres may be a scenario of corn being planted over a poor corn stand that was taken out, or more commonly now, a soybean crop being planted into fields intended for corn. Herbicides that were safe for corn may have already been applied. Late planting has led some growers to switch to soybean and it is important to review labels for rotational intervals. Many active ingredients with rotational restrictions are also in pre-packed herbicide mixes with various trade names (Table 1).

Corn Safety

The acetanilide herbicide family of active ingredients is used predominantly in corn in preemergence applications. Acetanilide herbicides sold in the United States are based on the following active ingredients- * Acetochlor (herbicides such as Degree Xtra®, Harness®, Harness® Xtra, TripleFLEX® Herbicide, and Warrant® Herbicide) * (S-)Metolachlor (herbicides such as Dual Magnum®, and Cinch®)
Dimethenamid (herbicides such as Outlook®)

Acetanilides break down in soil through microbial degradation. In tolerant corn plants, enzymes and proteins rapidly bind and inactivate acetanilide herbicides. A primary difference among acetanilide herbicides is their speed of binding. INTEGRATED PEST MANAGEMENT OF CABBAGE

| Cabbage LooperScientific Name: Trichoplusia ni | * | |
DESCRIPTION OF THE PESTLooper caterpillars can be distinguished from most other common caterpillars in cole crops by their distinctive looping movement in which they arch the middle portion of their body to bring the prolegs or hind legs forward to meet the front legs. Loopers are green, usually with a narrow white stripe along each side and several narrow lines down the back. Loopers are smooth-skinned with only a few long bristles down the back; they may grow up to 1.5 inches long. Mature larvae spin silken cocoons and pupate, usually attached to leaves. Adults are brownish moths with a distinctive silvery figure-8 on the front wings. Eggs are ridged and dome-shaped and usually laid singly on the undersurface of leaves. Loopers may have numerous generations and continue to develop all year long in cole crops growing areas of California with the highest populations usually occurring in fall.DAMAGEAlthough seedlings are occasionally damaged, most injury occurs after heading. Loopers eat ragged holes into leaves, bore through heads and contaminate heads and leaves with their bodies and frass. Young plants between seedling stage and heading can tolerate substantial leaf damage without loss of yield.MANAGEMENTCabbage loopers have many natural enemies that frequently keep loopers below economic levels, at least until heading, if they are not killed by insecticide treatments for other pests. Monitor to determine population levels of loopers and natural enemies and to determine the need for treatment following heading. If treatment is needed, use a selective material such as Bacillus thuringiensis. Biological Control
Important parasites include the egg parasite Trichogramma pretiosum, the larval parasites Hyposoter exiguae, Copidosoma truncatellum, and Microplitis brassicae, and the parasitic tachinid fly Voria ruralis. A nuclear polyhedrosis virus disease is also important under certain circumstances; the bodies of diseased caterpillars turn into shapeless sacks of dark liquid and can often be spotted hanging from leaves. Be sure to monitor for natural enemies; if looper populations are close to treatment thresholds but you find a significant percentage of parasitized or disease-killed individuals, delay treatment for a few days to see if these natural controls will bring populations down on their own. If treatment is necessary, use of Bacillus thuringiensis insecticide will minimize injury to natural enemies. Organically Acceptable Methods
Biological control and sprays of Bacillus thuringiensis and the Entrust formulation of spinosad are organically acceptable management tools. Monitoring and Treatment Decisions
Check 25 plants selected randomly throughout the field. Look for eggs and small larvae on the underside of lower leaves. If you find holes, search the general area for the caterpillar, opening damaged heads as necessary. Although damage can give you a general idea of where loopers may be and the seriousness of the infestation, do not base treatment on damage levels. Base treatment on numbers of healthy larvae present (include imported cabbageworms in counts, too, if they are also present). Treat seedlings or small plants if populations of medium-sized to large caterpillars are large enough to stunt growth. Before heading, well-established plants do not need to be treated unless you find more than 9 small- to medium-sized larvae per plant. Treat just before heading or at Brussels sprouts formation if counts show more than one looper or other caterpillar in 25 plants. Where possible, use a selective insecticide to avoid adverse impacts on natural enemies. Bacillus thuringiensis and most other selective insecticides are very effective against cabbage loopers, especially when applied to early-instar caterpillars (i.e., very young). Cabbage loopers are also controlled with the more toxic materials recommended for use against other lepidopterous (caterpillar) pests. If significant numbers of other caterpillars (armyworms or diamondback moths) are present, the use of a carbamate or pyrethroid may be warranted. |

Insect/Mite Management
Insect/Mite Pests
Insects are the principal pests found on cabbage in Florida. The greatest insect problem for Florida growers is the diamondback moth (Plutella xylostella). Cabbage looper (Trichoplusia ni) is also considered a major pest, but it has been less of a problem over the past decade. Some insect pests have been considered major in the past but are only occasional problems now, including aphids, beet armyworm (Spodoptera exigua), cabbage webworm (Hellula rogatalis), imported cabbageworm (Artogeia rapae), cutworms, and mole crickets. All of the minor pests on Florida cabbage have the potential to become major pests, but they are currently being controlled by treatments for diamondback moth. Silverleaf whitefly (Bemesia argentifolii) and vegetable leafminer (Liriomyza trifolii) are more significant in southern Florida, where silverleaf whitefly is considered a major pest and vegetable leafminer a minor pest (Hayslip et al. 1953; Webb 2010).
Additional insect pests that occasionally cause minor damage to Florida cabbage include southern armyworm (Spodoptera eridania), yellow-striped armyworm (Spodoptera ornithogalli), fall armyworm (Spodoptera frugiperda), blister beetles (Epicauta spp. and Macrobasisunicolor), cabbage budworm (Hellula phidilealis), corn earworm (Helicoverpa zea), cross-striped cabbageworm (Evergestis rimosalis), flea beetles (Phyllotreta spp.), grasshoppers, gulf white butterfly (Ascia monuste), harlequin bug (Murgantia histrionica), horned squash bug (Anasa armigera), onion thrips (Thrips tabaci), saltmarsh caterpillar (Estigmene acrea), southern cabbageworm (Pontia protodice), southern green stinkbug (Nezara viridula), spotted cucumber beetle (Diabrotica undecimpunctata howardi), tarnished plant bug (Lygus lineolaris), vegetable weevil (Listroderes costirostris obliquus), and yellow-margined leaf beetle (Microtheca ochroloma) (Leibee 1996; Webb 2010).
Diamondback Moth (Plutella xylostella)
Diamondback moth became the principal pest on Florida cabbage in the 1980s, and it remains one of the most serious problems for the state’s cabbage growers, occurring annually (Leibee 1996; Webb 2010). Plants at all stages of growth may be attacked. The moth lays its eggs in groups of 2–3 on the lower surface of leaves. In approximately one day the eggs hatch, and the larvae begin to feed on the leaves. Feeding results in many small holes that grow larger as the larvae increase in size. Often, feeding does not go through the entire leaf, leaving a thin layer of the leaf epidermis. In addition to the leaves, diamondback moth larvae can attack the developing cabbage heads, producing shallow tunnels on the tops of heads. The resulting damage deforms the heads and leaves entry points for decay pathogens (Hayslip et al. 1953).
The larval stage can range from 10 days to a month, depending on temperature. Diamondback moth larvae slow their feeding at temperatures below 50°F (10°C), and population growth is most rapid at temperatures greater than 80°F (26.7°C). The pupal stage is passed within a transparent, loose cocoon, usually attached to the underside of leaves. The moths emerge within 1–2 weeks after entering the pupal stage (Hayslip et al. 1953).
Diamondback moth is most abundant in southern Florida from December to February or March and can attack at any time during the crop cycle. By the end of May, moth counts in pheromone traps fall to near zero. Moth counts may rise in mid-fall through early winter, but activity is limited during that time. Populations build on winter weeds, such as wild mustard, before moving into winter and early spring cabbage plantings. From mid-winter through spring, diamondback moth is a serious pest, causing losses of up to 70% in the absence of control (Nuessly and Hentz 1999).
Aphids
Turnip aphid (Hyadaphis erysimi) and green peach aphid (Myzus persicae) are the most important aphids on Florida cabbage. Green peach aphid is a vector of turnip mosaic virus in Florida (Webb 2010).
Although cabbage aphid (Brevicoryne brassicae) may attack the crop at any stage, green peach aphid attacks cabbage mainly before heading begins. Aphids suck plant juices with their piercing-sucking mouthparts, resulting in yellowing and curling of leaves. Particularly when attacked as seedlings, plants may be stunted or die as a result of aphid feeding. Aphids can be protected from insecticide sprays within the curled leaves or inside the cupped leaves of headed plants (Hayslip et al. 1953).
Beet Armyworm (Spodoptera exigua)
Beet armyworm is a sporadic pest on Florida cabbage, and it is usually kept under damaging levels by controls targeted to diamondback moth. Beet armyworm populations in southern Florida are highest from late March through mid-June, with a small rise in population from mid-August through October. Population rise in the late summer and fall is thought to be related to beet armyworm activity on late summer weeds, while the population increase in the spring coincides with the leafy vegetable production season in southern Florida (Nuessly and Hentz 1999).
The beet armyworm has a wide host range, and, besides cabbage, it also attacks vegetables such as asparagus, bean, beet, broccoli, cauliflower, celery, chickpea, corn, cowpea, eggplant, lettuce, onion, pea, pepper, potato, radish, spinach, sweet potato, tomato, and turnip, and field crops such as alfalfa, corn, cotton, peanut, safflower, sorghum, soybean, and tobacco. Many weeds also serve as hosts, including lambsquarters (Chenopodium album), pigweeds (Amaranthus spp.), purslane (Portulaca sp.), parthenium (Parthenium sp.), and mullein (Verbascum sp.). Larvae feed on both foliage and fruit of host plants. On cabbage, beet armyworm larvae consume greater amounts of leaf tissue than the diamondback moth but not as much as the cabbage looper. An action threshold of 0.3 beet armyworm larvae per plant has been used for cabbage in Texas. Since adults can readily invade a field from nearby crops or weeds, monitoring the crop twice a week for beet armyworm presence and damage is recommended (Capinera 2011a).
The insect is active the entire year in southern Florida, and it migrates annually into north Florida and rest of the southeastern United States (Nuessly and Hentz 1999). Females can lay up to 600 eggs each, usually in groups of about 100. Eggs are laid on the underside of lower leaves and are covered with fuzzy, white scales. Under warm conditions, eggs hatch within 2–3 days. The larvae feed from one to three weeks, in groups when younger and scattered on the plant when larger. Full-grown larvae pupate in the soil in a cocoon constructed from sand and bits of soil. Adults emerge within a week in warm temperatures (Sorenson and Baker 1983; Capinera 2011a).
Cabbage Webworm (Hellula rogatalis)
Like beet armyworm, cabbage webworm is seen sporadically and is controlled by treatments for diamondback moth. It is more of a problem in southern Florida. The pest can attack cabbage both in seedbeds and the field (Hayslip et al. 1953).
Cabbage webworm eggs are usually laid in plant buds. Upon hatching, the larvae feed on the underside of the leaves in the bud area, producing small holes. The larvae cover themselves with webs, which become covered with dirt and excrement. Larger larvae can burrow into buds, stems, and leaves. The insect may feed on the growing point, which prevents head formation, and the plant may appear lopsided. When fully grown, larvae pupate in the buds, on the sides of stems, or on the soil surface (Hayslip et al. 1953).
Imported Cabbageworm (Artogeia rapae)
Another minor pest, imported cabbageworm produces large holes in leaves and may attack the head near maturity, leaving damage similar to diamondback moth (Workman 1983). Imported cabbageworm feeding results in large, irregular holes in the leaves and the head’s outer layers (Hayslip et al. 1953).
Female moths can lay several hundred eggs each within a cabbage field, attaching them to the underside of leaves. The egg stage lasts about one week, and the emerging caterpillars feed on the underside of the leaves. After about two weeks, the larvae attach themselves with silk to a support and pupate. The moths emerge in 1–2 weeks (Hayslip et al. 1953).
Cutworms
Cutworms affecting cabbage include black cutworm (Agrotis ipsilon) and granulate cutworm (Feltia subterranea). Cutworms are stout, gray caterpillars with a greasy appearance. They are active at night, feeding on cabbage stems and leaves and other plants. During the day, they take refuge in the soil at the base of the plants. Recently transplanted cabbage is particularly susceptible to attack by cutworms, which can cut thin-stemmed plants off at or slightly below the soil surface. They also cut large holes in leaves touching the soil surface (Hayslip et al. 1953). Several plants in a row are usually affected, and when feeding, cutworms often pull the end of a leaf into a protected area of the soil (Workman 1983). Cutworms also eat into heading cabbage and may remain within the head during the day. Overall, while some damage to leaves and heads occurs, greatest losses from cutworm damage are the result of reduced stands (Hayslip et al. 1953).
Black cutworm is one of the most destructive cutworms and attacks a wide range of plants. Although cutworm larvae can migrate into a field from adjacent areas, most migration occurs by adults flying into the field. The moth deposits eggs in groups of 1–30 on leaves or stems near ground level. The egg stage lasts from 5 to 15 days, the larval stage lasts from three to four weeks, and the pupal stage takes 12 to 36 days. At high temperatures, when development is more rapid, the life cycle can be completed in six or seven weeks. Granulate cutworm’s life cycle is similar to that of the black cutworm (Hayslip et al. 1953).
Mole Crickets (Scapteriscus spp. and Neocurtilla hexadactyla)
Mole crickets are a problem in cabbage seedbeds, where they produce raised tunnels on the soil surface. Although they do not feed on the plant, their tunneling may cause cabbage seedlings to fall over (Workman 1983). Their enlarged front legs are adapted for burrowing, and they can also damage newly-transplanted cabbage plants by burrowing around them, resulting in drying of soil and roots. Mole crickets spend their entire life cycle in the soil and are nocturnal, becoming active at night and remaining in their tunnels during the day. Greatest damage occurs during warm, moist weather (Hayslip et al. 1953).
Silverleaf Whitefly (Bemisia argentifolii)
Silverleaf whitefly, previously known as strain B of the sweetpotato whitefly, is a common pest on cabbage in southern Florida. However, it does not severely damage the crop. Under heavy infestations, when the outer leaves become covered with whiteflies, the leaves are removed at harvest.
Adult females produce an average of 160 eggs each, depositing them on the lower surface of host plant leaves. The first nymphal (immature) stage, the crawler stage, attaches itself to the leaf near the empty egg case. The whitefly passes through three more sedentary nymphal stages, appearing like transparent scales, before molting to the adult stage. Whiteflies feed by sucking the plant’s sap through their needle-like, piercing-sucking mouthparts. Like aphids, they extract large amounts of the plant’s sap (phloem), excreting the excess liquid as honeydew, upon which sooty mold can grow (Johnson, Short, and Castner 2005; Norman et al. n.d.).
With a host range of more than 500 species of plants, the silverleaf whitefly has been observed to reproduce on at least 15 crops and 20 weed species in Florida. Whitefly populations commonly peak on the state’s crops at the time of harvest, as the whitefly migrates from crop to crop throughout the year. In southern Florida, populations build on fall vegetables and move directly to overlapping spring crops. In west-central Florida, whitefly adults trapped in cabbage fields are highest from November until April or May. Cabbage can serve as a winter reservoir for whiteflies colonizing spring plantings of tomatoes and other vegetable crops in southwest Florida. Over the summer fallow period, whitefly populations are low because whiteflies are limited to weeds, such as water primrose, hairy indigo, and spurge. Weeds are poor hosts to the whitefly and usually harbor many natural enemies that reduce populations during that time (Schuster, Polston, and Price 1992; Stansly 1995; Norman et al. n.d.)
Chemical Control
In 2010, Florida growers applied insecticides totaling 7,400 pounds of active ingredient to 100% of the state’s cabbage acreage. During the years when usage data was collected, 95%–100% of cabbage acreage has been treated with insecticides each year, with total annual usage ranging from 6,100 to 55,200 pounds of active ingredient. The most commonly used insecticides on Florida cabbage are Bacillus thuringiensis (B.t.) and spinosad. In fact, B.t. was the only insecticide with published use values for Florida cabbage in 2002. Older insecticidal materials used in Florida for cabbage were bifenthrin, carbaryl, chlorpyrifos, dimethoate, emamectin benzoate, endosulfan, esfenvalerate, imidacloprid, indoxacarb, lambda-cyhalothrin, malathion, methoxyfenozide, methomyl, permethrin, thiodicarb, tebufenozide, azadirachtin, azinphos-methyl, Beauveria bassiana, cypermethrin (beta or zeta), diazinon, disulfoton, fenpropathrin, insecticidal soaps, insecticidal oils, naled, oxydemeton-methyl, polyhedrosis viruses for corn earworm and beet armyworm, pymetrozine, pyrethrins, and sulfur.
Other insecticidal materials recently registered for use in Florida cabbage include thiamethoxam, esfenvalerate, acetamiprid, beta-cyfluthrin, clothianidin, flonicamid, pheromones, tebufenozide, chlorantraniliprole, buprofezin, gamma-cyhalothrin, potassium salts of fatty acids, pyriproxyfen, piperonyl butoxide, s-methoprene, spirotetramat, cyfuthrin, spiromesifen, sodium tetraborohydrate decahydrate, emamectin benzoate, spinetoram, novaluron, dinotefuran, kaolin, flubendiamide, cyromazine, flubendiamide, extract of Chenopodium ambrosioides, Paecilomyces sp., and Chromobacterium subtsugae (USDA/NASS 2010b; CDMS 2013).
Bacillus thuringiensis
The biopesticide Bacillus thuringiensis (B.t.) is the most important insect management tool for Florida cabbage growers. Cabbage growers use B.t. every year to manage diamondback moth and other lepidopteran larvae. A naturally occurring soil bacterium, B.t.produces spores and crystalline bodies that act as stomach poison to the insects that consume it. The most common formulations are highly specific for lepidopterous larvae (caterpillars) and do not harm beneficial organisms. However, it is most effective against smaller larvae. B.t. may be applied up to the day of harvest, meaning the pre-harvest interval (PHI) is 0 days, and the restricted-entry interval (REI) under the Worker Protection Standard is 4 hours.
Spinosad
Spinosad is a reduced risk broad-spectrum insecticide derived from fermentation of the naturally occurring soil bacteriumSaccharopolyspora spinosa. Since its registration, spinosad has become one of the most important insecticides for Florida cabbage growers (Eger and Lindenberg 1998). It controls many lepidopterous larvae, dipteran leafminers, and thrips. It is used by Florida cabbage growers primarily to manage diamondback moth and cabbage looper but also to control imported cabbageworm and armyworm. Spinosad has low activity against most beneficial insects and is useful in integrated pest management (IPM) programs. Spinosad may be applied up to 1 day before harvest (PHI=1 day), and the restricted entry interval (REI) under the Worker Protection Standard is 4 hours. For purposes of resistance management, a maximum of six diamondback moth treatments may be made per year (since this is a lower rate than the maximum of 0.16 lb ai/A). Additionally, no more than three consecutive treatments may be made in a 30-day period, followed by a 30-day period free of spinosad. The maximum amount that can be used per crop is 0.45 lb ai/A (CDMS 2013).
Use of Insecticides in IPM Programs
Action thresholds for caterpillar pests of cabbage were developed in Florida and Georgia during the 1970s and 1980s to reduce insecticide sprays on cabbage. Until then, growers had been spraying using a fixed schedule of once or twice weekly. Using insecticides based on early thresholds (the number of larvae present) was shown to greatly reduce insecticide sprays and still produce marketable cabbage. Thresholds were developed based on visual damage ratings and percent of plants with new damage, and these were as effective as the more time-consuming larvae counts. However, thresholds based on new damage tended to result in more insecticide applications. All thresholds were shown to be less effective when an atypical pest was present or a particular pest was present in very high numbers (Leibee et al. 1984; Leibee 1996).
An IPM program to manage diamondback moth in cabbage has been under development in Florida for several years. The program consists of multiple strategies, including using B.t. insecticides, biological control (parasitoid releases), trap crops, and pheromone treatments for mating disruption (Hu et al. 1998).
Use of Insecticides in Resistance Management Programs
Insecticide resistance in the control of cabbage insects in Florida has been a problem for more than 40 years. Diamondback moth is an agricultural pest that has demonstrated the ability to quickly become resistant to insecticides. From the 1940s through the 1970s, growers applied DDT, toxaphene, parathion, methoxychlor, mevinphos, endosulfan, naled, methomyl, and methamidophos to cabbage for control of multiple cabbage caterpillars. Cabbage looper resistance to DDT, parathion, and toxaphene was documented in 1957, and populations resistant to methomyl were found in the early 1980s. When the pyrethroids became available in the early 1980s, growers switched to permethrin and fenvalerate to control cabbage looper and diamondback moth, which became more difficult to control with the earlier insecticides. Permethrin and fenvalerate effectively controlled those pests until resistance began in the mid-1980s. Within a few years, growers were experiencing difficulty in controlling diamondback moth with those insecticides. In 1987, central Florida diamondback moth populations were found to be resistant to fenvalerate and methomyl but susceptible to chlorpyrifos, acephate, endosulfan, and thiodicarb. By the early 1990s, thiodicarb had also become less effective (Leibee and Savage 1992a; Leibee and Savage 1992b; Leibee and Capinera 1995; Leibee 1996).
Pyrethroids and carbamates became less effective during the late 1980s, forcing growers to modify their insecticide use. Growers switched to several organophosphates, endosulfan, and Bacillus thuringiensis kurstaki. None of these was completely effective, and resistance to B.t. kurstaki was confirmed in Florida in the early 1990s. Growers then switched to the newly introduced Bacillusthuringiensis aizawai-based insecticides, which offered greater diamondback moth control. In addition, reduced pyrethroids use resulted in greater natural control due to the return of parasites. Diamondback moth populations also appeared to develop greater susceptibility toB.t. kurstaki, which growers began using again. However, resistance has apparently been developing to B.t. aizawai (Leibee and Capinera 1995; Leibee 1996).
Resistance in diamondback moth populations in Florida has been largely attributed to the frequent use of single insecticides or classes of insecticides over time, as well as to the nearly continuous production of cabbage in isolated areas. During the 1980s, cabbage producers in Florida began to harvest later in the spring and transplant earlier in the summer, reducing the previous crucifer-free period that had existed between June and September. At the same time, container-grown transplants began to be produced during the summer months, giving diamondback moth continuous access to crucifers. The principal transplant producers have operated in the main field production areas, allowing easy movement of insect pests from fields to transplant houses at the end of the spring season and back to production fields at the start of the fall season. Both the development of insecticide resistance and the loss of natural enemies because of insecticide treatments for cabbage looper control contributed to the increase in diamondback moth populations in Florida during the 1980s. During this time, the diamondback moth became a major pest on cabbage. Intensive use of insecticides during the 1980s, combined with the increasing use of infested transplants and resulting greater insect pest problems, most likely contributed to the high degree of insecticide resistance found among cabbage pests in Florida (Leibee and Capinera 1995; Leibee 1996).
A number of recommendations were made to limit development of greater resistance problems in diamondback moth in Florida. Those included 1) avoiding cabbage production during the warmest months, when B.t.-based insecticides are least effective and insect problems are greatest; 2) destroying crop residues to avoid pest movement into new plantings; 3) using pest-free transplants; 4) inspecting the crop frequently, beginning at the seedling stage, and using action thresholds to minimize insecticide applications; 5) using pheromone traps to monitor adult activity and time insecticide applications; 6) using B.t. kurstaki and B.t. aizawai as the main insecticides to control diamondback moth, rotating the two to reduce resistance selection; and 7) avoiding the use of carbamates and eliminating the use of pyrethroids (Leibee and Capinera 1995).
Cultural Control
Mating disruption with sex pheromones was effective in reducing diamondback moth and cabbage looper populations in Florida cabbage. In field trials in northeast Florida, treating cabbage fields with sex pheromones controlled diamondback moth populations for most of the cabbage season, minimizing the need for pesticide sprays. In the trials, mating suppression was confirmed, but the effect of pheromone treatment on larval counts of cabbage looper was not evaluated because cabbage looper larvae were not present in sufficiently high numbers. Using pheromones is considered a promising management tactic if cabbage looper populations increase considerably (Mitchell et al. 1997a).
The use of trap crops has also been investigated to manage diamondback moth in Florida cabbage. A trap crop is a plant more attractive to the insect pest that lures it away from the more valuable crop. In preliminary studies, collard plants, when planted between rows of cabbage, were shown to have potential as a trap crop for diamondback moth. Collards also play an important role in maintaining populations of the natural enemy Diadegma insulare (Mitchell, Hu, and Okine 1997). Recently in northeast Florida, planting collards around the perimeters of cabbage fields helped reduce pesticide sprays for diamondback moth on cabbage by 75%–100% over cabbage fields treated with conventional insecticides, producing equivalent quantity and quality of cabbage (Weaver 1999).
Another cultural control for cabbage looper moths is the use of row covers, which can prevent cabbage looper moths from laying their eggs on the plants. However, using row covers is not always economically feasible (Capinera 2011b). Cutworm damage can be reduced by plowing under weeds at least one month prior to planting (Hayslip et al. 1953).
Biological Control
Florida cabbage growers depend on the microbial insecticide Bacillus thuringiensis to manage diamondback moth and cabbage looper, among other caterpillar pests. Several other natural control agents help reduce pest population densities in cabbage. A nuclear polyhedrosis virus (NPV) naturally reduces cabbage looper populations in some years. Larvae consuming the virus inclusion bodies usually die within 5–7 days, after becoming blotchy, then creamy white and swollen, and eventually limp. The virus is spread from the disintegrated bodies of infected larvae, a process aided by rainfall. Cabbage looper mortality from NPV tends to be greater during years of greater rainfall. The Trichoplusia ni NPV can be effective, but it has a narrow host range and consequently has not been commercialized (Capinera 2011b). Another nuclear polyhedrosis virus highly specific to beet armyworm is considered to be the most important natural mortality factor for beet armyworm larvae (Capinera 2011a). A polyhedrosis virus is commercially available for corn earworm as well.
Other natural mortality agents of cabbage looper include tachinid parasitoids such as Voria ruralis, which attacks medium or large larvae, wasp parasitoids such as Trichogramma spp., which parasitize looper eggs, and predators such as the earwig Labidura riparia, which has been observed feeding on cabbage looper larvae and pupae in Florida crucifer fields. Bird predation of cabbage looper has also been observed in fields in Florida. Mass release of Trichogramma spp. has been studied in several crops and was effective in crucifers, but it has not been used by Florida cabbage growers (Leibee 1996; Capinera 2011b).
The parasitic wasp Cotesia plutellae was imported and released in Florida in 1990, and since then it has been released in Florida cabbage fields sporadically, but it has not established. Inundative releases of C. plutellae have also been evaluated for control of diamondback moth populations in cabbage fields in northeast Florida. While the release rates used in the latest study (about 3,082 per hectare over the season) were not considered to provide sufficient economic control of the pest, such releases in combination with other IPM tactics (trap crops, pheromones for mating disruption, and use of B.t. insecticides) may prove effective. Parasitism by C. plutellae was complementary to the natural parasitism occurring from the native Diadegma insulare (Leibee 1996; Mitchell et al. 1997b; Mitchell et al. 1999).
Diamondback moth populations in Florida also suffer a high parasitism rate by Trichogramma sp. and are affected by several pathogens, including Zoophthora spp. (Leibee 1996). Additional natural enemies of cabbage pests reported in Florida include the parasites Meteorus vulgaris, a braconid wasp that attacks granulate cutworm and fall armyworm in the Everglades area, the encyrtid wasp Copidosoma truncatellum, a parasite of the cabbage looper, the ichneumonid wasp Horogenes insularis, which attacks diamondback moth and cabbage looper, the braconid wasp Diatretus rapae, which parasitizes cabbage and turnip aphids, and the tachinid flies Archytas piliventris and Eucelatoria rubentis, which attack armyworms and cutworms.
Predators of cabbage insects reported in Florida include the pentatomid bugs Podisus maculiventris and Podisus mucronatus, which feed on cabbage loopers and imported cabbageworms, and the reduviid bug Zelus bilobus, which also attacks cabbage loopers and imported cabbageworms. Other predaceous bugs include the pentatomids Stiretrus anchorago and Euthyrhynchus floridanus, and the reduviidsArilus cristatus and Sinea diadema. In addition, ground beetles of the genus Calosoma have played a role in biological control of cabbage pests. The native species C. scrutator and C. sayi frequently feed on cutworms and armyworms. Finally, the ladybird beetles Cyclonedasanguinea immaculata, Hippodamia convergens, Ceratomegilla fuscilabris floridanus, Scymnus collaris, Scymnus terminatus,Exochomus marginipennis, Psyllobora sp., and Coccinella novemnotata are all aphid feeders found in Florida (Hayslip et al. 1953).
Disease Management
Disease Pathogens
Diseases are less of a problem than insect pests for Florida cabbage growers. Most of the diseases affecting cabbage are sporadic, but cabbage growers must contend with the presence of at least one major disease in most years. The most significant diseases on cabbage in Florida are black rot (caused by Xanthomonas campestris), sclerotinose (caused by Sclerotinia sclerotiorum), downy mildew (caused by Peronospora parasitica), and Alternaria leaf spot (caused by Alternaria spp.). Other diseases that occasionally affect cabbage in Florida include bacterial leaf spot (caused by Pseudomonas cichorii), damping-off (caused by Fusarium and Pythium spp.), turnip mosaic (caused by turnip mosaic virus), wirestem (caused by Rhizoctonia solani), and yellows (caused by Fusarium oxysporum f.conglutinans). White rust of foliage (caused by Albugo candida) and powdery mildew (caused by Erisyphe polygoni) are minor diseases on cabbage in Florida. Another condition characterized by tiny black specks on the foliage the week after harvest has an unknown origin. Additional minor problems include brown heart, caused by boron deficiency, and tipburn, caused by potassium deficiency (Momol, Raid, and Kucharek 2005).
Black Rot (caused by Xanthomonas campestris pv campestris)
Black rot is also called black spot or black leg. It is the most serious disease of cabbage in Florida and is most common when cabbage transplants are grown outdoors. It can occur at any time of the year, and it is difficult to control once it gets into a field. Losses of up to 10% are recorded annually. In addition to cabbage, black rot attacks other cole crops, including broccoli, cauliflower, kale, kohlrabi, Brussels sprouts, rutabaga, turnip, collards, radish, mustard, and water cress. It also attacks cruciferous weeds such as wild radish, pepper grass, and shepherds purse (Kucharek and Strandberg 2000a).
The bacteria that cause black rot enter cabbage through injuries or natural openings on the leaves. Mechanical injury during transplanting, particularly injury that results in wounds on the root system, is an ideal entry mechanism for the pathogen. Cracks in the tissue of older roots also offer an entry point, particularly when the soil is saturated with water. Injury from insect feeding is a minor source of entry. A more virulent strain of the bacteria, which is present in Florida, is more likely to enter plants through stomatal openings in the leaves (Kucharek and Strandberg 2000).
Symptoms may not appear on leaves until up to 43 days after infection. Early symptoms of black rot include stunting, yellowing leaves, and blackening veins. A yellow, wedge-shaped area may be produced at the ends of leaves. As the bacteria move down the leaf veins and into the plant’s vascular system, the disease becomes systemic, as bacteria move within the vascular system to healthy leaves. Both leaf veins and vascular tissue become darkened, and leaves wilt and die. Plants are later dwarfed and produce one-sided heads (Kucharek and Strandberg 2000).
Plants at any growth stage can be infected by black rot. The disease is seed-borne, and plants from infected seeds die quickly after germination. When young seedlings are infected, the plants do not produce heads, and heads from plants infected later will deteriorate after harvest. In addition to being spread on seeds, the bacteria are spread by rain, irrigation, and any water movement in the field. The disease is most severe under warm and wet conditions. The bacteria can survive in undecomposed debris of crucifer plants, and it grows at temperatures ranging from 40°F to 97°F (4°C–36°C), although optimum temperature for its growth is between 80°F and 86°F (27°C–30°C). After plants are infected, symptom expression is greatest at temperatures between 68°F and 82°F (20°C–28°C) (Kucharek and Strandberg 2000).
Sclerotinose (caused by Sclerotinia sclerotiorum)
Sclerotinose, also called Sclerotinia watery rot or watery soft rot, is another important cabbage disease in Florida, but it does not occur each year. Its sporadic occurrence coincides with the simultaneous presence of cool and damp conditions. Optimum conditions for rapid disease development include temperatures of 60°F–70°F (15°C–21°C) and high humidity with dew formation (Pernezny and Purdy 2009).
The disease is first seen on leaves and stems close to the ground. Small, water-soaked spots appear and enlarge, accompanied by a growth of white mycelium (the body of the fungus). As the disease develops, the fungus grows upward on the plant, often spreading over the head and creating a soft, dark water-soaked mass on the leaves. Within this mass, many small, black sclerotia (resting structures) are produced. The sclerotia are characteristic of diseases caused by S. sclerotiorum, and these are the fungus’ survival mechanism from season to season. Sclerotia may form on the surface of the head as the disease progresses and the fungus invades the whole plant. Plants with heavily infected stems will wilt, fall over, and eventually die. Sclerotinose often follows cold conditions or other types of plant injury (Momol, Raid, and Kucharek 2005).
Sclerotinia watery rot was a major disease in the Hastings area in the 1940s and 1950s, but declined in importance during the 1960s and 1970s. At that time, the disease was observed after hard freezes that damaged the cabbage plants at the soil line, where the fungus would enter the injured stem. Sclerotia were the source of inoculum for such infections. After falling to the ground and being incorporated into the soil during disking and bedding operations, the sclerotia produce fruiting bodies (apothecia) after at least 10 days of high soil moisture. The fruiting bodies produce spores that are carried by air currents to infect injured or dying cabbage tissue. From there, the infection can spread slowly by plant to plant contact. The fungus can also attack the plant from the soil without forming fruiting bodies and resulting spores. In that case, leaves in direct contact with the soil are infected, or infection occurs at the soil line. Infection resulting from spore dispersal can occur on all plant parts (Weingartner 1981).
Downy Mildew (caused by Peronospora parasitica)
Downy mildew occurs on cabbage in Florida in most years; however, losses are minimal (approximately 2%) because growers effectively control the disease with chemical management. Fungicides are the principal means of downy mildew control by Florida cabbage growers, and the use of non-systemic fungicides is recommended. Strains of the fungus resistant to systemic fungicides develop more rapidly (Kucharek 2000a).
The fungus that causes downy mildew of cabbage also attacks other crucifers, such as cauliflower, collards, Chinese cabbage, Brussels sprouts, broccoli, kale, and kohlrabi (McRitchie 1973). The first symptoms of downy mildew are black specks and yellow-brown spots forming on the upper leaf surface, accompanied by a fluffy mold growth developing on the lower surface. Young leaves may fall off when infected, and on older leaves, the spots may coalesce, producing large, sunken, tan spots. The disease can attack young seedlings or plants that have already headed. On older cabbage plants, the disease produces dark, sunken spots, which may appear purplish, on the head or wrapper leaves. Downy mildew infection of older plants may leave them susceptible to sclerotinose-causing bacteria (Kucharek 2000a).
Disease development can be extremely rapid, affecting an entire field in 3–4 days under favorable conditions (cool, moist weather). Although spore production can occur at temperatures of 39°F–85°F (4°C–29°C), optimum temperatures are 53°F–61°F (12°C–16°C). Germination and penetration of the spores are most rapid at temperatures of 42°F– 61°F (6°C–16°C) and can occur at any temperature from 39°F to 75°F (4°C–24°C). At temperatures around 75°F (24°C), symptoms occur in 3–4 days of infection. When temperatures are suitable, disease development progresses more rapidly under wetter conditions (Kucharek 2000a).
The fungus that causes downy mildew can be spread by infected transplants or windblown spores produced in the lesions on the lower leaf surface of infected plants. Another type of spore, functioning as a survival spore, is produced within infected plant tissue during crop senescence and may serve as a source of inoculum for later crops. However, the role of the survival spores in disease spread is considered minimal. The extent to which weeds serve as a source of inoculum is presently unclear (Kucharek 2000a).
Alternaria Leaf Spot (caused by Alternaria spp.)
Alternaria leaf spot can also be a major disease on Florida cabbage during the years it occurs. It tends to be more of a problem in northern Florida than in southern Florida. The fungus can be seedborne, being carried in the seed in the form of mycelium or on the outside of the seed in the form of spores. Alternaria spp. seedling infections are not common in Florida. However, when they occur, infected seeds can experience preemergent or postemergent seedling blight. If they don’t kill the plant outright, stem lesions at the seedling stage will result in inferior produce size (Kucharek 2000b).
Leaf spots produced by Alternaria spp. are much more common in Florida. Initial symptoms of Alternaria leaf spot include small, dark spots on the leaves. A target spot can be seen as spots enlarge. Older spots enlarge to 2–3 inches (5–7.5 cm) and may be black, brown, or tan. A yellow halo around brown lesions on the leaf edges can be used to distinguish Alternaria leaf blight from black rot. However, yellow halos are not always produced around leaf lesions. As disease development continues, leaves may yellow and die. Concentric bands or a solid mass of fuzzy dark green to black growth within leaf spots develops as a result of spore production. Spores are usually produced at night and released during the day. But during prolonged periods of overcast weather, spores may be produced continuously. Optimum temperatures for spore production are 75°F–82°F (24°C–28°C), and new spores can be produced in 7–10 days of infection under favorable temperature conditions for penetration and germination (Kucharek 2000b).
In addition to being produced on leaf spots, spores can be produced on crop debris. The fungus may survive on cruciferous weeds as well. Infections and lesion size increase substantially with longer periods of leaf wetting, as a result of prolonged dew periods or frequent rains (Kucharek 2000b).
Bacterial Leaf Spot (caused by Pseudomonas cichorii)
Bacterial leaf spot is only occasionally a problem on cabbage in Florida. The disease produces small, slightly sunken spots that are gray to dark brown and that may appear as target spots (concentric rings). Bacterial leaf spot occurs principally on the wrapper leaves, but it may affect internal leaves when conditions are favorable. Overhead irrigation and poor field drainage favor disease development (Momol, Raid, and Kucharek 2005).
Damping-Off (caused by Fusarium and Pythium spp.)
Damping-off occurs in transplant seedbeds. Plants either fail to emerge, or in the case of postemergence damping-off, a water-soaked lesion develops on the stem at or just below the soil surface. The seedling later wilts, falls over, and then dies (Momol, Raid, and Kucharek 2005).
Turnip Mosaic (caused by turnip mosaic virus)
Turnip mosaic, also called black ringspot, occasionally affects cabbage in Florida. The disease is transmitted by aphids, and it also affects other crucifers, beets, spinach, and tobacco, among others. Symptoms appear on infected plants when the temperature is 75°F–85°F (23.9°C–29.4°C). Leaves become mottled, and plants appear stunted. Mosaic symptoms develop first on the underside of leaves. As the tissue in the dark green spots dies, a ring spot pattern develops. Heads from plants that did not appear to be infected in the field may develop symptoms in postharvest storage (Momol, Raid, and Kucharek 2005).
Wirestem (caused by Rhizoctonia solani)
Wirestem is an occasional problem for Florida cabbage growers. The causal fungus can attack roots, stems, and leaves. In some cases, a seedling’s outer stem will shrivel, turn dark, and become tough. Under appropriate weather conditions, such seedlings can recover. However, if the fungus continues to grow up the stem, bottom rot and head rot may develop (Momol, Raid, and Kucharek 2005).
Yellows (caused by Fusarium oxysporum f. conglutinans)
Yellows is another occasional disease seen by Florida cabbage growers. The disease first occurs in the lower leaves, with the appearance of a yellow-green color. Yellowing of the tissue may move upward to the top leaves. Yellowed tissue turns brown, and premature shedding of the leaves occurs. Vascular tissue in the leaves and stems is blackened. Sometimes, only one side of the plant is infected, in which case the plant curls and bends (Momol, Raid, and Kucharek 2005).
Chemical Control
In 2010, Florida growers applied fungicides totaling 70,600 pounds of active ingredient to 99% of the state’s cabbage acreage. The most commonly used fungicide on Florida cabbage was chlorothalonil with total usage of 65,600 pounds. Older fungicidal materials registered for use in Florida cabbage are copper compounds, azoxystrobin, Bacillus sp., fosetyl-Al, hydrogen dioxide, mefenoxam, PCNB, and sulfur (USDA/NASS 2010b; CDMS 2013). Other recently registered fungicidal materials are acibenzolar-s-methyl, metalaxyl, potassium phosphite, potassium bicarbonate, potassium phosphate, potassium silicate, pyraclostrobin, mancozeb, boscalid, penthiopyrad, dimethomorph, chloropicin, 1,3-dichloropropene, cyprodinil, fludioxonil, fluazinam, fluopicolide, triflumizole, cyazofamid, fenamidone, mandipropamid, metalaxyl, thiram, laminarin, Streptomyces lydicus, Glicoladium virens, Trichoderma sp., and oils (neem, clove, rosemary, and thyme) (CDMS 2013).
Chlorothalonil
Chlorothalonil is a broad-spectrum chloronitrile fungicide used to manage Alternaria leaf spot and downy mildew (CDMS 2013). Chlorothalonil may be applied up to 7 days before harvest (PHI=7 days), and the restricted entry interval (REI) under the Worker Protection Standard is 12 hours. The minimum retreatment period is seven days, and the seasonal maximum application is 12 lb ai/A.
In 2010, Florida growers applied an average of 0.9 pounds of active ingredient per acre at each application to 99% of cabbage acreage, an average of 6.7 times. Total usage was 65,500 pounds of active ingredient.
During the years usage data was collected, Florida cabbage growers applied chlorothalonil at an average rate of 0.80–1.42 pounds of active ingredient per acre at each application to 39%–94% of cabbage acreage. Growers have made an average number of applications ranging from 4.6 to 5.7 each year, totaling 22,700–56,300 pounds of active ingredient annually (USDA/NASS 2012).
Copper Hydroxide
Florida cabbage growers use copper hydroxide to manage black rot, Alternaria leaf spot, and downy mildew. The restricted entry interval (REI) of copper hydroxide under the Worker Protection Standard is 24 hours (CDMS 2013).
Cultural Control
Black rot can be spread by infected seeds or during transplant production. Although resistant varieties are available, the most effective management is adequate sanitation while producing transplants. Cabbage growers can buy certified transplants as well as seeds that have been certified as being free of black rot. Regardless of the source of cabbage seeds, using the hot water treatment is an essential management tactic that effectively controls black rot; the hot water treatment is soaking the seeds at 122°F (50°C) for 30–35 minutes.
Specific practices during transplant production include the use of clean flats and disease-free seed, destruction of diseased plants and residue after the harvest, avoiding the movement of contaminated soil, fumigation, or rotation of transplant beds, irrigation from a well and not from an open ditch, elimination of cruciferous weeds, raising transplant beds for adequate drainage, avoiding handling of plants while wet, and avoiding placing transplant beds within one-quarter mile of crucifer production fields. In addition, transplants should not be wetted down before transplanting, since black rot spreads most easily under moist conditions (Momol, Raid, and Kucharek 2005).
While the use of clean planting material is the key cultural control for black rot, most growers also stay out of the fields as much as possible during wet conditions to avoid spreading the disease. Additional cultural practices that should be followed together include planting in fields that have not been in crucifer production for 12 months, plowing down crucifer fields just after harvest, and thoroughly cleaning all equipment and tools prior to use in the field or transplant bed (Kucharek and Strandberg 2000; Momol, Raid, and Kucharek 2005).
The best cultural control for sclerotinose, as well as most other diseases, is wider plant and row spacing, which cut down on long dew periods. However, to maximize yield, most growers do not increase spacing as a disease management practice. Additional recommendations for cultural control of sclerotinose include rotating with a crop that is not susceptible to Sclerotinia (e.g., sweet corn), turning the soil at least six inches when plowing, avoiding the use of overhead irrigation, and completely flooding or intermittently flooding for six weeks during the summer, if possible (Momol, Raid, and Kucharek 2005).
Destruction or burial of old plant beds and residues in harvested fields aids in managing Alternaria leaf spot and downy mildew. Elimination of cruciferous crops and weeds around seedbeds, rotation with non-cruciferous crops, and the use of disease-free seeds and transplants also aid in managing those diseases. Turnip mosaic can be managed by eliminating the weed hosts of the virus, particularly mustard-type weeds, both in the seedbed and field. Early control of aphids, especially in seedbeds, is the other key to reducing virus incidence. Cultural controls for wirestem include rotating crops on both seedbeds and fields, providing adequate drainage, cultivating as soon as possible after heavy rains to aerate and dry the soil, and avoiding planting in crop debris or a green manure crop that has recently been incorporated. For yellows, disease-free transplants should be used, and the only available control once soil has been infested is using resistant varieties (Kucharek 2000b; Momol, Raid, and Kucharek 2005).
Postharvest Decays and Their Management
Excessive trimming of wrapper leaves may cause the more susceptible inner leaves to wilt. In addition to contributing to wilting, cuts or breaks can provide an entry site for disease pathogens. The postharvest decays watery soft rot, bacterial soft rot, gray mold rot, Alternaria leaf spot, and black leaf speck can all affect cabbage during the postharvest period (Sargent et al. 2007).
Nematode Management
Nematode Pests
Plant-parasitic nematodes are microscopic roundworms found in soils. They primarily attack plant roots. General symptoms of nematode damage include stunting, premature wilting, leaf yellowing, and related symptoms characteristic of nutrient deficiencies. Stunting and poor stand developments occur in patches throughout the field as a result of the irregular distribution of nematodes within the soil. Root-knot, sting, stubby root, and awl nematodes are all important pests of crucifers, and cyst nematode is a serious pest in central Florida (Noling 2009; Noling 2012).
Root-Knot Nematodes (Meloidogyne spp.)
Many cruciferous plants are hosts to common species of root-knot nematodes. In greenhouse tests, moderate levels of root galling and egg masses occurred on cabbage plants infested with three species of root-knot nematodes (M. javanica, M. incognita races 1 and 3, andM. arenaria) (McSorley and Fredrick 1995).
Root-knot nematodes enter the host plant root as second stage juveniles and settle within the root to establish a feeding site. At the feeding site, nematode secretions cause the surrounding plant cells to enlarge and multiply, producing the characteristic galls associated with root-knot attack. As more nematodes enter the root and feeding continues, the galls fuse to form large tumors on the roots. Within the root, the developing female molts several times before developing into a swollen, pear-shaped adult. The adult may live in the host plant for several months, laying hundreds to several thousand eggs that are released into the soil. Low temperatures or very dry soil conditions can cause eggs to hatch more slowly. Root deformation and injury caused by root-knot nematodes reduce root area and interfere with water and nutrient uptake. Resulting symptoms include stunting, wilting, chlorosis, and yield loss. In addition to expending the plant’s resources, the gall tissue is more susceptible to secondary infections such as root rots (Stokes 1972; Crow and Dunn 2005; Noling 2012).
Sting Nematodes (Belonalaimus longicaudatus)
Cabbage in Florida often experiences severe yield losses because of sting nematodes (White and Rhoades 1992). Sting nematodes are ectoparasites, remaining outside the plant root and feeding superficially at or near the root tip. Affected root tips turn yellow and later necrotic, with cavities forming and the root tip swelling slightly. Damage from sting nematode feeding inhibits root elongation and causes roots to form tight mats and appear swollen, resulting in a "stubby root" or "coarse root" appearance (Christie 1959; Crow and Dunn 2005; Esser 1976; Noling 2012).
Sting nematodes are especially damaging to seedlings and transplants. Death of transplants on highly infested sites can leave gaps of missing plants or patches of plants lacking vigor. In northern Florida, sting nematodes are most abundant in April and May. Sting nematodes prefer sandy soils (with 84%–94% sand) and are most abundant in the upper 12 inches (30 cm). Optimum soil temperature for this nematode is 77°F–90°F (25°C–32°C), and optimum soil moisture is about 7% (Christie 1959; Crow and Dunn 2005; Esser 1976; Noling 2012).
Stubby-Root Nematodes (Trichodorus spp., Paratrichodorus spp.)
Stubby-root nematodes feed externally on the root surface and remain in the soil throughout their life cycle. They mainly feed at the tip of the growing root, stopping root elongation. The result can be a short, stubby root system with swollen root branches. Their feeding may also cause abnormal growth of lateral roots and increased production of branch roots. In Florida, stubby-root nematodes are found mainly in sandy or sandy loam soils, but also occur in muck soils. In addition to cabbage, the principal crops injured by this nematode in Florida include beets, corn, celery, cauliflower, chayote, and several grasses. Stubby-root nematode populations build up quickly in the presence of a suitable host and likewise decrease quickly when a host is no longer available (Christie 1959; Crow and Dunn 2005; MacGowan 1983).
Awl Nematodes (Dolichodorus spp.)
Awl nematodes also feed superficially from the outside of the plant root, inhibiting root elongation. Awl nematodes are similar to sting nematodes in appearance, habits, and the plant injury symptoms as a result of their feeding (Christie 1959; Crow and Dunn 2005).
Chemical Control
Ethoprop, fenamiphos, metam, and 1,3-dichloropropene are the nematicides registered for use on cabbage in Florida. Methyl bromide has been phased out with no critical use exemption for use in cabbage (CDMS 2013).
Cultural Control
The use of pest-free transplants is the most important cultural control for nematodes on cabbage. Transplants should be produced in a sterile growing medium or fumigated soil (Noling 2009).
Some resistance to sting nematodes has been found in several cabbage varieties tested in Florida. However, using nematode resistance alone has not been found to be economically feasible (White and Rhoades 1992).
Weed Management
Weed Pests
Weeds can reduce cabbage yields by competing for nutrients, water, and light. Managing weeds early in the season is particularly important to maintain crop vigor and yield. Since cabbage is largely a winter crop, many weed species are winter annuals. Wild radish is a major weed in Florida cabbage fields. Cutleaf evening primrose and Carolina geranium may also be present. In the early spring or late fall, there may be problems with summer annuals like amaranth, lambsquarters, and Pennsylvania smartweed, among others (Dittmar and Stall 2010).
Wild Radish (Raphanus raphanistrum)
Wild radish is a winter annual that dies when hot weather begins, although some plants can live for a whole year in Florida. It can reach 1 meter (3.28 feet) in height. The stems have prickly hairs when the plant is younger and become smooth as the plant matures. The plant has one to several branches, and the leaves are hairy, with deep, rounded lobes. Wild radish germinates rapidly. The plant is often confused with wild mustard (Brassica kaber), but wild mustard is not found as a weed in Florida crops (Hall, Vandiver, and Ferrell 2012a).
Wild radish can have different effects on cabbage yield depending on when it occurs in the season. For example, during the warmer part of the spring and fall, the presence of up to 16 wild radish plants per meter (3.28 feet) of cabbage row will not reduce cabbage yields. However, during the cooler part of the season, only one wild radish plant per meter will significantly reduce cabbage yields (Dittmar and Stall 2010). In general, wild radish is a poor competitor with cabbage during the spring planting season (Steed et al. 1998).
Cutleaf Evening Primrose (Oenothera laciniata)
Cutleaf evening primrose is an annual plant with hairy stems that branch at the base. Its long, narrow leaves are deeply cut near the base (Miller et al. 1975).
Carolina Geranium (Geranium carolinianum)
Carolina geranium is a winter annual with smooth, reddish stems that branch widely at the base (Miller et al. 1975). Its many branches form a circular growth from the plant’s center, and it may rise to 0.6 m (2 feet) tall from a tap root (Hall, Vandiver, and Ferrell 2012b).
Amaranth (Amaranthus spp.)
Amaranths (pigweeds) are summer annual broadleaf weeds with erect stems that grow to 2 meters (6.5 feet) tall. Several species of amaranth are present in Florida; the most important are smooth pigweed (Amaranthus hybridus), spiny amaranth (A. spinosus), and livid amaranth (A. lividus). Amaranths or pigweeds reproduce solely by seed, producing very small, dark seeds. Smooth pigweeds flower from July to November, and spiny amaranth flowers from June to October. They prefer open areas with bright sunlight (Lorenzi and Jeffery 1987).
Lambsquarters (Chenopodium album)
Lambsquarters is an erect, summer annual that grows to 2 meters (6.5 feet) tall. It grows well on all soil types and over a range of soil pH values (Hall, Vandiver, and Ferrell 2012c).
Pennsylvania Smartweed (Polygonum pennsylvanicum)
Pennsylvania smartweed is a summer annual with smooth, branching stems and smooth, pointed leaves. It grows from 1 to 4 feet (0.3–1.2 meters) tall (Miller et al. 1975).
Chemical Control
In 2006, Florida growers applied herbicides totaling 4,100 pounds of active ingredient to 52% of the state’s cabbage acreage. During the years when usage data was collected, 52%–89% of Florida’s cabbage acreage was treated with herbicides each year, with total annual usage ranging from 4,100 to 13,000 pounds of active ingredient (USDA/NASS 2010b). Historically, the most commonly reported herbicides used on cabbage in Florida are trifluralin, metolachlor, and napropamide, but DCPA, glyphosate, oxyfluorfen, sethoxydim, paraquat, bensulide, pelargonic acid, and clethodim are also available for use on cabbage (USDA/NASS 2012). Other active ingredients recently registered for weed management in cabbage include carfentrazone-ethyl, clomazone, pyraflufen-ethyl, oxyfluorfen, d-limonene, sethoxydim, pendimethalin, clopyralid, ammoniated soap of fatty acids and oils (cinnamon and clove) (CDMS 2013).
Cultural Control
Cultural weed management practices include crop rotation, cover cropping, high-density planting, mulching, cultivation, and flooding. Cabbage growers in northern Florida tend to cultivate more than those in southern Florida, who use herbicides to a greater extent (Dittmar and Stall 2010). INTEGRATED PEST MANAGEMENT FOR MANGO
Integrated pest management programs for mango must be based on sampling and on economic thresholds, and must take into account the effects of cultural practices, horticultural sprays and disease control on pest and natural enemy interactions. An analysis of the mass of information available on the different mango pests,viz., fruit flies(Bactrocera sp.,Ceratitis sp.,Anastrepha sp.), mango seed weevil(Sternochetus mangiferae), thrips(Frankliniella spp.), gall midges(Procontariniasp.), scales, mites and mealybugs is given, as well as different examples for future entomological research.
Integrated Pest Management
Mangoes are prone to insect infestation and disease infection at any stage of their development. Without proper pest management program, quality fruits may not be produced.

The current control measures for pests’ attacking mango still relies on the use of pesticides. Most insecticides and fungicides are applied as calendar spray in an excessive manner resulting to pest resistance, elevation of minor pests to major ones, destruction of natural enemies and contamination of environment. In addition, pesticides are expensive and have caused in increased production inputs.

Many of these problems can be minimized though Integrated Pest Management (IPM). This involves these of alternative measures in combination, to minimize pests.

The IPM strategies make use of cultural management (pruning, cultivation, sanitation, proper nutrition to enhance vigor and fruit bagging) conservation of beneficial insects (pollinators and bio-con agents) and proper pesticide management.

This brochure on IPM for mango production emphasizes prevention of pests through destruction of source and prevention of its spread.

INSECTS

Circular-white back borer
Common names: Circular-white back borer,

Leaf cutting beetle

Scientific name: Callimetropus sp

Parts Affected: Young twigs and terminal leaves

Destructive stages: Adults and larvae

Description: This is a long-horned beetle and has very similar habits to the twig borer/cutter. The adult scrapes the bark of young twigs causing the death of terminal parts. The insect is easily identified by the circular white mark on the back of its body. Aside from destroyingthe twigs, the insect also cut the leaves of mango especially those on the tips of the shoots, hence, the name of “leaf cutting beetle”. Pile of cut leaves on the ground is a common indication of the presence of the insects.

Prevention/control

Like the twig borer, adults of the circular white-back borer are attracted to young leaves of mango or flushes for egg laying. Insecticides recommended for a twig borer infestation can also be used to protect trees from circular white back infestation. To prevent or minimize damage, spray the whole canopy with Karate at 1 ½ tbsp per 16 L water. Repeat application after one month, especially during flushing. Other pyrithroids can be used.

Green beetle
Scientific name: Anomala sp.

Common names: Green beetle and Toy beetle

Alternate hosts: Santol, avocado, coconut, cashew etc.

Destructive stage: Adults

Parts affected: Mainly leaves and sometimes flowers and fruits

Description: Adults are metallic green. They feed mainly on the leaves and occasionally on the flowers of mango. these insects also attack young fruits by chewing bits and pieces of the peel or skin, particularly near the fruit stalk.

Prevention/control * Remove the adults from the tree by shaking the branches and spray insecticide. * Adults are attracted to light; hence, light trapping is an effective control measure. * Avoid piling compost or other organic matter near mango trees since these are preferred sites for egg-laying.

Mango cecid fly
Scientific name: Procantarinia sp.

Common names: Leaf gall midge, Gall fly and Mango leaf gall

Parts affected: Leaves and fruits

Destructive stage: larvae, adult

Description: Adults which are mosquito-like in appearance prefer to lay eggs on new flushes (young leaves). The larvae, which develop from eggs, mine the leaves producing galls or swelling tissues. Under heavy infestations, the leaves wrinkle and remain yellow. Close examination of the leaves shows dark green, circular galls randomly distributed on the leaves blade. When open, each gall contains yellow larvae of the cecid fly. When the adults emerged from this galls, the leaves produce circular spots or holes which are sometimes mistaken as fungal infections (anthrac nose). The latter, is however, irregular in shape.

While the damage of cecid fly is usually associated with galling of young leaves, infested fruits produce circular, brown to black, scab-like spots randomly distributed on the fruit surface. This damage is commonly called “buti”, armalite, and kurikong ’and‘saksak walis’ by growers the water –soaked spots with contains small, yellow larvae. Infested fruits retain the scabby lesions at harvest affecting their quality.

Prevention/control * Orchard sanitation is important. Clear weedy areas since adults prefer to stay on these plants. * Young leaves are very attractive for egg laying. Spraying Sevin, Decis, Karate, and Stingray (3 to 4 tbsp per 16 L of water) will minimize damage. Spray insecticides in the afternoon, preferably 5:00 to 6:00 pm. * Prune crowded branches (particularly irregular branches) to allow light penetration. * Bag the fruits 55 to 60 days after flower induction.

Mango thrips
Scientific name: Scitothrips dorsalis (Giard)

Selenothrips rubocinctus (Giard)

Common names: Mango thrips, Red-banded thrips

Destructive stage: Nymphs and adults

Parts affected: leaves and flowers

Description: Mango thrips are small insects with “fringe” wings. These are occasional insect pests of mango but maybe destructive in some areas. Adults and nymphs destroy the leaves by scrapping the surface and feeding on the plant sap. Affected leaves develop brown areas, and later dry up and fall to the ground. Burning effect on flowers is a common damage of thrip injury.

Prevention/control

Both young and adults insects are sensitive to light. Prune crowded branches to allow light penetration, which create an environment less favorable for their development. Many insecticides are effective in the control of thrips, provided these are sprayed in fine mist and sprayed underneath the leaves where they hide.

Mealybugs
Scientific name: Ferrisia virgata (Cockerell)

Planococcus lilacinus (Cockerell)

Common names: Grey mealy bug and Cottony cushion mealybug

Alternate hosts: Many ornamental plants and fruit trees.

Destructive stage: Nymphs and adults

Parts affected: leaves, flowers and fruits

Description: Mealybugs (white, cottony insects) feed on leaves, especially on the flushes by sucking the plant sap. Affected parts turn yellow, dry up, and eventually fall. Mealybugs also excrete a sticky fluid known as “honey dew” where the sooty molds grow. The latter covers the leaf area producing black papery film on the surface. Sooty molds affect the photosynthetic activity of the leaves.

Both adults and nymphs attack the flowers by feeding on the base, gradually moving up to cover the entire flower. The flowers dry up and drop off prematurely. The “honey dew” produced by mealy bugs attracts red ants and serves as medium for the growth of sooty molds.

Both adults and immature insects congregate on the stalk before moving to the lower portions of the fruit. Excessive feeding of the sap damages the stalk and result to fruit drop. On the fruit surface, the insect secretes “honey dew” from which sooty molds develop. This spoils the appearance of the fruit. At harvest, mealy bugs can persist on the fruit affecting quality. It can attract ants that are cumbersome during harvest.

Prevention/control * Pruning creates an environment that is not favorable for the growth of mealy bugs. * Like scale insects, mealy bugs have symbiotic relationship with red ants. Mealy bugs and scale insects provide food for the red ants through their excreta (honey dew). In return, ants offer protection and distribute to insect to the different part of the tree. Spray the red ants with Malathion (1 ½ to 3 tbsp per 16 L water), decis (1 to 5 tbsp per 16 L water), and karate (3/4 to 1 ½ tbsp per 16 L water) to prevent the spread of mealy bugs. * Bag the fruits at 55 to 60 days after induction to prevent damage from mealy bugs. The paper bag should be closed properly at all sides and should remain intact up to harvest.

June beetle
Scientific name: Leucopholis irrorata (chevrolat)

Common names: toy beetle, June beetle

Alternate hosts: Santol, avocado, Cashew, Coconut, and many other fruit trees

Destructive stage: Larva and adult

Parts affected: Roots, leaves and sometimes flowers

Description: The adults are brown ans are easily dropped down when foliage are shaken. They feed extensively on the leaves, leaving only the midrib. The larvae (grubs) feed on the roots. The insect gets into the root system from infested organic matter, affected plant wilts and if uprooted, small, curved larvae can be seen feeding on the roots.

Prevention/control * Before applying the organic matter as fertilizer for mango, dry it thoroughly to kill the eggs of the beetle. * Granular insecticide like Furadon can be applied in the soil to kill the larvae. * Adults can be controlled by spray application of insecticide.

Scale insect
Important species recorded on mangoes:

Oriental scale: Aonidiella orientalis (Newstead)

Tropical scale: Hemiberlesia palmae (Cockerell)

Alternate hosts: Several fruit trees

Destructive stage: Adults, nymphs (crawlers)

Parts affected: Almost all parts of mango particularly twigs, branches, leaves and fruits

Description: Scale insects are usually circular in form with scale-like appearance. In the nursery, leaves of grafted mangoes are readily infested with scale insects, causing them to dry and fall. On bearing trees, high population of scale insects cause blackening of the canopy due to growth of fungus “sooty mold” which develops from their excreta (honey dew). Affected leaves are covered with a thin, black, papery film which produces unsightly appearance. Moreover, the photosynthetic activity (food production) of the leaves is reduced considerably. The branches and twigs of mango are susceptible to attack of scale insects. While feeding, they inject toxic substances into the tissues which result in the production of galls (bulging of twigs) and distortions of affected parts. Damaged portions fail to heal or recover.

Prevention/control * Young scale insects are carried and distributed by red ants to different parts of the tree. To prevent spread, ants should be destroyed by spray application (Please see Annex for suggested control measures.) * Prune regularly to remove unhealthy and crowded branches. * Bag the fruits at 55 to 60 days after induction to prevent damage from scale insects. Seal the paper bag properly at all sides and let it remain intact up to harvest. * Use clean planting materials free from any infestation. * Trees which are sickly and crowded are susceptible to scale insect attack. * Under severe attack, prune affected parts, spray insecticide with sticker, fertilize and irrigate.

Termite
Scientific name: Macrotermes gilvus (Hagen)

Common names: White ants, termite

Alternate hosts: Several fruit trees

Destructive stage: Adults and immature

Parts affected: Roots, trunks and branches

Description: Similar to ants but have soft bodies and are whitish in color. Termites construct earthen tunnels visible near damaged plant parts. The barks may be partly or fully eaten. Termites multiply very fast and are capable of destroying the entire tree. After damaging the roots, termites go up the trunk through earthen tunnels. The workers feed on the bark and underlying tissues. Parts affected are partially or totally destroyed.

Prevention/control * Paint or brush the trunk with used diesel oil to discourage the movement of termites from soil to the upper parts of the tree. * Prune crowded branches to allow light penetration. This will provide unfavorable environment for the multiplication of the insect. Termites have soft bodies and die upon exposure to sunlight. * Insecticides can be sprayed to control termites. Be sure to destroy the earthen tunnels before applying insecticides. * For termite mounds, make a hole on one side, deep enough to reach the nest and pour kerosene.

Mango leafhopper
Scientific name: Idioscopus clypealis (Lethierry)

Common names: Blossom leafhopper, Mango leafhopper

Parts affected: Leaves, flowers and young fruits

Destructive stage: Nymphs and adults

Description: Adults are brown to gray with wedge-shaped body. Head is distinct with protruding eyes on the side. Nymphs are smaller, light brown and have no wings.

Nymphs and adults damage the flowers by piercing the tissues and sucking the plant sap which causes withering, drying and falling of individual flowers. Under severe infestation, no fruit develops. The insects excretes fluid called “honey dew”, an excellent medium for development of the fungus, “sooty mold”, which interferes with the photosynthetic activity of the leaves. It also disturbs flower fertilization and spoils the appearance of fruits. Under high insect population, the entire canopy is covered with sooty mold with the leaves and flowers turning black.

Prevention/control * Since hopper population is expected to be high in summer, early induction of mango trees (September, October and November) will minimize hopper problems in the field. * Use light traps during early stages of flower development to attract and kill adults which are ready to lay eggs. To install a light trap, hang the source of light (electric bulb or kerosene operated lamp) on the tree. Place a basin containing a mixture of soap and water (1:10) underneath the light. Hoppers which are attracted to the light are drowned in the solution. One light trap is required per hectare of mango plantation. * Prune crowded branches to discourage hoppers from staying in the tree. Pruning allows good light penetration and makes the habitat unfavorable for hopper development. * Spray insecticide directed to the nymphal stages rather than the adults, hence, detection of this stage is important. (Please see Annex for suggested control measures)

Mango tipborer
Scientific name: Chlumetia transversa (Walker)

Common names: Tipborer, shoot borer

Alternate hosts: Cashew, Guava

Destructive stage: Larva

Parts affected: Young shoots and flowers

Description: While mango tipborer is a common problem on young shoots, the insect is also observed to destroy early flowers. Newly-developed flowers are damaged entirely while mature flowers are cut into half, with the upper portion being destroyed. The insect is becoming a serious problem of mango flowers, especially during early induction.

Prevention/control

The adult destroys flowers from bud emergence to elongation. Hence, early spraying of insecticides is necessary to protect these stages especially during summer. Insecticides recommended for hopper control will also protect the flowers of mango from tip borer infestation. If infestation is minimal, cut affected portion. (Please see Annex for suggested control measures)

Tent caterpillar
Scientific name: Orthaga melanoperalis (Hampson)

Common names: Web worm

Alternate hosts: Cashew

Destructive stage of the pest: Larvae

Parts affected: Leaves and flowers

Description: The insect is destructive to mango leaves. However, when flowers are present, these are also destroyed by the larvae which secrete a web-like structure and feed on individual flowers.

Prevention/control * Prune crowded branches and damaged leaves. * Control infestation on the foliage by insecticidal spray to prevent transfer of the insect to the flowers during the productive stage.

Mango fruit fly
Scientific name: Bactrocera philippinensis sp. n. And

Bactrocera accipitalis (Bezzi)

Common names: Mango fruit fly

Alternate hosts: Guava, Santol, Sineguelas, Starfruit, Guyabano, Chico, Papaya, Passion fruit, Macopa

Parts affected: Fruits

Destructive stage: Adults and larvae

Description: Damage on fruits starts during egg-laying of adult that resembles colorful housefly. Fresh punctures may not be readily recognized until after 3 to 5 days when soft brownish spots appear on the skin and the underlying tissues start to spoil. The larvae cause the major problem since continuous feeding destroys large portions of the flesh. Breakdown of tissues makes the mango fruits unsuitable for consumption. In the field, infested fruits drop to the ground and decay. Under severe infestation, as much as 70 percent of the crop is damaged.

Prevention/control * Collect and bury fruit droppings at least half a meter below the ground to prevent the development of the insect. * Avoid bruising of fruits during spraying since damaged fruits are susceptible to fruit fly attack. * Bag the fruits using newsprint at 55 to 60 days after induction (chicken egg size) to minimize damage from fruit fly. (Please see Annex for suggested control measures) * Avoid planting papaya, guava, sineguelas or santol as intercrops for mango since these fruits are preferred hosts of the insect. On the other hand, cashew and calamansi are less preferred. * Spray the insecticides at 90 to 105 days after induction since fruits at these stages are attractive for egg laying. Recommended insecticides to prevent fruit fly infestation are Baythroid (1 to 1 ½ tbsp per 16 L water), Karate (3/4 to 1 ½ tbsp per 16 L water) and Decis (1 to 5 tbsp per 16 L water). Last spraying should at least be 15 days from harvest. (See annex for recommended control measures)

Mango seedborer
Scientific name: Noorda albizonalis (Hampson)

Common names: Red-ringed mango caterpillar,

Red boring caterpillar, Fruit boring caterpillar, Mango fruit borer and Mango seed borer

Parts affected: Fruits and seeds of mango

Destructive stage of the pest: larvae

Description: Unlike the fruit fly which feeds mainly on the flesh, the mango seed borer consumes both the flesh and seed. Damage starts when the newly-hatched larvae enter the fruit by boring holes through the apex or the narrow tip of the fruit. As the larvae develop, they feed on the tissues beneath the skin. The damaged area later collapses causing the apex to burst and the fruits eventually fall to the ground.

Serious damage is brought about by the destruction of the seed, since a single larva is capable of consuming the entire mango seed in a short period of time.

Prevention/control * Pick the fruits showing damage. Otherwise, larvae will transfer and destroy adjacent healthy fruits. * Collect and dispose infested fruits on the ground by burying them to prevent the insect from completing its life cycle. * Bag the fruits with newsprints a 55 to 60 days after the induction will also minimize damage of the borer. * Adults can be controlled by spray application of insecticide in the afternoon. (Please see Annex for suggested control measure)

Mango pulp weevil
Scientific name: Sternochetus frigidus (Fabricius)

Common names: Pulp weevil

Parts affected: Flesh or pulp of the fruit

Destructive stage: Adult andlarvae

Description: adults lay their eggs on young fruits while the larvae feed on the flesh. Affected fruits fall to the ground. Damage is not visible externally. However, inner tissues are destroyed by the feeding larvae. The pest is found in Palawan which resulted to the quarantine of the province.

Prevention/control * Adults stay away from light. Pruning is a practical means to discourage movement of insect to mango trees. * Collect and properly dispose dropped fruit by burying them half a meter below the ground to prevent the insect from completing its life cycle. * Bag the fruits at 55 to 60 days after induction. * Several pyrethroids can be sprayed to control pests. * Do not allow other mango varieties to flower since they serve as alternate host of the pest.

Twig cutter/borer
Scientific name: Niphonoclea albata (Newman) and N.capito (Pascoe)

Common names: Twig borer (old); Twig cutter (new)

Parts affected: Branches and twigs

Destructive stage: Adult andlarvae

Description: Before laying eggs, the adult cuts or girdles the branch/twig. This is done by nipping the branch halfway, then turning around to make another cut just as deep as, but slightly lower, than the first cut. Affected parts fail to transport nutrients and water causing the terminal leaves to dry up. Dried leaves on the tress canopy are a common sign of twig borer infestation.

Prevention/control * Avoid planting corn as intercrop for mango. * Under brushing of surrounding areas (grasses, creeping vines, etc.) is recommended since these plants serve as habitat for the adult insect. * The insect can be controlled by spraying insecticide.

Mango trunk borer
Scientific name: Plocaederus sp (Guerin)

Common names: Mango bark borer

Parts affected: Cashew

Destructive stage: Larva and adult

Description: The insect belongs to the family of the long-horned beetle. Damage is done by adults who bore holes into the trunk during egg laying. Larvae which develop, feed on inner wood. Water and nutrients are prevented from going up the tree, depriving the leaves and twigs of nourishment. Affected parts dry up and eventually die. Damage is pronounced on neglected and abandoned trees.

Prevention/control * Trees with poor growth are easily attacked and damaged by the mango trunk borer. Hence, application of fertilizer and proper pruning are necessary. Yearly application of 3 to 5kg complete fertilizer is recommended to improve vigor. * Crowded branches should be pruned to provide good light penetration and air circulation within the canopy. * Infected parts should be removed and burned.

Helopeltis/capsid bug
Scientific name: Helopeltis sp.

Common names: Capsid bug

Tea mosquito

Parts affected: Cacao, guava and Cashew

Destructive stage: Leaves and fruits

Description: the adult is a medium-sized bug, black in color with orange marking in the thorax. It is a sucking insect and feeds on young leaves and fruits.

Affected leaves shows dark brown, irregular spots that result in the wrinkling of leaf and blade. On fruits, corky raised irregular brown spots resembling scab-like damage is prominent. Unlike damage of cecid fly, the damage from capsid bug is characterized by spots which are irregular and dry.

This is a result of a substance in the saliva, introduced in the tissue during feeding. The damage is also known as ‘kurikong’, or ‘armalite’.

Prevention/control * Spray insecticide on young flushes.
(See Annex for suggested control measure)

* Prune for field sanitation purposes * Bag fruits * If insecticides are to be sprayed, apply these in the afternoon.

DISEASES

Anthracnose

Causal organism: Colletotrichum gloeosporioides Penz

Parts affected: Leaves, flowers and Fruits

Description: Considered as the most seriousfungal disease of mango in the Philippines, anthracnose occurs in all mango growing areas. It attacks the different parts of the tree, but major damage occurs at flowering and after harvest. It is serious during the wet seasons and usually occurs as a post harvest disease of mango fruits.

The disease is characterized by the appearance of tiny spots on the leaves. These later enlarge to form discrete, rounded or angular spots which come together to form large, irregular shaped-patches with light brown to black spots. However, during advance stage of the disease, the spots give way and produce “shot hole” appearing in various shapes and sizes. This must be differentiated from the “shot holes” produced by the cecid fly which are small and circular.

Anthracnose is the most devastating disease of mango flowers, especially when induction is done early in season (August to October). The presence of rain and high humidity favors the development of disease, thus, the flowers are easily infected. Common signs of the disease are black streaks on the main stalks and branches of the flower, which later become large, black patches. Under severe infection, entire flowers turn black and fail to develop.

Young fruits are also affected and fall prematurely. Symptoms are, however, not visible since the fungus does not have proper condition for development (hard and acidic fruits). After harvest, when fruits start to ripen, the fungus is reactivated and spread over the surface (latent infection)

Early symptoms of the disease are black, pin-pricked lesions. Later, the lesions form bigger black spots, until the whole fruit is covered. The disease is most serious during wet seasons and usually occurs as important post-harvest disease of mango fruits.

Prevention/control for leaves

As a fungal disease, the development and spread of anthracnose are facilitated by high relative humidity within the tree canopy. Young leaves are susceptible to the disease.

* Prune crowded branches to allow light penetration and good air circulation that will create an environment unfavorable for disease development. * Remove dead and diseased branches to reduce the source/reservoir of fungal spores. * Ring cultivation can lessen humidity underneath the trees, which discourage germination of spores.

Prevention/control for flowers * Prune after harvest to increase ventilation and reduce humidity inside the canopy. * Collect and burn trashes to reduce sources of disease inocula. * Some farmers practice shaking of branches after blooming to remove morning dew deposited on the flowers. By doing so, the relative humidity is reduced and male flowers are eliminated, providing enough space for development of hermaphrodite flowers which produce fruits after pollination. * Several chemicals such as Benlate (1 to 2 tbsp per 16 L water). Maneb (4 to 6 tbsp per 16 L water), Dithane (4 to 7 ½ tbsp per 16 L water), and Manzate (½ tbsp per 20 L water) have given varying degree of protection for flowers against anthracnose. These are applied singly or in combination in a sequential spray program. It is also suggested to incorporate any of these fungicides in the flower inducers, especially when flower induction is done early in season. (Please also see Annex for other suggested control measures)

Prevention/control for fruits * Apply protectant fungicides such as Daconil (1/2 tbsp per 16 L water), Manzate (1/2 tbsp per 16 L water), Dithane (4 to 7 ½ tbsp per 16 L water), a week after bud break, at fruit set and 20 days before harvest. (Please also see Annex for other suggested control measures) * Bagging of fruits at 60 days after flower induction can minimize the problem. * Hot water treatment (HWT) by dipping newly harvested fruits in heated water (52 to 55°C) for 10 minutes, followed by hydro-cooling and air drying.

Die-back
Causal organism: Colletotrichum Gloeosporioidez (Penz)

Parts affected: Terminal shoots

Description: The symptoms are discoloration and darkening of the twig at some distance from the tip. As the disease advances, the twig withers, dropping it leaves. Dead twigs are often seen protruding from the tree canopy like “sticks” devoid of leaves. When split often, the twigs show internal discoloration with gummy sap. Blackening of the twigs is associated with the presence of the fungus.

Prevention/control

The fungus multiplies in crowded and shady canopies. To minimize fungal infection, prune to make the environment less favorable for their growth and spray fungicide.

Gummosis
Causal organism: Phytophthora palmivora Butl

Common names: Crown rot, Root rots

Parts affected: Trunk, branches

Description: The disease occurs during both wet and dry seasons causing slow death of mangoes. In seedling stage, infection starts from the roots while in the nursery. In bigger trees, infections extend from the trunk upward and laterally to the branches, where early symptoms are manifested. In seedlings, infection results in root rot, while on big trees, infection is largely confined to the bark with profuse gumming/bleeding. When scraped, the affected part is brown in contrast to green healthy tissues.

Water soaked lesions first appear, followed by ruptures. Latex or gummy sap comes out and hardens to form short colored strips along trunk and branches of the affected area. Affected parts become watery and rot. The whole tree sheds leaves and later dies.

Prevention/control

· Since the fungus is soil inhabiting, sterilize the potting media by pouring boiling water to reduce the source of infection before bagging the seedlings.

· Field planning is recommended in well drained soil (avoid water logged areas)

· Tree spacing of less than 10x10 m apart should be discouraged.

· Avoid root and trunk injuries during cultivation.

· Fungicide can be applied as ‘slurry’ over the affected parts. This is done by mixing the fungicide in water to form a ‘paste’. The later is applied to affected areas.

Seedling wilt
Causal organism: Pythium sp and other soil-borne fungi

Common names: Seedling wilt

Parts affected: Roots and young stem

Description: Considered a serious disease of seedlings grown in plastic bags. The disease is caused by overcrowding, too much shading, and excessive watering. Leaves become dull. Light green with brownish spots near the base. Within a few days, the leaves bend downward, curl, and die. Infected seedlings shoe excessive decay of roots.

Prevention/control * Use sterile soil media. This is prepared by cooking the soil in a half-drum container for 1 to 2 hours or by pouring boiling water on it before bagging the seedlings. * Avoid arranging grafts too closely in the nursery. Early sunlight should reach the plants through partial shading. * Water only when necessary * Remove infected plants from healthy ones to prevent further spread of the sisease.

Scab
Causal organism: Elsinoe mangiferae (Brit and Jenkins)

Common names: Scab

Parts affected: Flowers and Fruits

Description: The disease is more prominent on fruits as compared to leaves and flowers. Young fruits are susceptible to scab and fall to the ground. When examined, irregular, raised corky structures are present on the surface. During rainy days, the scab becomes dark brown but remain light brown during the dry months. On mature fruits, the scabby lesions remain very distinct and persist after harvest affecting quality.

Prevention/control

· Like anthracnose, scab produces spores which remain dormant on dead twigs/branches and trashes below the tree. Prune and collect dead leaves and branches and burn them.

· Apply protectant fungicides a week after bed break, fruit setting and during fruit enlargement. (Please see Annex for suggested control measures)

· Avoid mechanical injuries during harvesting

· Bagging of fruits at 55 to 60 days after induction is a practical way of preventing damage and reduce incidence of scab in the field. Seal the paper bag properly at all sides and let it remain intact up to harvest.

Sooty molds
Causal organism: genus-tripospermum, Limaculina, Trichopelteca, Chaetothyrium,Capnodendron and Polychaeton

Parts affected: Flowers and Fruits

Description: Sooty mold is caused by a fungus which grows andobtains its nourishment from ‘honey dew’ excreted by sucking insects like leafhoppers, scales, and mealy bugs. Although no direct damage is done to the plant, the photosynthetic activity of the leaves is adversely affected. Thus, the trees bear fruits poorly and show a general reduction in vigor. The fungus grows as a thin, black papery film on the surface of the leaves.

The disease develops on the leaf surface as black, papery film especially in areas where honey dew deposits are present. Unlike anthracnose which covers the entire flowers, sooty mold development is randomly distributed on the different parts of the flowers. Affected fruits have a dirty appearance.

Prevention/control * The disease develops from excreta (honey dew) deposited on the leaves by sucking insects like mango leaf hoppers, scales, and mealy bugs. A practical approach to prevent the occurrence of sooty mold is to destroy the sucking insects by spray application of recommended insecticides such as Malathion (1 ½ to 3 tbsp per 16 L water), Decis (1 to 5 tbsp per 16 L water), or Karate (3/4 to 1 ½ tbsp per 16 L water) * Prune infected branches * Bag the fruits to minimize infection of sooty mold * Hot water treatment has been found to facilitate cleaning of fruits affected by the disease, provided it is treated immediately after harvest.

Stem end rot
Causal organism: Dothiorella dominicana (Dd)

Common names: Stem end rot, DSER

Parts affected: Fruits

Description: The disease is considered next to anthracnose in importance and is responsible for about 2-6 percent of storage and transit rots of mango.

The disease is characterized by appearance of dark discoloration near the fruit stalk (pedicel). Under warm and moist conditions, the infected area extends towards the end of the fruit. Later, the symptom turns from dark brown to purplish-black and the tissues become soft and watery. The disease produces soft rot in contrast to the hard rot produced by anthracnose.

Prevention/control * Remove and burn primary sources of the disease such as dead twigs, barks and other trashes. * Since high incidence of stem-end rot occurs on fruits without stalks, harvest the fruits with about 1.0 to 2.0 cm of the stalk attached. * The pre-harvest sprays of fungicides recommended for anthracnose can also be used to prevent stem-end rot. * Avoid the use of organic materials as liners for mango during packaging

OTHER PESTS
Parasitic flowering plants
Scientific name: Family: Loranthaceae

Common names: Parasitic plants

Phanerogamic parasites

Alternate hosts: Many shade trees and fruit trees

Parts affected: Branches and twigs

Destructive stage: All stages of development

Description: Parasitic plants grow on branches of mango trees to obtain nutrient and water. Affected parts are starved, decay, and eventually die

Prevention/control * Prune crowded branches for good light penetration to discourage the growth of parasitic plants * Avoid planting the trees too close (less than 10x10 m) to prevent in the later years. * One percent herbicide has shown to control these parasitic plants. Direct spray on the parasitic plant is recommended to prevent herbicide injury to mango

Mango gallmite
Scientific name: Unidentified

Common names: Mango galls, gall mite

Parts affected: Leaves

Description: The pest is not an insect, but a small mite related to spiders. Mites feed on both young and old leaves which result in the production of galls. The damage is similarly observed in the leaves of ‘Bangkok’ santol. Affected leaves produce galls, curl and photosynthetic activity is reduced. During heavy infestation, growth is stunted. The problem is confined to Mindanao, particularly in Davao and Cotabato.

Prevention/control * Avoid planting ‘Bangkok’ santol as intercrop for mango since the leaves are very susceptible to mite infestation * Prune affected parts * Acaricides are sprayed to control mites

Rat
Scientific name: Rattus argentiventer, R. mindanensis (Mearns)

Common names: field rat, rice land rats

Alternate hosts: Rice, corn, grains and fruit trees

Destructive stage: Both young and adult stages

Parts affected: Stems of newly planted trees and occasionally, fruits

Description: Newly planted trees are attractive to rats, especially when rice or corn fields are vacant. Rats prefer to eat or chew stems of mango resulting to cutting of young plants. As high as 12 percent damage have been reported in the field. In some instances, fruits about to be harvested are also destroyed by rats.

Prevention/control * Cut down and remove weeds in surrounding areas. This will reduce shelter and burrowing sites * Avoid planting rice and corn as intercrops for mango, especially in rat infested areas * Rats can be controlled by baits made of rice bran and rodenticide

FRUIT ABNORMALITIES

BIOKO

Description: a common complaint among growers is the abnormal development of fruits despite good management. Those affected remain small, round and green at harvest. In the Visayas, this abnormality is called ‘Bioko’, in Luzon, ‘ Paninglon’. Bioko can affect 2 to 10 percent of the total fruit production. The cause(s) of this problem is not known although there are speculations that is associated with lack of micro-elements, lack of water during fruit development and side effects of chemicals particularly insecticides. Some growers value the fruits because they command high price in the market and are believed to have aphdisiac properties.

Prevention/control * Apply foliar fertilizers containing micro-elements like zinc, boron, magnesium, and copper. Spray at 42 to 45 days induction and 3 to 4 weeks later. The following foliar sprays are recommended for mango: Albatros (1 ½ to 2 tbsp per 4 L water), Nutraphos (4 tbsp per 16 L water), Wokozin (1 to 2 ml per L), Crop giant (4 tbsp per L) and Agrowel (2 to 3 tbsp per gallon). * Apply water during fruit development at 15 day intervals and stop irrigation one month before harvest.

KASOY-KASOY

Description: ‘Bioko’ fruits are round and green. However, sometimes fruits are malformed and curved, resembling a cashew seed. These symptoms are referred to as ‘Kasoy-kasoy’. Commonly, the fruits split on the curved side and the tips are usually yellow in color. Fruits produced on the tips of the flowers usually exhibit the symptom. Kasoy-kasoy fruits do not grow big and fall to the ground prematurely.

There are indications to show that this problem is associated with parthenocarpy, a process of fruit development without fertilization. This happens when pollination does not occur because of the absence of flies, bees, ants etc. however, other factors such as lack of water and nutrients have to be considered.

Prevention/control * Avoid spraying insecticides during full bloom to protect insect pollinators (flies, wasp, bees, ants etc.) * Maintain flowering plants in the orchard as source of food for pollinators. * Encourage pollinators to visit the trees by spray application of 10 percent honey or sugar solution. Apply the solution at full bloom as spot treatment.