IP-1 taro root aphid - Advanced Crop Science 55

Transcription

Insect Pests
Dec. 1997
IP -1
Cooperative Extension Service
Taro Root Aphid
T
aro root aphid, Patchiella reaumuri, is one of the
most destructive insect pests of dryland (upland)
taro. Taro root aphids feed on the taro roots, and this
can greatly reduce plant vigor, yield, and quality. Crop
losses of up to 75–100 percent have been known with
‘Lehua’, ‘Chinese’, and dasheen taro on the island of
Hawaii.
Damage from taro root aphid feeding is often exten
sive during drought conditions, and it can be especially
severe on young plants in new plantings. The damage
can be extensive because the aphid feeding activity may
go undetected under ground.
The yellow-gray aphid usually is covered with a
mass of fine, white, cottony, waxy threads. Signs of in
festation appear as white mold on the fibrous taro roots
(Figure 1). When populations are high, colonies are
1
Dug-up taro roots with taro root aphids.
found both on roots and around the basal sections of the
leaf sheaths, just above the top of the corm (Figure 2).
The taro root aphid is highly host-specific. It appar
ently infests only taro (and, possibly, closely related
plants of the family Araceae). This aphid has been
present on upland taro on the island of Hawaii since
1971. It has been present on Oahu since 1995, when it
was found in commercial plantings in the Kahuku and
Mililani areas. An infestation was observed in a com
munity garden plot on Lanai in 1994, but prompt de
struction of the infested plants prevented further spread
and establishment there.
The taro root aphid apparently does not attack taro
grown under wetland conditions.
In Hawaii, this species does not produce winged
sexual forms, and reproduction occurs without fertili
2
Taro root aphid infestation on taro leaf petiole and sheath.
Issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in cooperation with the U.S. Department of Agriculture. Charles W. Laughlin,
Director and Dean, Cooperative Extension Service, College of Tropical Agriculture and Human Resources, University of Hawaii at Manoa, Honolulu, Hawaii 96822. An
Equal Opportunity / Affirmative Action Institution providing programs and services to the people of Hawaii without regard to race, sex, age, religion, color, national
origin, ancestry, disability, marital status, arrest and court record, sexual orientation, or veteran status.
IP-1
Taro Root Aphid
zation by males. Taro root aphids have been observed
to be associated with numerous attending ants, which
probably move the aphids around, enabling them to de
velop damaging populations.
Control
Spread of the taro root aphid occurs mainly by the plant
ing of infested hulis (cormels, used as seedpieces). A
hot-water dip treatment to disinfest taro hulis of root
aphids has been developed by entomologists at the Col
lege of Tropical Agriculture and Human Resources. Dip
ping taro hulis for 6 minutes in water held at 120°F
(49°C), followed by immersion in cool water, will dis
infest them of root aphids without significant effects on
the hulis.
A taro crop planted with infested hulis will never
get off to a good start, and subsequent yield will not
reach adequate levels, especially if periods of drought
occur. It is very important, therefore, to plant clean hulis
and to grow upland taro only in unaffected areas.
No effective insecticide is currently available for
use against root aphids on taro.* Should an insecticide
become available, it will likely be most useful as a con
trol measure when applied during the early growth phase
of the taro crop.
CTAHR — Dec. 1997
If a heavy infestation of taro root aphid occurs, the
crop should immediately be removed and destroyed,
with care to include all culls and unharvested cormels.
The field should be given a thorough and deep cultiva
tion to drive away ants and to promote root degrada
tion. After cultivation, fallow the field or grow non-taro
crops for at least one year.
Quarantine regulations prohibit shipment of taro
hulis from the island of Hawaii to other islands in the
state. To reduce the risk of introducing the taro root aphid
to other locations in the state of Hawaii where taro is
grown, these regulations should be revised to include
Oahu. In the meantime, shipping taro planting materi
als (or taro corms with hulis attached) from Oahu is not
recommended.
The College of Tropical Agriculture and Human
Resources has done the research necessary for approval
of pesticides for control of the taro root aphid. If ap
proved by regulatory agencies, these pesticides may
become available for use. Contact your local Coopera
tive Extension Service office for current information on
the status of pesticides for use against the taro root aphid.
Prepared by Dwight M. Sato1 and Arnold H. Hara2 with the assis
tance of Ronald F.L. Mau 2 , Dick M. Tsuda 2, and Randall T.
Hamasaki3, this publication revises and replaces Commodity Fact
Sheet TA-4(A), Taro root aphid, by Sato, Hara, and Jack Beardsley2,
published in 1989. Photos courtesy of Julie Coughlin4.
1
CTAHR Cooperative Extension Service, Hilo
2
CTAHR Department of Entomology
3
CTAHR Cooperative Extension Service, Kaneohe
4
CTAHR Department of Environmental Biochemistry
*Certain insecticidal soaps labeled for use against aphids on root and tuber vegetables are currently available. Insecticidal soaps may control
some aphids, but their efficacy on taro root aphid is uncertain because it is a very waxy aphid, which may protect it. Also, aphids on roots
underground are difficult to contact with spray solutions. Furthermore, insecticidal soaps have been observed in some cases to burn taro
leaves, particularly when applied during the heat of the day.
Unless the label says otherwise, insecticidal soaps labeled for use on root and tuber vegetables may be used as a dip for treatment of hulis
before planting. Again, their efficacy against taro root aphids is uncertain. The dip should be at the spray concentration given on the label
(generally about 1% active ingredient in the spray solution). Any remaining unused solution should not be dumped on the soil but rather should
be sprayed over areas bordering the growing area, where pest reinfestation is likely.
2
Insect Pests
Nov. 1998
IP-2
Bougainvillea Looper
B
ouganvillea loopers, as the
name suggests, feed primarily
on bougainvillea, but they have also
been reported to feed on other plants
in the Nyctaginaceae family, such
as the four-o’clock (Mirabilis
jalapa). This looper has most often
been observed feeding on the com
mon purple bougainvillea, but it
does not appear to have a preference
for one bougainvillea variety over
another—it likes them all.
Disclisioprocta stellata (Guenee)
Order Lepidoptera, family Geometridae
Description
The bougainvillea looper is a green or brown caterpillar
about 1 inch long. It is also called “inchworm” or “mea
suring worm” because it moves in alternate contractions
and expansions suggestive of measuring. The looper
larva mimics stems and branches very well and feeds
primarily at night, which is why you may see the dam
age but fail to find the culprit on the plant.
The adult is a moth, a very fast flyer with a wing
span of about 1 inch. The moth does not feed on the
foliage. Like the larva, it also is
active at night, when it is believed
to lay its eggs on the underside of
bougainvillea leaves.
Damage
The bougainvillea looper feeds from the edges of the
leaves, which results in severe scalloping of the foliage.
Attacks begin on the young, tender shoots and leaves
before progressing down the stem. The loopers may
move down the stems during the night and take shelter
on the larger interior branches during the day. As the
population multiplies, entire shrubs can be defoliated.
To date, the bougainvillea looper has not generally been
regarded as a serious pest. The insect will cause signifi
cant visual damage to bougainvillea, although this does
not apparently result in the death of the plants.
Distribution
The bougainvillea looper is a very
wide-ranging, migratory species
from tropical America. It is a rela
tively new pest in Hawaii, first re
ported on Oahu in 1993, and since
then has spread to Maui, the Big Is
land, Kauai, and probably Molokai.
Although it could have been intro
duced to Hawaii with nursery stock,
it is possible that it became estab
lished naturally through long-range
dispersal, because the moths can
travel great distances on air currents.
Control
Bacillus thuringiensis (BT, or Dipel®) and neem-based
biological insecticide products should be effective on
the loopers without harming other insects that may bio
logically control them, such as parasitic, mud, and pa
per wasps. Insectical oils and soaps will not control cat
erpillars such as the looper.
Most synthetic insecticides with labels permitting
use against caterpillars on landscape ornamentals, such
as carbaryl (Sevin®), will likely kill the bougainvillea
looper, although these products are often destructive to
beneficial insects as well.
Spraying insecticides late in the evening is recom
mended. This is when the bougainvillea looper caterpil
lars and adult moths are active, and also when the ben
eficial insects are not likely to be active.
James Tavares1, David Hensley2, Jay Deputy2, Dick Tsuda3,
and Arnold Hara3
1
Cooperative Extension Service, Kahului; 2Department of
Horticulture; 3Department of Entomology
Mention of a trademark or proprietary name does not constitute an endorse
ment, guarantee, or warranty by the University of Hawaii Cooperative Exten
sion Service or its employees and does not imply recommendation to the
exclusion of other suitable products. Pesticide use is governed by state and
federal regulations. Read the pesticide label to be sure that the intended use
is included on it, and follow all label directions.
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and issued in furtherance of Cooperative Extension work, Acts of May 8 and June
30, 1914, in cooperation with the U.S. Department of Agriculture. Charles W. Laughlin, Director and Dean, Cooperative Extension Service, CTAHR, University
of Hawaii at Manoa, Honolulu, Hawaii 96822. An Equal Opportunity / Affirmative Action Institution providing programs and services to the people of Hawaii without
regard to race, sex, age, religion, color, national origin, ancestry, disability, marital status, arrest and court record, sexual orientation, or veteran status.
ALIEN
PEST
ALERT!
Red Imported Fire Ant
A Seriously Harmful Potential Invasive Species
T
Neil J. Reimer and Carol Okada, Hawai‘i Department of Agriculture
he red imported fire ant, Solenopsis invicta, na
tive to South America, is a serious pest of agri
cultural, urban, and native environments in areas that it
has invaded. This species is not known to be present in
Hawai‘i but is related to the tropical fire ant, Solenopsis
geminata, which is present in Hawai‘i. The red imported
fire ant, however, is much more aggressive.
Workers and queen, relative sizes
Infested areas
Potential areas
of infestation
Distribution in the United States
The red imported fire ant was accidentally introduced into
Alabama in the 1930s and has since spread throughout
the southern USA. It now occurs in Alabama, Arkansas,
California, Florida, Georgia, Louisiana, Mississippi, New
Mexico, North Carolina, Oklahoma, South Carolina, Ten
nessee, Texas, and Puerto Rico. There have been spot in
festations in Arizona, but these have been eradicated. This
pest will continue to spread on the Mainland. Its distri
bution appears to be limited by temperature and mois
ture: it does not tolerate freezing well, and it does poorly
in areas that receive less than 10 inches of rain per year.
Distribution in Hawaii
At present, the red imported fire ant is not found in Ha
waii. However, conditions in Hawaii are definitely con
ducive to its survival. The Hawaii Department of Agri-
Mounds in a pasture
Workers, actual sizes
culture regards it as a high priority to prevent the red
imported fire ant from establishing in Hawaii.
Life cycle and biology
The life cycle of this ant is similar to many other pest
ants. The colonies (“mounds”) can contain 10–100 or
more queens, which each lay up to 800 eggs per day.
After 7–10 days, the eggs hatch into larvae, which de
velop over a 6–10-day period before pupating. After
another 9–15 days, the adult emerges from the pupa.
Soil from excavation of the colony nest is mounded
at its entrance. The ants will nest in any soil and habitat,
but they prefer sunny, open areas such as pastures, fields,
parks, and golf courses. Pasturelands may have 250
mounds or more per acre, each containing from 80,000
to 500,000 worker ants.
30, 1914, in cooperation with the U.S. Department of Agriculture. Andrew G. Hashimoto, Director/Dean, Cooperative Extension Service/CTAHR, University
CTAHR publications can be found on the Web site <http://www.ctahr.hawaii.edu> or ordered by calling 808-956-7046 or sending e-mail to [email protected].
ALIEN
PEST
ALERT!
2
Red Imported Fire Ant
Human health risks
Red imported fire ants are very aggressive toward any
thing that disturbs their mound. They can sting repeat
edly. Typically, the ant grasps the skin with its jaws and
inserts its stinger into the flesh, injecting venom from
its poison sac. Pivoting its head, it can inflict an average
of seven to eight stings in a circular pattern.
Typical red imported fire ant sting symptoms.
Symptoms of each sting are a burning and itching that
lasts about an hour. A small blister will form in a few hours,
followed by a white pustule in a day or two. Scratching
the stings can lead to infection and scarring. Reaction to
the sting ranges from localized swelling with pustule for
mation to severe, life-threatening anaphylactic shock.
Individuals who have a severe reaction to the venom
may suffer chest pains, nausea, swelling of the face and/
or throat, sweating, loss of breath, or slurred speech.
Diabetics and others with circulatory disorders includ
ing varicose veins and phlebitis are at risk for complica
tions. In 1988, 32 human deaths were attributed to these
ants in the United States.
Agricultural impacts
Domesticated animals attacked by red imported fire ants
are susceptible to anaphylactic shock, and their sensitiv
ity can vary with age and amount of exposure. Young ani
mals, if they are unable to escape, may be blinded or killed.
The ants feed on germinating seeds and can destroy
buds and developing fruits, thus causing serious dam
age to crops. They also cause extensive damage to seed
lings and saplings by girdling stems and branches.
Mounds built in clay soils become hard as rock and dam
age farm machinery.
Environmental impacts—
urban and recreational
The red imported fire ant is a serious problem in urban
and recreational environments. Its presence will deter
people from outdoor recreational activities. Playgrounds,
athletic fields, parks, and golf courses must either be
heavily treated with pesticides to control these ants, or
they are best left unused.
These ants often form nests near buildings and for
age into the buildings for food and water. They will oc
casionally nest in electrical equipment, such as air condi
tioners, traffic signal boxes, and other devices, causing
shorts. Fire ants have a major impact on ground-nesting
species, such as birds, rodents, and insects. The decima
tion of insects will reduce the food supply of native wild
life and negatively impact the pollination of native plants.
What to look for and who to contact
The red imported fire ant looks very much like the fire
ant already present in Hawai‘i. The two species can be
accurately differentiated only by an expert, but there are
some characteristics which may help distinguish them:
Red imported fire ant
Tropical fire ant
Solenopsis invicta
(Not present in Hawai‘i)
Solenopsis geminata
(Present in Hawai‘i)
Builds mounds
Never builds mounds but
may form small dirt piles
Very aggressive;
expect many stings
Less aggressive;
expect just a few stings
Sting causes small
blister followed by
white pustule
Sting causes small red
swelling
Found in any environment
including dry coastal areas
Generally restricted to
dry coastal areas
No large-headed workers
Some workers with
large, bi-lobed heads
If you suspect that you have seen a red imported
fire ant, or to obtain more information, contact the Ha
wai‘i Department of Agriculture on O‘ahu at 586-PEST
(586-7378); on Moloka‘i and Läna‘i at 800-468-4644;
on Hawai‘i at 974-4000 ext. 67378; on Kaua‘i at 274
3141 ext. 67378; or on Maui at 984-2400 ext. 67378.
UH-CTAHR publication IP-3 (revised)—Nov. 2004
Insect Pests
Sept. 1999
IP-4
Managing Fruit Flies on Farms in Hawaii
Russell Messing, Department of Entomology
F
ruit flies have become serious pests in Hawaii since
the first species was found here in about 1895. They
are widespread, occurring from sea level to above 7000 ft
elevation, and feed on hundreds of host plant species, many
of which are economic crops.
Four species of fruit flies in the family Tephritidae are
now known in Hawaii. The melon fly is commonly found
in commercial and backyard vegetable gardens at low el
evations. The Mediterranean fruit fly (“medfly”) moved
away from most lowland areas (except low-elevation cof
fee fields) when the oriental fruit fly arrived in 1945, and it
is now found more frequently in upper elevations. The ori
ental fruit fly is found in most elevations and climates. The
solanaceous fruit fly survives in both cool and hot climates
but so far has been found only in dry areas of Hawaii (<100
inches of rain per year).
This publication provides information to help farmers
and gardeners identify pest fruit flies, learn about their
habits and life cycles, and implement strategies to manage
them and reduce crop damage. A glossary defining some
of the terms used is on page 7.
Most control strategies use a combination of tech
niques—no single, “one-answer” solution to the fruit fly
problem is available. The postharvest treatments required
for export of commodities affected by fruit flies are not
covered in this publication.
Damage caused by fruit flies
Plant injury. Fruit fly adults most often lay their eggs in
the fresh flesh of fruits and vegetables. The eggs hatch into
larvae (maggots), which most often feed on the inside of
the fruit, resulting in a soft, mushy mess. Look for wig
gling white larvae the next time you pick a very ripe guava
or other fruit.
Economic injury. Fruit flies can often be present at
low levels without causing significant economic problems,
so control may not be necessary. If high fruit fly popula
tions are causing more severe damage, management prac
tices may need to be implemented.
Key steps in managing fruit flies
• Prevention—practice sanitation techniques.
• Monitor the levels of pests; determine if you have eco
nomic injury; evaluate and use the best strategies.
• Identify the fruit fly species and become familiar with
its life cycle and host plants.
• Determine which other plants in the area are fruit fly hosts,
and determine when these plants are fruiting.
• If possible, rotate your crops so they do not fruit when
other hosts are fruiting and pest populations are peaking.
• Harvest fruits under-ripe when possible (e.g., papayas are
usually fruit fly–free if picked when less than 1⁄4 ripe).
• If fruit flies cause economic injury, apply appropriate
controls.
• Divert pests with poisoned border plants, baits, or lures.
• Monitor pests again and reevaluate your strategies.
Life cycles of fruit flies
Fruit fly development (life cycle) depends on temperature.
Cool temperatures slow the development cycle, while warm
temperatures speed it up. Information on life cycles given
here is derived from laboratory-raised fruit flies grown at
77°F at 50% relative humidity, except for the solanaceous
fruit fly (80°F at 60% RH); wild flies will most likely be
different. Traits common to all four species include
• eggs are white, up to 1⁄l6 inch long
• larvae range from 1⁄l6 to 3⁄8 inch long (just before pupat
ing, the larvae often “pop” and flip to leave the fruit)
• pupation normally occurs 1–2 inches under the soil
• adults usually rest in shady locations unless feeding,
mating, or laying eggs; most feed at dawn and mate at
dusk.
This publication replaces HITAHR Brief no. 114, 1995, Introduction to managing fruit flies in Hawai‘i, by Laurel Dekker and Russell Messing.
30, 1914, in cooperation with the U.S. Department of Agriculture. H. Michael Harrington, Interim Director/Dean, Cooperative Extension Service/CTAHR, University
IP-4
Melon fly
CTAHR — Sept. 1999
Mediterraneanfruit fly
Four species of tephrited fruit fly are found in Hawaii. Wing pattern is the best distinguishing characteristic; color is
inconsistent and not always reliable. See “key characteristics” in the descriptions below for distinguishing features.
Melon fly
Scientific name: Bactrocera cucurbitae; native to Asia; de
tected in Hawaii around 1907.
Key characteristics: Wing pattern has stripes and a large
black spot at the wing tip. Abdomen is usually brown with
a gold to brown horizontal band and a faint black “T”. Ovi
positor (egg-laying tube) has a plump, straight sheath (outer
covering) and is about 1⁄l6 inch long.
Distribution: parts of Africa, Burma, Sri Lanka, China,
Guam, Hawaii, New Guinea, Rota, Commonwealth of the
Northern Marianas, Southeast Asia, and South Asia; sea
level to 4500 ft.
Hosts: Over 100 known. Preferred hosts are cucurbits
(squash, melon, etc.). Other hosts include solanaceous
plants (tomato, eggplant, pepper, etc.) and papaya.
Life cycle: One generation takes around 37 days; egg
to adult in 15–18 d; eggs hatch in about 30 hr; larvae de
velop in 7–8 d; adults emerge in 9–10 d; pre-oviposition
period is 7–8 d; females lay an average of 15 eggs /day,
singly or in clusters.
Special notes: Known to feed on stem shoots and buds
of squashes and melons.
2
Mediterranean fruit fly
Scientific name: Ceratitis capitata; native to sub-Saharan
Africa; first reported in Hawaii in 1895.
Key characteristics: Wing pattern is very complex and
multicolored (gold and black) with black stripes and de
tailed markings. Black spots are on the back or thorax. Ab
domen is usually brown. Adult is about 2⁄3 the size of the
other fruit flies.
Distribution: Africa, Mediterranean countries, Hawaii,
western Australia, Central and South America; the domi
nant fruit-fly pest in Hawaii above 3000 ft and in low-el
evation coffee; prefers dry regions.
Hosts: Over 300 hosts. Preferred hosts include coffee,
peach, plum, loquat, orange, guava, rose apple, solanaceous
plants (pepper, Jerusalem cherry), and the sapote family,
among others.
Life cycle: One generation takes around 18–31 days;
egg to adult in l9 d; eggs hatch in about 2–3 d; larvae de
velop in 7–8 d; adults emerge in 9–10 d; the pre-oviposi
tion period is about 3 d; females lay an average of 10 eggs/
day, singly or in clusters of up to 10.
IP-4
Oriental fruit fly
Solanaceous fruit fly
Photographs are from the website of the USDA Agricultural Research Service’s Tropical Fruit, Vegetable, and Ornamental Crops Laboratory, Hilo, Hawaii.
Oriental fruit fly
Scientific name: Bactrocera dorsalis; native to Asia; in
troduced to Hawaii in 1945.
Key characteristics: Wing pattern has two solid black
lines stemming from the point of attachment, without a
black spot at the tip as in the solanaceous fruit fly. Abdo
men is gold to brown with gold to brown horizontal band
and prominent black “T”. Ovipositor has a slender, straight
sheath.
Distribution: Asia, Australia, Surinam, and islands of
the Pacific; the major fruit fly pest in Hawaii at low eleva
tions, except for coffee fields.
Hosts: Over 200 wild and cultivated hosts. Preferred
hosts include guava, mango, papaya, starfruit, passion fruit,
citrus, fig, rose apple, tomato, and many more.
to adult in 19 d; eggs hatch in about 38 hr; larvae develop
in 7–8 d; adults emerge in 10–11 d; the pre-oviposition
period is 6–7 d; females lay over 130 eggs /day, usually in
groups of 10 but as many as 100 or more.
Solanaceous fruit fly
Scientific name: Bactrocera latifrons (also known as Ma
laysian fruit fly); native to South and Southeast Asia; first
detected in Hawaii in 1983.
Key characteristics: Wing pattern has two solid black
lines stemming from the point of attachment, plus a black
spot at the wing tip that differentiates it from the oriental
fruit fly. Abdomen is usually brown, without a prominent
“T”. Ovipositor is tri-lobed, to 1⁄16 inch long.
Distribution: China, Taiwan, Malaysia, Thailand, Laos,
India, Pakistan, and Hawaii.
Hosts: 33 reported hosts, mostly solanaceous (pepper,
tomato, eggplant, apple of sodom), and occasionally cu
curbits.
to adult in 21 d; eggs hatch in about 2 d; larvae develop in
8–9 d; adults emerge in 10 d; the pre-oviposition period is
10–11 d; females lay an average of 10 eggs/day, one at a
time.
Special notes: Occurrence is generally in low num
bers with a patchy distribution.
3
IP-4
Prevention strategies
Exclosure. Crop damage can be prevented by keeping fruits
out of reach of female fruit flies. Screen-houses can pro
duce fruit-fly–free crops. Local research has found that an
economical structure (~$1.20/sq ft) was cost-effective
within the first harvest for tomato production. Netting
(floating row covers or lightweight netting from a fabric
store) can be placed directly on plants or on a frame of
PVC tubes for temporary cover of crops like zucchini. To
matoes and self-pollinating cucumbers are pollinated by
the wind, but some other crops may need hand pollination
if plants are covered by screen. A possibility that has not
been fully explored is to add bee hives to large screenhouses
to provide ample pollination. (Note: secondary insect or
weed problems may arise from reduced air circulation and
lack of beneficial insect populations in enclosed areas.)
Another method of exclosure is bagging individual
fruits with newspaper, paper bags, or other barriers. This
method works well but is labor-intensive.
Sanitation. Remove fruits as they ripen. If they fall to
the ground, be sure to kill any larvae in them by burying
the fruit deeply or putting them in an air-tight container for
four days or until no movement is found. Check for pupae
(and destroy them) before adding fruit to compost piles.
Sanitation by itself will not be effective in many situations,
because fruit flies can fly in from outside areas. Melon fly
pupae buried as deep as 2 ft have managed to emerge as
adults from dry sand, wet sand, and soil. When composting,
the pile must achieve internal temperatures of at least 120°F.
Mowing or shredding ground fruit can provide sanitation
by killing the larvae or exposing them to other predators.
Harvest early. By harvesting early, you can sometimes
prevent infestation (e.g., fruit flies do not usually sting
papayas or ‘Sharwil’ avocados that are less than 1⁄4 ripe).
However, some fruits lose flavor when harvested too early,
as they will not ripen fully.
Reduce populations. If fruit flies are present in your
field prior to crop ripening, you can try to reduce their popu
lation by attracting the adults to a poisoned bait. This can
be done by spraying a protein-bait–insecticide mixture onto
nearby non-crop plants, windbreaks, or a border of corn
plants. Farmers and researchers have observed reduction
of melon flies in zucchini, cucumber, and watermelon fields
when using bait sprays on border crops. Suppression sprays
have also been used in Australia, Israel, Mexico, Florida,
and California. Mass trapping with protein baits (for male
and female fruit flies) or with chemical lures (for males) is
being researched as a method of fruit fly reduction.
4
Create an “isolated” area. Planting between other
crops or rotating to opposite ends of a field has been tried
for a few crops (melon fly hosts). Often, fruit flies do not
find the crop during the first half of the harvest. This strat
egy should not be repeated in consecutive plantings in the
same place.
Plant resistance can help. High levels of citrus oil in
immature citrus peels can be toxic to larvae, so research
ers are investigating the use of plant growth hormone
(giberellic acid) to delay peel ripening and reduce suscep
tibility to fruit flies. Mango cultivars are being developed
to have flesh that is harder and crisper when ripe. Small
tomatoes (Roma and cherry) can be infested by fruit flies,
contrary to popular belief; however, many growers have
found that small tomato varieties can be harvested with
less infestation than large varieties.
Don’t confuse fruit flies with vinegar flies. Note that
the fresh-fruit–eating fruit flies discussed here (tephritid
family) are not the same as the tiny “fruit flies” that feed
on yeasts and decaying fruit. These tiny flies called vin
egar flies belong to the drosophilid family and can often
by found on soggy fruits on the ground or overripe, fer
menting fruits.
Control of fruit flies
Note on using pesticides: Read the pesticide label com
pletely. Apply according to manufacturer’s recommenda
tions only to crops specified on the label. If in doubt, con
tact your local Cooperative Extension Service office or the
Hawaii Dept. of Agriculture, Pesticides Branch. If infor
mation given in this publication is different from the label
directions, follow the label directions.
Cultural and chemical controls
Bait spray. In fruit-fly–infested areas, a protein hydroly
sate compound, such as Nu-lure® or Staley’s® bait, can be
combined with insecticide and applied to plants that are
associated with the resting and feeding areas of the adults,
rather than on the crop to be protected. Bait sprays use small
amounts of chemical and are not generally attractive to ben
eficial insects that may be natural enemies of fruit flies and
other pests.
To apply with a knapsack sprayer, find a malathion
product cleared for use on the target site. Follow the direc
tions for fruit-fly control on the pesticide label. For ex
ample, mix the appropriate amount of malathion 25% WP
with 1 qt Nu-lure and 3 gal water; or 1 part malathion 57%
EC with 3 parts Nu-lure. To apply with a conventional
IP-4
power sprayer of 20–100-gal capacity, mix 1 qt Nu-lure
with the appropriate amount of malathion. Agitate during
application. Spray with concentrated, coarse droplets on
border plants that are listed on the pesticide label. Apply
weekly (for high populations) to bi-weekly (for low popu
lations). Reapply after rain. Researchers and farmers have
observed good control of melon flies with this technique.
The Hawaii Department of Agriculture’s Pesticides Branch
has allowed application of pesticide bait sprays to other
border plants and windbreaks under certain conditions.
Note that this policy may change—contact your HDOA
Pesticides Branch district office for current information.
Note also that the mixtures described above have a pH of
4.7; recent research indicates that a pH of 9.2 is more at
tractive to the flies, so researchers are looking at ways to
raise the pH.
Spot treatments with bait spray–insecticide mixtures
have been used successfully elsewhere, but these methods
may not be included on current labels in Hawaii. In Mexico,
bait spray has been applied to orchard tree trunks with good
results. Israeli producers have found spot treatments ef
fective for medflies in or around fields when applied at 2
oz per spot, spaced at 40–80 spots per acre, with 16–33
feet between spots.
Insecticide sprays. Insecticides applied to kill fruit
flies directly should be used only as a last resort and only
on crops allowed on the pesticide label. At least 40 pesti
cides have been found toxic to fruit flies, including
malathion and naled. Pyrethrum is not as toxic to fruit flies
as malathion. Most pesticides, including permethrin, are
more toxic to beneficial insects (such as parasites of pest
insects) than to fruit flies.
Approved organic controls
Neem. In research tests, neem-treated sand was found to
be toxic to oriental fruit flies and medflies but not to sev
eral beneficials. This suggests potential for soil treatment
to inhibit fruit fly development in fields (however, adults
may still invade from outside areas). Azatin® is a neem
product registered for use in Hawaii as a soil treatment
against fruit fly larvae. The National Organic Standards
Board has approved use of neem in certified fields, but it is
still investigating the inert ingredients in Azatin.
Biological controls
Chickens and guinea hens may eat some fruit-fly larvae
found at the top of the soil. Wild birds have also been seen
digging through infested fruits for larvae. Birds and fowl
may also help with sanitizing infested fruits.
Ants are known to feed on most life stages of fruit
flies (research reports up to 40% kill), and earwigs have
been reported to feed on fruit fly larvae.
Nematodes are among the soil-borne organisms that
feed on insect pests in the soil. Nematodes are microscopic
roundworms with a broad host range, including fruit fly
larvae. Currently, commercial use of the nematode
Steinernema carpocapsae is not permitted in Hawaii, but
in the future this may become a viable control for areas
heavily infested with fruit flies.
Fruit fly parasites are tiny wasps that attack only fruit
flies. Parasites can lay their eggs in the egg, larva, or pupa
of a developing fruit fly. The parasite develops within the
immature stages until the fruit fly pupa is consumed, and
then the adult parasite emerges from the soil. Parasites can
be very effective in controlling fruit flies—reports have
indicated up to 90% kill of oriental fruit flies in unsprayed
guava.
Species that parasitize tephritid fruit flies have become
established in the state of Hawaii after being introduced
for biological control. All evidence indicates that these re
ported parasites do not harm any other species besides fruit
flies. Many additional parasites exist in Africa, Asia, and
South America. Do not attempt to bring in beneficial in
sects yourself; to do so violates stringent import regula
tions that protect Hawaii from alien species.
Rearing for identification—Get to know your pests
Raising larvae to adulthood is the best way to identify the
fruit fly species attacking your crops. An easy home method
uses a wide-mouth plastic container with a lid. Make some
small air holes in the top. Place a small amount of infested
fruit with wriggling larvae inside the clean container. Ob
serve regularly, making sure there is no liquid collecting
on the bottom. Soil or sand can be added to prevent drown
ing. As the larvae age, they will leave the fruit to pupate.
You can remove the fruit after the pupae are formed. The
adults will emerge after 9–11 days. Compare them with
the descriptions given on pages 2–3.
Beneficial wasps that are parasites of fruit flies can be
reared in the same way. Because the wasps are small, make
the holes in the top smaller than 1⁄16 of an inch, or put a
tissue or small-mesh screen between the top and bottom of
the cup. Adults will emerge in 2–10 days from ripe fruit.
5
IP-4
Trapping strategies
Monitoring with traps
Monitoring helps identify fruit fly pests, keeps track of
changes in their population levels, and indicates when or
whether to use controls. The best way to detect the pres
ence of fruit flies and evaluate the effectiveness of control
measures is to monitor fruit infestation.
Liquid traps with food bait attract males and females.
Put 1–2 inches of bait mix into the trap, and check weekly.
Yeast tablets: mix five Torula® yeast tablets in 2–21⁄2 cups
water; stir to dissolve tablets. Protein hydrolysate: mix 11
fluid oz Nu-lure® or Staley’s Fly Bait®, 7 fluid oz borax, and
31⁄2 qt water. Fruit: blend cucumber or other primary host
with water; place small amount in trap; change often.
Parapheromone lure traps use highly volatile lures
which attract male flies; these traps need to be checked
frequently. The amount of lure determines how attractive
and long-lasting these traps will be. Lures catch only males,
leaving the females in the field to infest the fruit. At present,
only methyl eugenol for oriental fruit fly is available in
Hawaii.
To attract male fruit flies, initially use 3–5 drops of
lure in a trap. Adding an insecticide to the lure provides a
better catch than traps without insecticide. Use 1 drop of
an insecticide approved for use on your crop for every 20
drops of lure used. Replenish the lure as needed, using more
lure to attract males over longer distances and for longer
time periods. Only insecticides that are EPA-registered and
labeled for use on that crop may be used. The Hawaii De
partment of Agriculture Pesticides Branch has agreed that
parapheromone lures with insecticide may also be used in
fields with non-approved crops to collect fruit flies for sur
vey purposes only in properly labeled traps (this policy may
change).
Yellow spheres or sticky panels are also used to moni
tor fruit flies in crop fields. Check them regularly, and
change them when the trapping surface is full or becomes
dusty.
Mass trapping
High-density trapping is being explored to reduce or sup
press populations of fruit flies. USDA researchers have not
produced evidence that small-scale trapping helps reduce
infestation. However, mass trapping is used in other areas.
In Crete, it resulted in substantial reduction of insecticides
used against a fruit fly. Local research is needed to deter
mine if small-scale suppression of fruit flies can be effec
tive.
6
Types of attractants
Food baits are effective, mild attractants for males and
females of all four fruit fly species. Food baits are not very
volatile, so bait traps typically have lower catches than the
parapheromone lure traps, but food baits can be used di
rectly in the field.
Torula® yeast tablets are more effective than Nu-lure
over time, because the pH is stable at 9.2. The level of pH
in the mix plays an important role in attracting fruit flies.
Fewer fruit flies are attracted to the mix as the pH becomes
more acidic. USDA researchers are testing a combination
of Torula® yeast and dyes commonly used in cosmetics
and drugs to improve population reduction of medflies and
oriental fruit flies.
Nu-lure® (a yeast extract) and Staley’s Fly Bait® (a
corn extract) are hydrolyzed proteins. They are not effec
tive over time as the pH drops from its initial state of 8.5.
Promar®, an experimental hydrolyzed protein developed
in Australia, has been very effective against a species similar
to the oriental fruit fly in Malaysia, where starfruit orchards
with Promar® spray applications rather than insecticidal
cover sprays have doubled yields, mostly due to more bees
being available for pollination.
Farmers report that homemade baits (cucumber or zuc
chini blended with water, or vinegar plus yeast) have at
tracted both males and females of the melon fly.
Parapheromone lures are very volatile and longer last
ing than protein baits. They attract only males, and each
fruit fly species in Hawaii is attracted to a different kind.
The amount of lure used depends on whether the trapping
is for monitoring or for mass trapping. A few drops may be
effective to sample the population over a short period of
time, but more is needed for mass trapping over a longer
period. The kind of lure also affects the amount needed. In
California, detection traps with methyl eugenol are set at
two per square mile, whereas with tremedlure 10 traps are
needed for the same area (6 ml of lure per trap in both
cases). In Hawaii, three to five drops of methyl eugenol
have been used in within-field traps.
Parapheromone lures for male fruit flies
Type of lure
Strength
Fly attracted
methyl eugenol
Cue-lure
Ceralure
Trimedlure
Latilure
very volatile and persistent
moderately persistent
persistent
moderately persistent
mildly persistent
oriental
melon
medfly
medfly
solanaceous
IP-4
The effectiveness of traps varies with their color and shape.
Yellow is the most attractive color to males and females of
oriental fruit fly, melon fly, and medfly. They are attracted
to yellow and white flat panels as well as spheres. In field
tests, researchers collected both females and males from
the colored traps.
lure traps have been spaced 100 ft apart outside the field.
The visual range of fruit flies is about 15–20 ft. Yel
low traps should be placed within that distance from the
host plants and at greater density than lure traps. Monitor
ing programs on the U.S. mainland recommend that traps
be placed 4–6 ft above the ground.
Types of traps
All traps used for catching fruit flies must be properly la
beled with the name of the bait or lure and date the trap
was set. Keep traps out of reach of children.
Glossary
Bait An attractant and food source (sometimes mixed with
insecticide) for treating fruit-fly–infested areas.
Beneficial organisms Birds, insects, nematodes or other
organisms that aid in controlling pests.
Development Growth through life stages or life cycle. Fruit
flies have four life stages: egg, larva, pupa, and adult.
Generation The time it takes to complete all stages of de
velopment, including the pre-oviposition period.
Host A plant or animal that provides food for larval growth
and development.
Infestation The presence of a fruit fly in a host.
Integrated pest management A control strategy that in
tegrates cultural, biological, and chemical techniques to
manage pests.
Larva Maggot; juvenile stage of fly development; plural:
larvae.
Nu-lure® A commercial formulation of corn protein that
acts as a broad-spectrum food attractant for male and fe
male fruit flies.
Ovipositor Egg-laying tube.
Parapheromone lure Mild to very strong attractants that
attract only male fruit flies; many are produced by plants.
Persistent Relates to how long-lasting a lure is.
Pre-oviposition period Time period after adults emerge,
before egglaying begins.
Protein hydrolysate Extracts of yeasts or grains that act
as a broad-spectrum food attractant for male and female
fruit flies (and many other protein-feeding insects).
Pupa The transformation stage of fly development, after
larva and before adult; a hard, brittle case covers the pupa;
plural: pupae; pupation: the act of transformation.
Sheath Outer covering of ovipositor.
Staley’s® Fly Bait No. 7 A commercial formulation of corn
protein that acts as a broad-spectrum food attractant to male
and female fruit flies.
Thorax Back or top of the mid-body.
Torula® yeast tablets A commercial formulation of yeast
protein that acts as a broad-spectrum food attractant for
male and female fruit flies.
Volatile Readily vaporized; refers to lures, affects how well
they can be carried on the wind.
Commercial traps
• Protein bait—glass or plastic McPhail traps can be used;
flies enter from below and cannot get out.
• Lure—the waxed cardboard Jackson trap, or tent trap, is
popular; it has a removable, sticky insert floor to catch
flies and a cotton wick for the lure.
• Yellow sticky board—rectangular, yellow, sticky boards
are used with or without other attractants.
Home-made
• Protein bait—use a clear plastic bottle with several 1
inch holes; add a liquid bait mix.
• Lure—use a clear plastic bottle with a few 1⁄4-inch holes;
put cotton inside to absorb the lure.
• Harris trap—a tall container with a clear, wide cover and
1-inch diameter holes; can be used with any attrac
tant; easier to use than sticky traps, but when used with
lures, it must have insecticide to kill the flies before
they escape.
• Sticky panels—paint cardboard or wood panels bright
yellow; cover with Tanglefoot®.
Placement of traps
The location and placement of monitoring traps may be
more critical for medflies than other fruit flies. Research
has shown that medflies can effectively be trapped in their
mating areas, such as the upwind side of crowns of trees
receiving some light. Traps for the other fruit flies should
be placed in their resting or feeding areas.
Protein traps and other mild attractants should be
placed in a shady area close to the host plants. Lure traps
should be placed at the borders, corners, and outside of the
field before flies move into the field. Color attractants
should be placed in the open for best effectiveness.
Trap density (number per area) and spacing depends
on the type and amount of attractant used. Traps for moni
toring do not need to cover the entire area evenly. Protein
bait traps have been used at 15–30 ft in-field spacing, and
7
IP-4
References
Liquido, N. 1993. Reduction of oriental fruitfly (Diptera:
Tephritidae) populations in papaya orchards by field
sanitation. J. Agric. Entomol.10(2):163–170.
Liquido, N., E. Harris, and L. Dekker. 1994. Ecology of
Bactrocera latifrons (Diptera: Tephritidae) populations:
host plants. natural enemies. distribution. and abun
dance. Ann. Entomol. Soc. Am. 87(1):71–84.
Mau, R. F. L. 1983. Watermelon insecticide guide for com
mercial producers. Univ. of Hawaii, HITAHR Brief No.
044.
Robinson, A. S., and G. Hooper (eds). 1989. World crop
pests: Fruit flies—their biology, natural enemies and
control. vol. 3A and 3B.
Steiner, L., W. C. Mitchell, and K. Ohinata. 1959. USDA
recommends poisoned-bait sprays for fruit flies. Hawaii
Agriculture. March, 1959. p. 25–30.
United States Department of Agriculture, Animal and Plant
Health Inspection Service. Emergency programs manu
als. Mediterranean fruit fly action plan (l982); Melonfly
action plan (1984); Oriental fruit fly action plan (1989);
Malaysian fruit fly action plan (1993).
A list of additional references is available upon request.
Trap sources
Great Lakes IPM, 10220 Church Rd., NE, Vestaburg, MI
48891.
Pest Management Supply Inc., 311 River Drive, MA 01035.
Information and assistance for the development of the first
edition of this publication, HITAHR Brief no. 114 (1995), was
provided by Deborah Ward, Terry Sekioka, Vince Jones, and
Ken Kaneshiro of UH-Manoa; Robert Boesch, Lance
Kobashigawa, and Pat Conant of the Hawaii Dept. of Agri
culture; and Norman Makio, Tane Datta, Michael Rassa, Joe
Rosenova, Bart Jones, Kert Hamamoto, Jack Banks, and Jim
Frazier. Other support was received from the LISA for Ha
waii Project, the Hawaii County Research and Development
Department, the Hawaii Fruit Fly Committee, and the Big
Island Resource, Conservation and Development Council. The
present revision was prepared by Russell Messing and the
staff of the CTAHR Publications and Information Office.
Mention of a trademark, company, or proprietary name does
not constitute an endorsement, guarantee, or warranty by the
University of Hawaii Cooperative Extension Service or its
employees and does not imply recommendation to the exclu
sion of other suitable products or companies.
Caution: Pesticide use is governed by state and federal
regulations. Read the pesticide label to ensure that the
intended use is included on it, and follow all label direc
tions.
This and other publications of the College of Tropical Agriculture and Human Resources, University of Hawaii at Manoa, can be found on the
Web site <http://www2.ctahr.hawaii.edu/oc/> or ordered by calling 808-956-7046 or sending e-mail to [email protected].
8
Insect Pests
July 2000
IP-5
Destructive Turf Caterpillars in Hawaii
Jay Deputy1 and Arnold Hara2
Departments of Tropical Plant and Soil Science and 2Plant and Environmental Protection Sciences
1
T
he most common insect pests of turfgrasses in
Hawaii are “lawn caterpillars,” the larvae of lepi
dopterous insects (moths and butterflies). Major pests
in this group are three moths and one butterfly. The moths
are the grass webworm (Herpeto
gramma licarsisalis Walker), the lawn
armyworm (Spodoptera mauritia
acronyctoides Guenee), and several
species of cutworm including the black
cutworm (Agrotis ipsilon Hufnagel).
The butterfly is the fiery skipper
(Hylephila phyleus Drury). All of
Hawaii’s turfgrasses are susceptible to
attack by these four pests, although
some of these insects prefer a particu
lar type of turf. The grass webworm
does the most damage and is therefore
the most important turf pest in Hawaii,
and the lawn armyworm also causes ex
tensive injury. Serious outbreaks of
damage by the black cutworm and fi
ery skipper occur less frequently.
hatch into the stage called the larva or, more commonly,
the caterpillar. The larva is the feeding stage of the in
sect that causes all of the damage to turf, and it is the
primary target of a pest management program. The larva
goes through several developmental
stages called instars, and its size, color,
and markings may change drastically.
This part of the life cycle lasts for sev
eral weeks to a month or more, depend
ing mainly on the temperature. As the
last instar of the caterpillar matures, it
burrows into the soil and enters the
stage called the pupa. The pupa is tor
pedo-shaped and, in moths, is sur
rounded by a “cocoon” of silk and de
bris. The pupal stage is difficult to kill
with pesticides because it does not feed
and is not likely to come in contact with
pesticide sprays. The pupating insect
will undergo developmental changes
lasting for several weeks to several
months, depending on the temperature,
Grass webworm feeding.
before emerging as the adult moth or
The insect life cycle
butterfly. In Hawaii, the life cycles of
All moths and butterflies have a similar four-stage life
these insects are accelerated because of consistently warm
cycle. The adult is the familiar moth or butterfly, the
temperatures. They are capable of completing at least five
reproductive stage of the life cycle. The adult insects
or six life cycles per year.
mate and, depending on the species, the female lays eggs
The insects can be most easily identified by the ap
either singly over a period of several weeks or once or
pearance of the adult moth or butterfly or the larval cat
erpillar. The feeding habits of the caterpillars of each spe
twice in masses of several hundred eggs. The eggs take
cies are also characteristic, and preliminary identifica
from two to five days to hatch, depending on the spe
tion of the pest is often made by observing their damage.
cies and, more importantly, on the temperature. The eggs
30, 1914, in cooperation with the U.S. Department of Agriculture. H. Michael Harrington, Interim Director/Dean, Cooperative Extension Service/CTAHR, University
CTAHR publications can be found on the Web site <http://www2.ctahr.hawaii.edu> or ordered by calling 808-956-7046 or sending e-mail to [email protected].
IP-5
Turf Caterpillars
CTAHR — July 2000
Grass webworm caterpillar, about 1 inch long when fully grown.
Grass webworm pupae; the lower one is a later pupal stage.
Eggs of grass webworm on the upper side of a grass blade.
Grass webworm
The grass webworm, the major turf pest in Hawaii, is
thought to have come from Southeast Asia. It was first
found on Oahu in 1967 and has spread to all of the ma
jor islands. It prefers bermudagrass lawns and kikuyu
grass pastures but has 13 other host grasses including
centipedegrass and St. Augustinegrass. Grass webworm
feeding injury spreads more rapidly on fine textured
grasses than coarse textured ones. Common bermuda
grass and ‘Tifway’ bermudagrass are more resistant to
its infestation than the other host turfs. The grass web
worm is not a serious problem on zoysiagrass lawns in
Hawaii.
Its eggs are laid in small groups on the upper sur
face of leaves and stems, along the midrib near the base
of the blade. They are flat and elliptical and are laid sin
gly or in masses overlapping each other like shingles.
Just before hatching, the black head of the larva is vis
ible through the eggshell. Egg development ranges from
2
4 to 6 days, and hatching takes place at night. Grass
webworm eggs have been collected on grasses up to 4000
feet in elevation.
The grass webworm caterpillar develops through
five larval instars. The newly hatched first instar is about
1
⁄16 inch long, translucent with a black head capsule. It is
amber colored until feeding begins, when it changes to
light green as a result of the ingested plant material. The
other four instars have darker brown head capsules, and
pairs of small dark brown spots extend along the back
of the body. When fully grown, the grass webworm is
slender and about 1 inch long.
The first and second instar caterpillars begin to feed
on upper leaf surfaces, leaving the lower surface intact.
Their initial feeding produces grass blades with ragged
edges, the first visible sign of grass webworm infesta
tion. The third, fourth, and fifth instar caterpillars notch
leaf edges, eat entire leaves, and spin large quantities of
IP-5
Turf Caterpillars
CTAHR — July 2000
Two views of the adult, moth stage of the grass webworm; both views are about twice life size.
silk webbing. Feeding in these later instars occurs on
stems, leaves, and crowns, extending to ground level
and resulting in irregular brown patches in the turf. The
caterpillars can be found at the edges of the feeding area,
and they leave characteristic webbing and fecal pellets
throughout the area. All stages feed at night and hide
curled up in thread-lined tunnels in the turf thatch dur
ing the day. When disturbed, the caterpillar becomes
active and rapidly moves away.
These caterpillars prefer sunny areas and are often
found on south-facing or steep slopes where conditions
are hot and dry. The damage caused by the webworm is
often mistaken for drought stress, and the resulting turf
thinning is often accompanied by weed infestation. The
caterpillar reaches maturity in about 14 days.
Prior to pupation, the fifth instar larva becomes qui
escent and slightly shorter in length, and it burrows into
the soil to form a reddish-brown pupae about 1⁄2 inch
long. Pupation and development usually take 6–7 days
before the adult moth emerges.
The grass webworm moth has a wingspan of about
3
⁄4 inch when at rest with its wings spread in a triangular
shape. The body is approximately 1⁄2 inch long and var
ies from uniformly light to dark brown, with small black
dots scattered over the wings. The moths are gregarious
and often are found clustered on vegetation. They are
attracted to light and may be a nuisance around the home
when their populations are high. They are active at night
and rest during the day, when they often can be found
on flat surfaces in or near grass areas. The moth emerges
from the pupal stage at night, and mating generally oc
curs that first night. Three to six days after mating, the
female lays from 250 to 500 eggs over a period of five
to seven nights. The adult moth has a life span of about
13 days.
3
IP-5
Turf Caterpillars
CTAHR — July 2000
Eggs of the lawn armyworm: egg mass (left) and numerous emerging first-instar caterpilars (right).
Lawn armyworm
The lawn armyworm is a native of Southeast Asia, IndoAustralia, and the South Pacific. It was first recorded on
Oahu in 1953 but apparently arrived well before that
time. It is now found on all major Hawaiian islands.
Hawaii’s particular species of armyworm is apparently
not found anywhere else in the USA. During the 1960s
it was considered Hawaii’s most severe lawn pest, but
recently its populations have stabilized, possibly held
in check by various parasites and predators.
In Hawaii the lawn armyworm is a serious problem
mainly on bermudagrass lawns, but it will also feed on
sedges, sugarcane seedlings, seashore paspalum, and
zoysiagrass. Severe damage to lawns is characterized
by a completely denuded circular area sharply defined
by a front of undamaged turf. With heavy populations
of actively feeding larvae, this destruction may advance
4
about 1 foot each night.
The eggs are laid in masses of 600–700 eggs that
are covered with long, light brown hairs from the abdo
men of the female. These felt-like egg masses are ce
mented to leaves of trees and shrubs or on buildings
close to lights. They are often found on eaves and open
lanai ceilings. Brushing the egg masses off helps to
physically control the insect. The eggs hatch in three to
five days.
The larva of the lawn armyworm has seven to eight
instars. The first instar upon hatching is a tiny, green
caterpillar about 1⁄16 inch long, which spins a silken thread
to reach the ground and begin feeding on grass blades.
The caterpillars tend to remain in the same area and feed
together, forming the characteristic steadily increasing
circle of destruction. Close examination of armyworm
IP-5
Turf Caterpillars
CTAHR — July 2000
Two views of the lawn armyworm caterpillar (life size is about 11⁄2 inches); the one at right in its later-stage coloration.
Two views of the moth (adult) stage of the lawn armyworm, about twice life size; wings open (left) and wings folded (right).
infested stands of turf will reveal clipped or skeleton
ized grass blades mingled with green fecal pellets. Lar
vae will be found feeding near the edges of the dam
aged area. Occasionally, large numbers of armyworms
will develop in one area, then migrate to another after
exhausting their food supply.
As they grow, the caterpillars become brownish, with
a pair of pale stripes down the length of their backs.
They reach a mature length of approximately 11⁄2 inches
in about 28 days. The young caterpillars feed on the grass
during the night and day, but older and larger ones feed
only at night and hide in the thatch during the day. When
disturbed, the caterpillars will become active and jump
around rapidly.
The mature final-instar caterpillar burrows into the
soil and forms a hardened, reddish-brown casing (pupa)
around itself. The average length of the pupa is 5⁄8 inch.
The caterpillar pupates in the soil and emerges as an
adult moth in 10–14 days. The total life cycle can be
completed in about 43 days in Hawaii, with about 8 gen
erations per year.
The adult lawn armyworm is a grayish-brown, thick
bodied moth with a wingspread of about 11⁄2 inches. The
forewings are marked with several dark lines and a con
spicuous black spot. The moth emerges from the pupa
and mates within one day. The female begins laying eggs
about four days later. The adult moth lives for about 12
days. The females fly at night and are attracted to lights,
often laying their eggs near one. Populations of lawn
armyworms may be locally controlled by reducing night
lighting adjacent to sensitive turf areas or by using yel
low light bulbs, which are less attractive to the moth.
5
IP-5
Turf Caterpillars
The variegated cutworm caterpillar is similar
in size and appearance to the black cutworm;
actual size is 11⁄2–2 inches long.
CTAHR — July 2000
Moth stage of the variegated cutworm.
Cutworms
Cutworms, including the black cutworm or some closely
related species, are found in practically every part of the
world. “Cutworm” is the common term for the larval
stage (caterpillar) of various moths of the genus Agrotis
and other related genera of the family Noctuidae. They
feed on many plants including trees, turfgrass, rice and
other cereals, and the seedlings of tomato and crucifers,
cutting off stems, buds, and young leaves. They feed at
night and burrow into the soil during the day.
Black cutworm eggs are laid singly or in small clus
ters. The female prefers to deposit her eggs on curly
dock and mustard plants. One method of control of black
cutworm is to eliminate or reduce these broadleaf weeds.
The eggs hatch in 3–6 days.
6
The larva is brownish on top with a broad, pale gray
band along the midline. It has gray-green sides with lat
eral, blackish stripes. The head capsule is brownish-black
with two white spots. The mature instar is a plump, black
ish caterpillar 11⁄2–2 inches long. The caterpillars remain
in shallow holes during the day and curl up when dis
turbed. They emerge at night to feed on grass blades
and stems of young seedlings, shearing them off at
ground level. The feeding causes browning in turf. Cut
worms are solitary feeders, and their infestations are
usually in much smaller numbers than infestations of
lawn armyworms or grass webworms, and cutworm
damage, therefore, is usually not as serious. However, a
small population of black cutworms can devastate a
IP-5
Turf Caterpillars
CTAHR — July 2000
The moth caterpillars compared
Actual-size comparison of caterpillars of the black cutworm
(top), lawn armyworm (middle), and grass webworm (bottom).
These late instars are approaching full size. The ruler is in
millimeters; the bar is 1 inch long.
newly emerged bed of flower or vegetable seedlings in
a very short time.
The caterpillar lives for 28–34 days before boring
into the soil to form the pupa, which is dark brown and
about 3⁄4 inch long with a posterior spine. The pupal stage
lasts 10–14 days before the adult moth emerges.
Black cutworm adults are large, thick-bodied, noc
turnal moths with a wingspan of 11⁄2–2 inches. The body
is gray and the forewings are gray with dark brownish
black markings. The hindwings are almost all white but
have a dark fringe. The moths mate within two to four
days after emergence. The female lays 1200–1600 eggs,
singly or a few together, over a 5–10 day period. The
adult moths may live for up to 30 days.
7
IP-5
Turf Caterpillars
Lawn damage from the fiery skipper.
CTAHR — July 2000
The fiery skipper lays its
eggs singly on a blade of
grass.
The fiery skipper caterpillar has a prominent head due to its
narrow neck.
Fiery skipper
The fiery skipper is a butterfly that is active during the
day and is almost always found in open lawns, gardens,
and fields in populated suburbs rather than in undisturbed
rural areas. This butterfly has the rapid, skipping flight
common in the insect family Hesperiidae. The fiery skip
per caterpillar prefers bermudagrasses. The larvae de
velop more slowly on zoysiagrasses and centipedegrass
and are seldom seen on St. Augustinegrass.
Fiery skipper eggs are laid singly on the undersurface
of grass leaves and stems. They hatch 2–3 days after
being laid.
8
The larva has distinctive, reddish markings on the
front of its oversized, black head. It has a narrow neck,
followed by a dark thoracic shield and a greenish-pink
body with a granulated texture. The caterpillars spin silk
shelters in the thatch and are not readily seen unless
flushed out by a pyrethrin or detergent test. The average
length of time to complete the larval stage is approxi
mately 16 days. Fiery skipper damage in turf is a 1–2
inch round spot from which all the grass has been eaten
by a single larva. If there is a large population, these
spots will combine into larger dead patches. Damage
IP-5
Turf Caterpillars
This fiery skipper pupa is surrounded by thatch debris.
usually appears on turf located near flowerbeds, where
the adult skippers feed.
The mature fiery skipper caterpillar burrows into
the soil to form a pupa that is light yellow but otherwise
similar in appearance to the pupa of the grass webworm
or the black cutworm. The adult butterfly emerges in 7–
10 days.
The adult has a wingspan of 11⁄4–11⁄2 inch. The wings
are orange-brown. The outer margins of the male’s wings
are black and toothed above, the forewing has a wide
black stigma, and the underside of the hindwing is scat
CTAHR — July 2000
The fiery skipper butterfly is found mostly in open suburban
or urban landscapes.
tered with small black spots. The upper side of the fe
male is dark brown with a very irregular orange band,
and the underside of the hindwing is pale brown with
paler checks. Skippers are distinguished from other but
terflies by having a hooked knob at the end of their very
short antennae. The adult butterflies are strongly attracted
to nectar-producing flowers, such as lantana. The males
pursue the newly emerged females, and mating takes
place within a day. Three to four days later, the female
begins laying 50–150 singly spaced eggs.
9
IP-5
Turf Caterpillars
CTAHR — July 2000
Control measures
Insecticide application is usually the first line of defense
when there is a sudden, widespread increase of defolia
tion by a turfgrass pest. There are few alternatives but
to depend upon a recommended chemical or biological
insecticide. Numerous insecticides for application to
turfgrasses have been registered with the EPA and the
Hawaii Department of Agriculture for use in Hawaii
(Table 1, pp. 12–13). Pesticide registrations are con
stantly changing and are often different from state to
state. Always read the label before applying a pesticide
to be sure that the intended use is stated on it.
Before taking any control measures, identify the pest
that is present and estimate how many caterpillars are
feeding in a given area. If caterpillars are not readily
seen, flush them out with soapy water or a solution of
pyrethrin insecticide poured over the area where activ
ity is suspected. Use 1–2 oz of dish soap or 1⁄2 oz of a
pyrethrin-containing insecticide in about 2 gallons of
water. Soak an area of about 1 square yard with the mix
ture and wait for a few minutes. The caterpillars will be
irritated by the solution and come to the surface. Col
lect the caterpillars in a can and count them after you
are sure there are no more hidden in the thatch (after 10
minutes). Repeat the test in other areas where infesta
tion is suspected. Depending on the caterpillar species
and the population density, treatment with insecticide
may or may not be necessary. Spot treatments in severely
affected areas may be the best approach, or a much larger
treatment may be necessary if the test indicates that the
entire lawn is heavily infested. Treatment is recom
mended if flushing of 1 square yard of turf reveals more
than five or six caterpillars of black cutworm or lawn
armyworm and more than 15 caterpillars of grass web
worm or fiery skipper. These threshold levels are a gen
eral recommendation. Some experts believe that only
four or five caterpillars of the grass webworm are enough
to warrant treatment. Even one larva may be unaccept
able in a highly manicured golf green.
Proper timing of a pesticide application to direct it
against the most vulnerable stage of the turfgrass pest is
necessary for effective control and usually reduces the
number of applications needed for complete control. All
four of Hawaii’s major turfgrass pests described here
are destructive to turf only during the larval (caterpil
lar) stage. All of these pests are more easily controlled
10
in the younger caterpillar stages (the first few instars).
They become much more difficult to eradicate as they
near pupation, and they are resistant to pesticide treat
ment in the pupa stage.
Pesticide application techniques
Before applying an insecticide, the turf should be mowed
and the clippings removed to enhance pesticide penetra
tion into the turf canopy. A thorough irrigation before
application moves insects out of the thatch and soil and
brings them to the surface. For night-feeding larvae
(grass webworm, lawn armyworm, black cutworm),
apply the insecticide in the late afternoon or early
evening. Light irrigation after spraying rinses the insec
ticide off grass blades and into the turf where thatch
active caterpillars reside. A heavier irrigation should
follow granular insecticide applications to wash the gran
ules into the thatch and activate the insecticide. After
this initial post-application irrigation, do not irrigate
again or mow for at least 24 hours. Some biological con
trol agents and newer chemical pesticides may require
special handling and application techniques. Always read
and follow the pesticide label instructions.
Liquid pesticide formulations are mixed with water
and sprayed on the grass. If a compressed-air sprayer is
used, mix the recommended amount of pesticide with 3
gallons of water for every 1000 square feet (sq ft, or ft2)
to be treated. If a watering can is used, mix the recom
mended amount of insecticide with 12 gallons of water
for every 1000 ft2. If using a hose-end sprayer, put the
recommended amount in the jar and follow the direc
tions for the particular model of sprayer. Wettable pow
der formulations are mixed in the same manner but the
solution must be shaken frequently during application.
Granular formulations should be applied with a mechani
cal spreader and watered in well. Apply these insecti
cides only to the lawn area, avoiding all other plants or
ornamentals. Observe and follow label directions for
reentry to treated areas. Allow several days for the full
effect of the treatment to take place.
Types of insecticides
Systemic insecticides are absorbed and translocated
through the plant. They may be applied as a liquid spray
or root drench or granular soil application. Foliar spray
IP-5
Turf Caterpillars
applications usually result in poor systemic activity.
Systemic insecticides usually have a long-lasting residual
effect. The usual mode of action for this type of insecti
cide is by affecting the nervous system of the insect af
ter it is ingested when plant sap is sucked or leaves are
eaten.
Contact insecticides are not absorbed by the plant
and are effective only if they make direct contact with
the insect, or if the insect eats the treated leaves or comes
into contact with the residue before the insecticide is
washed off by rain or overhead irrigation. Many of these
insecticides contain additives (“stickers”) that improve
adherence to the plant and provide a certain amount of
resistance to being washed off. However, contact insec
ticides are usually not as long lasting as systemic types.
A contact insecticide is usually applied as a foliar spray
and affects the nervous system of the insect upon direct
contact or ingestion.
Insecticides sometimes damage plants due to phyto
toxicity. It is advisable to test any product on a small
scale before making large-scale applications. Spray ac
cording to label directions, and spray again a week later
(unless the label prohibits such a frequency). Allow 5–
7 days for symptoms to appear; for systemic insecti
cides, allow 14–21 days for symptoms to appear.
Insect growth regulators (IGRs) are normally ap
plied in a foliar spray. Some may have a limited absorp
tion and translocation in the plant and therefore exhibit
a local systemic type of action. Others enter the insect
by contact. The mode of action for IGRs is by interfer
ence with the metamorphosis and adult development of
the insect.
Microbial control options in Hawaii are limited. One
that is commercially available is a species of bacterium
(Bacillus thuringiensis, “Bt”) which produces an endo
toxin that is ingested by insects.
Another type of microbial agent is insect-parasitic
nematodes. These soil-inhabiting, microscopic round
worms parasitize caterpillars and certain other insects,
reproduce inside of them, and then emerge to reinfect
other hosts. Insect-parasitic nematodes thrive only in
moist environments where they will not dry out. They
are therefore effective against soil and boring insect pests
including cutworms, armyworms, webworms, wire
worms, and caterpillars occurring in moist, humid mi
CTAHR — July 2000
croenvironments. Nematodes are not effective against
foliar feeding (sucking or chewing) insects. Several spe
cies of insect-parasitic nematode are in agricultural use.
Steinernema carpocapsae has recently been condition
ally approved for use as a microbial control agent in
Hawaii, and it may soon be commercially available.
Other naturally occurring microbial control agents
include pathogenic bacteria, fungi, and viruses that at
tack the caterpillars.
Cultural control
Remove heavy thatch to eliminate much of the daytime
resting habitat for the nocturnal larvae. However, the
grass webworm can be present in large numbers with
out much thatch cover. Do not promote thatch buildup
with heavy nitrogen fertilization or excessive watering.
Core aerating the soil followed by top-dressing with
organic matter also helps prevent thatch build-up. Avoid
stressing turfgrasses by overmowing or underwatering.
The lawn armyworm tends to lay eggs in damp areas
with rank growth, so eliminating such areas helps con
trol this pest.
Biological control
The four turf pests described here have numerous other
natural enemies in Hawaii, including parasites and preda
tors. Parasites include a trichogrammatid wasp, which
attacks the eggs, and paper wasps (vespids) and mud
dabber wasps (sphecids), which feed their young by
stinging caterpillars, then stocking their nest with the
paralyzed caterpillars to serve as future food for the
newly emerged young wasps. Ichneumonid, braconid,
and chalcid wasps are also common parasites of cater
pillars in Hawaii.
A long list of predators includes several species of
ants, carabid beetles, the giant bufo toad, and many spe
cies of birds including the common mynah bird, cattle
egret, Brazilian cardinal, and golden plover, all of which
feed on the caterpillars and the adult moths and butter
flies. Often these control agents naturally occur in suffi
cient numbers to effectively keep populations of turf
insect pests under control.
11
Some common insecticides labeled for use
in Hawaii that are effective against turf
caterpillars
Turf Caterpillars
CTAHR — July 2000
IP-5
The insecticides listed are effective against the larval (caterpillar) stages of the insects and are most effective when applied early
in the development of the larval instars before damage from their feeding becomes too severe.
The accuracy and completeness of this information is not warranted. The products mentioned as examples were licensed for sale
in Hawaii as of January, 2000. Pesticide registrations and allowed uses frequently change. Pesticide users should read the
product label to be sure that the intended use (target pest and site of application) is included on it, and follow all label directions,
precautions, and restrictions. If label information differs from that provided here, follow the label. Listing of products is for information
purposes only and should not be considered a recommendation.
Caution: insecticides may damage certain plants; make a test application on a small area before large-scale application (see p. 11).
*An asterisk by a pesticide name indicates that it is a Restricted-Use Pesticide (RUP) that can be purchased and applied only by
people with appropriate certification from the Hawaii Department of Agriculture.
†
Indicates toxicity to fish and aquatic organisms.
Chemical control agents
Active
ingredient
Chemical
class
Example
product name
spinosad
spinosyns
Conserve SC
bifenthrin
synthetic
pyrethroid
Talstar® GC Flowable Turfgrasses; check label for other registered application sites.
Contact poison. Some products for use by commercial
applicators only.
cyfluthrin
synthetic
pyrethroid
Tempo® 20 WP * †
Turfgrasses; check label for other registered application sites.
For use by commercial applicators only. Cyfluthrin is a contact
poison that is fast to intermediate acting, effective within 1–7 days;
residual effect lasts 3–6 weeks.
lambda-cyhalothrin
synthetic
pyrethroid
Scimitar® GC * †
Contact poison. For use by commercial applicators only.
deltamethrin
synthetic
pyrethroid
DeltaGuard® GC * †
Contact poison. For use by commercial applicators only.
acephate
organophosphate
Orthene®
Systemic and contact poison. Odorous, fast acting, effective within
1–3 days, residual effect lasts 1–2 weeks.
chlorpyrifos
organophosphate
Dursban® 50W *
Contact poison. Odorous, intermediate acting, effective within
3–7 days, residual effect lasts 3–6 weeks.
Many products for use by commercial applicators only.
May be fatal if swallowed or absorbed through skin.
diazinon
organophosphate
Diazinon 4E * †
Contact poison, fast acting, effective within 1–3 days, residual
effect lasts 1–2 weeks. Not for use on golf courses and sod farms.
Some formulations for use by commercial applicators only.
halfenozide
insect growth
regulator (IGR)
Mach 2™
Systemic insect growth regulator that acts as a molt-accelerating
compound (MAC).
12
Comments
Contact poison. May cause phytotoxicity.
Recommended for IPM programs; does not significantly impact
the natural predaceous arthropods including ladybird beetles,
lacewings, minute pirate bugs, and predatory mites.
IP-5
Turf Caterpillars
CTAHR — July 2000
Chemical control agents (continued)
Active
ingredient
Chemical
class
Example
product name
imidacloprid
chloronicotinyl
Merit® 75 WP
Contact and systemic poison. Applied as foliar spray or soil drench.
carbaryl
carbamate
Sevin® *
Contact poison. Intermediate-acting, effective within 3–7 days,
residual effect lasts 3–6 weeks. Some formulations for use by
commercial applicators only.
Organism
Product name
Comments
Bacillus thuringiensis (Bt)
(bacterium)
DiPel® 2X
Insect stomach poison (through production of spores and
endotoxins). Only effective on early-instar larvae; repeat
applications may be necessary. Breaks down rapidly in sunlight
and washes readily off leaves.
Organism
Product name
Comments
Steinernema carpocapsae
(nematode)
Millenium Biological
Insect Control
An insect parasite recently labeled for use in Hawaii.
Comments
Microbial control agents
Biological control agents
Mention of a trademark, company, or proprietary name does not constitute an endorsement, guarantee, or warranty by the University
of Hawaii Cooperative Extension Service or its employees and does not imply recommendation to the exclusion of other suitable
products or companies. Caution: Pesticide use is governed by state and federal regulations. Read the pesticide label to ensure that the
intended use is included on it, and follow all label directions.
13
IP-5
Turf Caterpillars
References
Lai, P.-Y., and C.Y. Funasaki. 1986. List of beneficial
organisms purposely introduced and released for bio
logical control in Hawaii: 1830–1985. Hawaii Dept.
of Agriculture, Division of Plant Industry, Plant Pest
Control Branch. Honolulu.
Lai, P.-Y., and C.Y. Funasaki. 1990. List of beneficial
organisms purposely introduced and released for bio
logical control in Hawaii: Addendum I: 1985–1990.,
Hawaii Dept of Agriculture, Division of Plant Indus
try, Plant Pest Control Branch. Honolulu.
Marsdan, David A. 1979. Turf caterpillars. CTAHR In
sect Pest Series no. 12. 7 pp.
Mitchell, Wallace. Pest management guidelines. Univ.
of Hawaii, unpublished manuscript.
Murdoch, C.L., H. Tashiro, J.W. Tavares, and W.C.
Mitchell. 1990. Economic damage and host prefer
ences of lepidopterous pests of major warm season
turfgrasses of Hawaii. Proceedings of the Hawaiian
Entomological Society 30:63–70.
UC pest management guidelines. 1997. <http://
www.ipm. ucdavis.edu/PMG/r785300811.html>.
A color version of this publication can be viewed on the Web site of the
College of Tropical Agriculture and Human Resources,
<http://www2.ctahr.hawaii.edu/oc/freepubs>
14
CTAHR — July 2000
Insect Pests
March 2001
IP-6
Root Mealybugs
of Quarantine Significance in Hawaii
Arnold H. Hara, Ruth Y. Niino-DuPonte, and Christopher M. Jacobsen
Department of Plant and Environmental Protection Sciences
S
even species of root mealybug are found in Hawaii,
and three of them are of quarantine significance.
These root mealybugs are a serious problem for Hawaii’s
export potted-plant industry because root infestations
are not easily detected unless the plants are removed
from their pots. Potted palms and other slow growing
plants are more susceptible to infestation by root mea
lybugs because they require lengthy bench time to at
tain marketable size.
Damage caused by root mealybugs is not specific.
The most common plant symptoms are slow growth,
lack of vigor, and subsequent death. Unless the infesta
tion is unusually heavy, it is not evident until the plant’s
pot is removed and the root ball is examined. A white,
waxy substance and adult female mealybugs will be
noticeable, especially between the pot and the root ball.
Plants that are growing slowly, root-bound, or under
environmental or nutritional stress are more susceptible
to root mealybug infestation.
Due to their cryptic habit (preference for dark, hid
den places), little is known about root mealybug biol
ogy. In general, depending on the species, the adult fe
males (Figures 1–3) live from 27 to 57 days. White, cot
tony masses containing egg-laying females and/or eggs
Mealybug Quarantine Pests
•
The rhizoecus root mealybugz, Rhizoecus hibisci
Kawai & Takagi (Figure 1), was discovered in Ha
waii in 1992 and has since spread to the state’s major
potted foliage plant production areas. This mealybug
has been found on palms, calathea, Serrisa spp., and
‘Tifdwarf’ bermudagrass.
•
The coffee root mealybug, Geococcus coffeae
Green, was discovered prior to 1908. It has a very wide
host range, including aglaonema, citrus, cacao, cof
fee, croton, cyperus, dieffenbachia, ferns, mango, ole
ander, palms, philodendron, pineapple, schefflera, and
syngonium.
• The pineapple mealybug, Dysmicoccus brevipes
Cockerelly (Figure 2), was first mentioned as occur
ring in Hawaii in 1910. It can be found on the lower
stem or stalks and exposed roots of pineapple and other
bromeliads, as well as on coffee, banana, caladium,
sugarcane, canna, citrus, eggplant, and palms.
The rhizoecus root mealybug is widely distrib
uted in East and Southeast Asia and has also been
found in Puerto Rico and Florida. The coffee root mea
lybug occurs throughout the tropics and subtropics,
including Central America, South America, Africa,
Micronesia, India, Sri Lanka, Philippines, and Florida.
The pineapple mealybug is found in South America,
Africa, Jamaica, Madagascar, the Dominican Repub
lic, Florida, Louisiana, and Massachusetts.
The adult female rhizoecus root mealybug is
snow-white and has an elongated oval shape up to
about 2.35 mm long. The adult female coffee root
mealybug is also a snow-white, elongated oval shape
varying from 2 to 2.5 mm in length; it can be distin
guished from other root mealybug species by the pres
ence of anal hooks, which are prominent, stiff, up
turned spines at the tip of each anal lobe. The adult
female pineapple mealybug is pale pink or white,
broadly oval, and approximately 3 mm long.
z
Although six species of Rhizoecus are known in Hawaii, in this publication we use the common name, rhizoecus root mealybug, to refer
only to R. hibisci. yFormerly called Pseudococcus brevipes Cockerell.
IP- 6
Root Mealybugs of Quarantine Significance
CTAHR — March 2001
2
1
Rhizoecus root mealybugs on palm roots.
Pineapple mealybugs on palm roots.
are normally visible on the outside of the root mass when
an infested plant is lifted from its container. Males of
the three species discussed here have not been observed
in Hawaii, although male pineapple mealybugs have
been collected in Madagascar, Martinique, and the
Domican Republic. The newly hatched, immature crawl
ers (nymphs) are the dispersal stage and are highly mo
bile. Once crawlers find a suitable site, they settle down
and begin to feed on roots with their sucking mouth
parts. The entire life cycle of a root mealybug ranges
from one to four months, depending on the species, cli
matic conditions, and availability of a food source.
Root mealybugs can be spread by irrigation water,
re-use of previously infested pots, re-use of contami
nated media, and crawlers moving from infested plants
to other plants. Infestation of greenhouse bench plants
by root mealybugs can occur by introducing nursery
stock that was already infested when purchased or from
crawlers that move in from host plants near the green
house.
Pest management
Biological control
Adult
27–57 days
Life cycle of a
root mealybug
Hawaii has no known natural predators or parasites that
are specific to rhizoecus and coffee root mealybugs.
Natural predators and parasites of the pineapple mealy
bug in Hawaii include several encrytid (parasitoid) wasps
and lady beetles.
Cultural control
Because root mealybugs are very difficult to detect and
control, every effort should be made to prevent their
spread and establishment. The following practices are
recommended:
• Inspect roots of newly purchased plants by removing
them from their pots.
• Inspect roots of suspected plants, especially slow
growing ones.
• Avoid pot-bound plants by re-potting when necessary.
Nymphal
stages
Crawler
(dispersal
stage)
Egg
< 24 hours
Drawing by James Baker, NCSU; photos by Julie Ann Yogi-Chun and A. Hara.
2
IP- 6
Egg
Nymph
3
Adult
4
Life stages of the rhizoecus root mealybug.
Palm roots in the pot not treated with copper hydroxide
(right) are more compacted and infested with mealybugs.
• Use
Chemical control
Pineapple mealybug populations are tended by sev
eral species of ants, and ant-control measures (physical
barriers, ant bait or spray) help prevent serious mealy
bug infestations.
Chemical control of root mealybugs requires saturation
of the root ball and potting medium to a degree that al
lows the pesticide to penetrate the pests’ white, waxy
secretion. Research has demonstrated that dipping or
drenching with liquid insecticide is more effective than
applying a granular formulation. Dursban® Turf & Nurs
ery Product applied twice as a drench or dip at two-week
intervals controls coffee root mealybug; however, it may
take four to six months before the cottony, waxy secre
tions deteriorate completely, making it difficult to de
termine treatment efficacy. This may pose a potential
risk of shipment rejection by quarantine inspectors.
Research trials ranked Dursban ® 50 WP and
Dursban® Turf & Nursery Product as the least phyto
toxic to palms and indicated that watering palms prior
to drenching application significantly reduced phytotox
icity. A small group of plants should be treated at the
recommended rate under the anticipated growing con
ditions and observed for phytotoxic symptoms for at least
14 days before a large number of plants are treated.
In the dip method, research findings indicated that
submerging the plant’s entire root ball without the pot
in a diluted Dursban solution (1 pint per 100 gallons)
for about 30 seconds with slight agitation is nearly twice
as effective as dipping the plant while still in its pot. In
the drench method, after premoistening with irrigation
or rainfall, the diluted Dursban solution is poured into
each potted plant container (without removing pots from
pots with inner coatings of copper hydroxide
(Spinout®), which prevents root matting and thereby
minimizes root mealybug infestations (Figure 4).
• Separate pots from the ground on raised benches or
with plastic film over the soil.
• Do not allow water from infested areas to run onto
clean areas.
• Remove alternate host plants from around the green
house, or control mealybugs on them.
• Use clean pots and soil; if infested, wash pots with
soap and water.
• Keep the growing area clean of plant debris.
Biorational control
CTAHR research has demonstrated that hot-water dips
are as effective as insecticides against mealybugs. Ex
periments showed that submerging potted rhapis palms
in water held at 120°F (49°C) until the internal root ball
temperature reached 115°F (46°C) was 100 percent ef
fective in killing root mealybugs. Only minor phytotox
icity to raphis was observed, a chlorosis (yellowing) of
older leaves. Drenching potted palm roots in hot water
at 120°F for 15 minutes will not only control mealy
bugs but will also eliminate burrowing nematodes.
3
IP- 6
plants) to saturate the soil at a rate of 10–12 fluid ounces
of solution per gallon of container size.
Marathon® 60 WP is applied only as a drench and
can be incorporated with a surfactant or wetting agent
to ensure thorough distribution of solution in the pot
ting medium. Drench rates are determined by plant con
tainer size. Over 95 percent control was observed for up
to 12 weeks in manufacturer’s trials. Residual activity
of Marathon should control most emerging mealybug
nymphs.
Follow safety precautions given on the product la
bels. Used drench solution should be disposed of by
applying it to approved crops and sites in accordance
with the pesticide label directions.
Precautionary statement
Pesticide use is governed by state and federal regula
tions. Read the pesticide label to be sure that the in
tended use is included on it, and follow all label direc
tions. Consult a chemical sales representative, the Ha
waii Department of Agriculture, or the University of
Hawaii Cooperative Extension Service for updated in
formation on available formulations. The pesticide user
is responsible for the proper use, application, storage,
and disposal of the pesticide.
Disclaimer
Mention of a product name does not imply endorsement
or recommendation by the Cooperative Extension Ser
vice, College of Tropical Agriculture and Human Re
sources, University of Hawaii or the United States De
partment of Agriculture and does not imply its recom
mendation to the exclusion of other products that may
be suitable.
4
References
Baker, J.R. (ed.) 1978. Insect and related pests of flowers and foli
age plants. North Carolina Agric. Extension Service, AG-136.
Beardsley, J.W., Jr. 1965. Notes on the pineapple mealybug com
plex, with descriptions of two new species (Homoptera: Pseudo
coccidae). Proc. Hawaiian Entomol. Soc. 14(1):55–68.
Beardsley, J.W., Jr. 1966. Hypogaeic mealybugs of the Hawaiian
Islands (Homoptera: Pseudococcidae). Proc. Hawaiian Entomol.
Soc. 14:151–155.
Beardsley, J.W., T.H. Su, F.L. McEwen, and D. Gerling. 1982. Field
investigations on the interrelationships of the big-headed ant, the
gray pineapple mealybug, and the pineapple mealybug wilt dis
ease in Hawaii. Proc. Hawaiian Entomol. Soc. 24(1):51–68.
Beardsley, J.W., Jr. 1995. Notes on two Rhizoecus species new to
the Hawaiian Islands, with a revised key to Hawaiian hypogaeic
mealybugs (Homoptera: Pseudococcidae:Rhizoecinae). Bishop
Museum Occasional Papers No. 42, pp. 28–29.
Dekle, G.W. 1965. A root mealybug (Geococcus coffeae Green)
(Homoptera: Pseudococcidae). Florida Dept. of Agric., Div. of
Plant Industry, Entomology Circular No. 43.
Hara, A. 1988. Control of the coffee root mealybug in potted plants.
University of Hawaii at Manoa, College of Tropical Agriculture
and Human Resources, Horticulture Digest, No. 86.
Kuitert, L.C. and G.W. Dekle. 1966. Control of root mealybug,
Geococcus coffeae Green. Proc., Florida State Horticultural So
ciety 79:484–488.
Linquist, R.K. 1991. Identification of insects and related pests of
horticultural plants. Ohio Florists’ Association, Columbus.
Merrill, G.B. 1953. A revision of the scale-insects of Florida. State
Plant Board of Florida, Gainesville, Bulletin 1.
Poe, S.L. 1973. Infestation and spread of root mealybugs in con
tainer-grown ornamentals. Institute of Food and Agricultural Sci
ences, University of Florida, Florida Foliage Grower 10(2):1–4.
Snetsinger, R. 1966. Biology and control of a root-feeding mealy
bug on Saintpaulia. J. Econ. Entomol. 59:1077–1078.
Williams, D.J. 1996. Four related species of root mealybugs of the
genus Rhizoecus from east and southeast Asia of importance at
quarantine inspection (Homoptera: Coccoidea: Pseudococcidae).
J. Natural History 30:1391–1403.
Zimmerman, E.C. 1948. Pseudococcus brevipes (Cockerell). In: In
sects of Hawaii; a manual of the insects of the Hawaiian Islands,
including enumeration of the species and notes on their origin,
distribution, hosts, parasites, etc. Volume 5. (Homoptera: Sterno
rhyncha), pp. 189–201.
Insect Pests
April 2001
IP-7*
Hibiscus Erineum Mite
1
Arnold Hara , Dick Tsuda1, James Tavares2, Julie Yogi3, and David Hensley3
Department of Plant and Environmental Protection Sciences, 2Cooperative Extension Service–Kahului, and 3Department of
Tropical Plant and Soil Sciences
1
Common name
Hibiscus erineum mite, hibiscus leaf-crumpling mite
Bumpy growths (galls) and distorted leaves are the re
sult of feeding by the hibiscus erineum mite.
Scientific name
Aceria hibisci (Nalepa)
Hosts
The hibiscus erineum mite seems to prefer the Chinese
red hibiscus (Hibiscus rosa-sinensis L.), but it will also
attack other hibiscus species and hybrids. Like most gall
(plant-feeding) mites, the hibiscus erineum mite’s host
range is narrow and confined primarily to hibiscus spe
cies; however, it has also been recorded on okra, a plant
in the same family.
Distribution
In Hawaii, the mite was first discovered on hibiscus at
Wheeler Air Force Base, Wahiawa, Oahu, in November
1989, and it is now found on all major islands. It has
been collected in other Pacific areas, such as Tonga, Fiji,
and parts of Australia, and it has also appeared in Bra
zil, but its full range of occurrence is unknown.
Damage
Hibiscus erineum mite feeding on plants results in un
sightly leaf, stem, and twig galls. The damage is most
noticeable on the leaves and developing vegetative buds.
The galls are localized growth reactions of the host plant
to the mite feeding. They are rounded, puckered bumps
that form irregular domes on the leaf surface. The galls
vary in size and are frequently connected and crowded
together, giving a lumpy appearance to the leaf surface.
Because active plant growth is necessary for mite es
tablishment, young leaves and buds are most vulner
able to mite infestation. Older, hardened growth will
not develop galls.
Biology
In warm and tropical areas, mites usually develop from
an egg through two nymph stages to the adult stage.
The nymphs resemble the adults but are smaller. The
hibiscus erineum mite develops inside the “pouch” of
the gall. Based on the extensive gall formation that can
occur on hibiscus in a relatively short period of time,
the life cycle of this mite seems to complete itself in
less than three weeks.
The adult hibiscus erineum mite is very small—
invisible to the unaided eye. The mite is soft-bodied and
wormlike, with two body regions: the gnathosoma
(mouthparts), and the idiosoma (remainder of the body).
Hibiscus erineum mites are unique among mites because
they have only two pairs of legs, compared with the
four pairs of other mite species.
*Revised by Arnold Hara from Instant Information no.18, 1996; information on cultivars provided by James Tavares and Marilyn Couture.
IP-7
Hibiscus Erineum Mite
Behavior
Hibiscus erineum mites rely on wind, insects, and birds
to carry them. The adult female is probably the most
mobile in terms of dispersal. Flying insects, especially
those that like the same plants as the mites, are believed
to be the most common means of aiding the movement
of hibiscus erineum mites.
Management
Biological control
Predatory mites are well known biological control agents
of the galling, plant-feeding mites. Predatory mites en
ter the galls and presumably are preying on the hibiscus
erineum mite. When predatory mites are present, gall
ing damage to hibiscus is reduced. Predatory mites can
be seen by the unaided eye and are recognized by their
fast-moving action. If a number of fast-moving mites
are observed on hibiscus with galls, then applying a
miticide is not recommended because it will kill the
predatory mites. Prune to remove severely affected
branches and leaves, and discard them promptly by burn
ing, burial, or dumping them enclosed in a plastic bag.
Cultural control
To prevent the spread of hibiscus erineum mite infesta
tions, avoid taking cuttings from known infested areas,
even from apparently healthy plants. If damage to Chi
nese hibiscus cannot be tolerated, consider replacing the
plant with another hibiscus type less preferred by the
mite, or another type of plant.
Preliminary trials conducted over two years at
CTAHR’s Low Elevation Experimental Farm in
CTAHR — April 2001
Kahului, Maui, indicated that the cultivars ‘Apricot’,
‘Empire’, ‘Pink Hibiscus’, ‘Itsy Bitsy Peach “Monch”’,
‘“Zahm” Chinese’, and ‘Apple Blossom’ are less sus
ceptible to hibiscus erineum mite infestation than ‘Chi
nese Red’, ‘Herman Shierman’, ‘Orange Hibiscus’, ‘Nii
Yellow’, and ‘Kardinal’. Most of these cultivars are suit
able to grow as hedges.
Chemical control
If biological or cultural methods do not control the hi
biscus erineum mite on the plant or in the overall land
scape, then pesticides can be used. Prune all severely
affected branches before applying miticides, and repeat
the miticide applications at least two to three times at
weekly intervals. Repeat applications are necessary be
cause modern pesticides are made not to last in the en
vironment.
Specific recommendation of a miticide is difficult
because of pesticide label restrictions. There are some
suitable pesticides registered for use only by licensed
landscape or nursery professionals. Homeowners may
consider miticides registered for general outdoor orna
mentals or specifically for hibiscus in the landscape. For
information on miticides currently registered for use by
homeowners or commercial growers and landscape
managers, contact your local Cooperative Extension
Service office.
Reference
Carson, Cynthia, and Neil Gough. 2000. Hibiscus
erineum mite. H00054. Queensland Horticulture In
stitute, Department of Primary Industries,
Queensland, Australia.
Mention of a trademark, company, or proprietary name does not constitute an endorsement, guarantee, or warranty by the University
of Hawaii Cooperative Extension Service or its employees and does not imply recommendation to the exclusion of other suitable
products or companies.
Caution: Pesticide use is governed by state and federal regulations. Read the pesticide label to ensure that the intended use is in
cluded on it, and follow all label directions.
2
Insect Pests
May 2001
IP-8
Scouting for Thrips in Orchid Flowers
Robert G. Hollingsworth1, Arnold H. Hara2, and Kelvin T. Sewake3
1
2
U.S. Pacific Basin Agricultural Research Center, USDA-Agricultural Research Service,
CTAHR Department of Plant and Environmental Protection Sciences, 3CTAHR Cooperative Extension Service
T
hrips are the most common insect pest of orchid
flowers in Hawaii. Thrips can be controlled using
appropriate pesticides either in the field or as a post
harvest dip.
Some growers apply insecticides to orchid crops on
a calendar basis, without checking first to see if pests
are present. But not all orchid farms in Hawaii have prob
lems with thrips, and therefore this practice may not be
cost-effective and might result in the eradication of ben
eficial insects that normally keep pests such as white
flies and aphids under control. In general, pesticide ap
plications should be made only when the number of pests
has exceeded a certain tolerance level, or threshold, as
determined by pest scouting.
In deciding on a threshold level for thrips in orchids,
a grower should consider how the crop is to be marketed.
If the flowers are to be sold within the state of Hawaii, a
light thrips infestation (an average of < 1 insect per or
chid spray) can be tolerated, because light infestations
will not damage flowers. However, if the number of thrips
appears to be increasing, growers should consider ap
plying insecticides. Once thrips become well established,
they will be hard to control, because their pupae in the
soil may escape treatment. Successive, carefully timed
insecticide sprays are needed if this occurs.
If the flowers are to be exported from Hawaii, the
grower is responsible for shipping flowers that are free
of thrips and other pests. Otherwise, quarantine inspec
tors may reject flower shipments and the Cut Flower
Compliance Agreement stamp issued by USDA-APHIS
may be revoked. Obviously, growers who export must
adopt a pest tolerance threshold of zero, or else use an
effective treatment after harvesting.
Scouting methods
Three methods can be used for pest scouting; their ad
vantages and disadvantages are summarized in Table 1.
Direct observation
Direct observation of thrips in blossoms is a good, non
destructive method, but it is relatively time-consuming.
Thrips typically hide deep within the blossoms. The lip
Table 1. Comparison of three methods for counting thrips in orchids.
Detection
method
Direct
observation
Efficiency of counting*
Advantages
Disadvantages
Use when goal is
To monitor the
level of the
thrips population
Adults
Nymphs
79%
14%
No equipment required,
consistent results if
same person counts
Time-consuming,
requires good eyesight
Flower
shake
Fast method,
instant results
May damage sprays if
shaken too hard
To detect
thrips
48–93%
4–22%
Berlese
funnel
Produces
consistent results
Time and expense of
funnel construction,
limited amount of plant
material can be processed
To monitor the
level of the
thrips population
34–59%
14–17%
*Efficiency data from Hollingsworth et al. (2000).
IP- 8
of each blossom must be gently pulled down to detect
whether thrips are inside. An experienced person can
generally examine all of the blossoms of one orchid spray
in less than a minute. The person sampling must have
good eyesight and be able to distinguish between thrips
and other small insects commonly encountered, includ
ing other pests, such as aphids, as well as beneficial in
sects, such as parasitic wasps and other predatory in
sects, that feed on thrips. Use of a hand lens is helpful to
distinguish among these insects. In general, thrips adults
will be more easily seen on light colored flowers, while
thrips nymphs (which are light in color) will be more
easily seen against a dark background.
Flower shake
Shaking flowers to dislodge thrips is a second sampling
method that is much faster than visually inspecting in
dividual flowers. A single orchid spray should be shaken
for 5 seconds within a white bucket or plastic bag. The
shake should be of moderate intensity—too vigorous a
shake will bruise or damage the flowers, reducing shelf
life. Research has shown that flower shakes remove from
about half to almost all of the adult thrips present but
less than a quarter of the nymphs (Table 1).
Berlese funnel
Another method for scouting thrips involves putting the
flowers into a specially constructed funnel beneath a
brooder lamp, using the heat from the light source to
drive the thrips down to a collecting jar at the bottom of
the funnel. This sampling device is called a Berlese fun
nel; it can be easily constructed from locally available
materials. Insects collected in the jar can be examined
and counted with the aid of a microscope or hand lens,
or they may be taken to an expert for identification.*
Identification could be important, because there are sev
eral different species of thrips commonly found in or
chid flowers, and certain species may be harder to con
trol with insecticides.
Berlese funnels are particularly good for detecting
the presence of thrips while they are still small, as these
stages would probably be overlooked using the other
two scouting methods.
*For a small fee, insects can be identified by CTAHR’s Agricultural Diagnostic
Service Center (ADSC). Samples can be submitted through any CTAHR Coop
erative Extension Service office. On Hawaii, the ADSC is located at the Komohana
Agricultural Complex in Hilo. Thrips should be submitted in 70% alcohol.
2
CTAHR — May 2001
Some growers have asked if sticky cards can be used
for sampling thrips in orchids. Yellow and blue sticky
cards have been used effectively to sample western
flower thrips in vegetable and flower greenhouses. We
tested a wide variety of card colors (including blue and
yellow) in an orchid shadehouse known to be infested
with western flower thrips, suspending the sticky cards
just above the plant canopy. We had very little success
collecting thrips of any kind. Therefore, we cannot rec
ommend this technique at this time.
How to collect samples
Regardless of the method you choose, you should col
lect the samples evenly throughout any area you are
managing as one unit. The number of samples to collect
depends on your reason for scouting and the level of
thrips present in the crop.
Growers who seldom find thrips may decide to
implement control measures when only a small percent
age of orchid sprays are infested. Those who take this
approach will want to sample a large number of orchid
sprays, and this can be done most efficiently by the
flower shake method. Our research indicates that orchid
sprays infested with adult thrips are randomly distrib
uted in an orchid crop, not clumped together. Using this
information, it is possible to calculate the probability of
detecting adult thrips for a given number of orchid sprays
sampled, provided that the infestation rate and the effi
ciency of the sampling method are specified. This is
shown in the graph in Figure 1, which assumes a 70%
efficiency of counting, such as might be obtained using
the shake method of sampling.
Other growers may have chronic problems with
thrips because thrips are constantly flying in from sur
rounding areas. These growers may want to use scout
ing results to monitor the level of thrips in the crop, in
order to better time pesticide sprays. In such circum
stances, thrips will likely be relatively easy to find us
ing any method. The emphasis should be on using a
method that produces consistent results. Berlese funnel
extractions might be most appropriate, because consis
tent results can be achieved even if different people col
lect the samples. The best way to determine how many
orchids sprays to sample is to compare results obtained
using several different sample sizes. The minimum
sample size that produces consistent results should be
selected.
IP- 8
Figure 1. Probability of detecting adult thrips as a function
of sample size and the percentage of orchid sprays
infested with adult thrips.
Probability of detecting adult thrips
1.2
10%
20%
1
5%
0.8
2%
0.6
1%
o.4
0.2
0
0
20
40
60
80
CTAHR — May 2001
3. With the hole saw bit, cut a hole in the center of the
jar lid. Use Liquid Nails adhesive to glue the lid onto
the spout of the funnel about 1⁄4 inch up from the bottom
of the spout so that the jar can be screwed onto the lid.
4. Bend four pieces of plumber’s tape so that when
evenly spaced around the lamp they will hold the lamp
just above the funnel. Drill 1⁄8-inch holes in the lamp
and rivet the plumber’s tape to the lamp. Adjustments
can be made by bending the plumber’s tape so that the
lamp rests just above the funnel.
5. The funnel cannot stand on the small jar at the bot
tom; therefore, it needs to be supported in a box or
bucket. A frame constructed from wood or galvanized
pipe can be used to support one or more funnels.
100
Number of orchid sprays sampled
Constructing a Berlese funnel
(from Tenbrink et al. 1998)
Materials needed:
• 10-inch
automotive funnel (Balkamp brand, Napa
Auto Parts, part #8211 126)
• 1 square foot of 1⁄4-inch-mesh galvanized hardware
cloth
• 4-ounce jar with screw-on lid, such as a baby food jar
• 10-inch brooder lamp (Woods Wireproducts brand,
Ace Hardware, item #30715)
• 40-watt incandescent light bulb (do not substitute a
bulb brighter than 60 watts)
• 4 pieces of 3⁄4-inch galvanized plumber’s tape, each
41⁄4 inches long
• 8 1⁄8-inch aluminum rivets
• Liquid Nails® adhesive
Tools needed:
• Electric drill with 1⁄8-inch drill bit
• Hole saw bit the same size as the funnel spout diameter
• Rivet gun
• Tin snips
• Pliers
Procedure for construction:
1. Remove the filter screen from the funnel.
2. Cut the hardware cloth to fit and place it in the funnel.
Using the Berlese funnel
Additional supplies needed: a hand lens or magnifying
glass (least 10X) and 70% isopropyl alcohol.
Pour 1–2 fluid ounces of alcohol into the jar. Screw
the jar onto the lid. If you plan to have the thrips identified,
use a mixture of half alcohol, half water, and add a drop of
detergent. This keeps the thrips from getting too stiff.
Harvest enough sprays to yield 50–100 blossoms.
Write down the date, cultivar, and number of sprays used.
Removing blossoms from stems speeds drying, but
handle them gently during removal to prevent thrips es
caping. Put the blossoms into the funnel, place the lamp
on the funnel, and turn on the light. Heat from the bulb
drives the thrips down and they fall into the alcohol.
After 8 or more hours, turn off the light and remove
the jar. Pour the alcohol into a flat dish. Using a hand
lens or magnifying glass, inspect the alcohol for thrips.
If aphids or mealybugs are on the flowers, they will also
be in the jar. Moths and beetles may be attracted to the
light and fall into the funnel. If this occurs, check the fit
of the lamp and adjust the plumber’s tape to minimize
the space between the lamp and the funnel. If the prob
lem continues, seal the space with tape.
Record the number of thrips and divide by the num
ber of sprays. The result of this calculation is the num
ber of thrips per spray. This number, when compared
with the numbers from other surveys, shows whether
the population is rising or falling.
Finally, clean the funnel and the jar. This is impor
tant to avoid contamination of later samples.
Mention of a trademark, company, or proprietary name does not constitute an endorsement, guarantee, or warranty by the University of Hawaii Cooperative Extension
Service or its employees and does not imply recommendation to the exclusion of other suitable products or companies.
3
IP- 8
CTAHR — May 2001
Adult thrips (center) are very small. At left, four of them are seen through a hand lens. At right, there are two within the
circle and another one less visible in the shadows within the blossom (arrow).
Berlese funnels are easily constructed from locally available materials. Multiple funnels can be placed together on a rack.
References
Hara, A.H., T.Y. Hata, V.L. Tenbrink, and B.K.S. Hu. 1995. Postharvest treat
ments against western flower thrips [Frankliniella occidentalis
(Pergande)] and melon thrips (Thrips palmi Karny) on orchids. Ann. Appl.
Biol. 126:403–415.
Hara, A.H., and T.Y. Hata. 1999. Pests and pest management. In: K. Leonhardt
and K. Sewake (eds), Growing dendrobium orchids in Hawaii, produc
tion and pest management guide. College of Tropical Agriculture and
Human Resources, University of Hawaii at Manoa. pp. 29–45
Hata, T.Y., A.H. Hara, and J.D. Hanson. 1991. Feeding preference of melon
thrips on orchids in Hawaii. HortScience 26: 1294–1295.
Hata, T.Y., A.H. Hara, B.K.S. Hu, R.T. Kaneko, and V.L. Tenbrink. 1993.
Field sprays and insecticidal dips after harvest for pest management of
4
Frankliniella occidentalis and Thrips palmi (Thysanoptera: Thripidae)
on orchids. J. Econ. Entomol. 86(5):1483–1489.
Hollingsworth, R.G., A.H. Hara, and K.T. Sewake. 2000. Pesticide use and
grower perceptions of pest problems on ornamental crops in Hawaii. Jour
nal of Extension 38(1). 11 pp. <http://joe.org/joe/2000february/rb1.html>.
Hollingsworth, R.G., K.T. Sewake and J.W. Armstrong. 2000. Scouting meth
ods for detection of thrips (Thysanoptera: Thripidae) on dendrobium or
chids in Hawaii. Manuscript submitted to Journal of Economic Entomol
ogy.
Tenbrink, V.L., A.H. Hara, T.Y. Hata, B.K.S. Hu, and R. Kaneko. 1998. The
Berlese funnel, a tool for monitoring thrips on orchids. CTAHR publica
tion IP-3, College of Tropical Agriculture and Human Resources, Uni
versity of Hawaii at Manoa.
Insect Pests
June 2002
IP-9
Anthurium Thrips Damage to Ornamentals in Hawaii
Arnold H. Hara, Christopher Jacobsen, and Ruth Niino-DuPonte
A
nthurium thrips, Chaetanaphothrips orchidii
(Moulton), (Thysanoptera: Thripidae), formerly
known as the orchid thrips, was first collected in Ha
waii in 1926 and has since become a common pest of
ornamentals. It is a widely distributed species, infesting
greenhouses and outdoor landscapes in the Dominican
Republic, South America, Australia, Japan, Puerto Rico,
India, many European countries, and, within the USA,
it has been reported in Florida, Kentucky, Washington
DC, New York, Louisiana, Illinois, and California, as
well as Hawaii. The anthurium thrips is similar in ap
pearance to two other introduced Chaetanaphothrips
species, the banana rust thrips, C. signipennis (Bagnall),
and C. leeuweni (Karny), that share similar hosts includ
ing banana, ti, and anthurium.
Hosts
While the anthurium thrips shows a preference for an
thuriums, it is a polyphagous feeder, attacking many
other flowers, ornamentals, herbs, fruits, vegetables,
grasses, and weeds. Its host plants include dendrobium
orchid, begonia, bird-of-paradise, bougainvillea, chry
santhemum, night-blooming cereus (Peniocereus greggi),
wandering jew (Tradescantia fluminensis), parsley, cit
rus, sweetpotato, lychee, banana, and corn.
Damage
The appearance of feeding damage caused by anthurium
thrips varies among host plant species. In most cases,
thrips prefer to feed on very young, succulent, imma
ture fruits, flowers, and foliage.
Adult and immature thrips begin feeding within the
unopened anthurium spathe soon after the bud emerges
from the leaf axil. Damage to anthurium appears as white
streaks or scarring on the front and back of the spathe,
deformed spathes, and, with age, bronzing of injured
tissues. Generally, the white streaks and scarring on
spathes caused by anthurium thrips are wider than those
caused by banana rust thrips. In severe cases, anthurium
spathes fail to open, foliage may be deformed with bronz
ing and streaking, and plant growth may be reduced (see
Fig. 1).
Biology
No male anthurium thrips has been observed; reproduc
tion occurs without mating and is continuous through
out the year. The adult uses a sharp ovipositor to deposit
up to 80–100 eggs into a bud or sheath. After 6–9 days,
the eggs hatch into nymphs that are whitish and look
like adult thrips but are smaller and lack wings. The
nymphs crawl and feed on the plant tissues for about a
week, causing damage with their sucking-rasping mouth
parts. Late-stage nymphs are yellow to orange and mi
grate off the host plant to molt into the prepupal stage.
Prepupae look similar to nymphs but have wing pads;
pupae have longer wing pads. Pupation occurs in the
soil or growth medium beneath the host plant, and nei
ther the prepupal nor the pupal stage feeds. In severe
infestations, prepupae can occur in silken cocoons on
the plant. The adult (Fig. 2) emerges from the pupal cells
after approximately 20 days and reinfests the host plants.
It is yellow with banded wings and is about the size of
the period at the end of this sentence (1⁄25 inch). The en
tire life cycle (egg to adult, Fig. 3) is completed in ap
proximately 28–32 days, but it may extend to 3 months,
depending on the temperature. Higher temperature and
humidity and new growth of host plants appear to be
favorable to thrips’ feeding and breeding, leading to
heavier infestations and greater damage during the sum
mer months (Fig. 4).
IP- 9
Control
Anthurium thrips are a serious pest to the anthurium in
dustry. Damage occurs 6–8 weeks before flower har
vest. Feeding by only a few thrips can cause white streaks
on spathes. Since thrips prefer feeding in unopened buds
and unfurled leaves and pupate in the medium or soil
beneath the host plant, they are concealed throughout
most of their life cycle and may be difficult to detect. To
avoid ineffective control measures, it is important to
identify the particular thrips species in an infestation.
Using a hand lens, check in rolled leaves and unfurled
buds, or collect samples to submit for professional di
agnosis and species identification to the Hawaii Depart
ment of Agriculture or to the CTAHR Agricultural Di
agnostic Service Center via any CTAHR Cooperative
Extension Service office.
Biological control
In Hawaii, anthocorid bugs (Orius tristicolor, O. perse
quens, and O. insidiosus) are general thrips predators,
although the extent of their effectiveness against anthu
rium thrips is not documented. Certain lacewings, lady
bird beetles, and predatory mites may also exert some
control on nymph and adult thrips, while ants may prey
on pupae in the soil. Several fungi, including Paecilo
myces species and Verticillium lecanii, have been iso
lated from other thrips species and may infect anthu
rium thrips as well.
Cultural control
Remove infested flowers and foliage from the field or
greenhouse to eliminate existing sources of thrips. Con
trol weeds and grasses, and remove old stock plants that
may serve as hosts to anthurium thrips. Obtain thrips
free propagative material when restocking.
There are no reports of anthurium cultivars that are
resistant or susceptible to anthurium thrips, but injury is
more noticeable on pastel shaded cultivars such as
‘Marian Seefurth’.
Biorational control
A hot-water dip before planting at 120°F (49°C) for 10
minutes can disinfest anthurium propagative material of
thrips. Anthurium cultivars that tolerate hot water treat
ment as rooted plants with leaves include ‘White Lady’,
‘Blushing Bride’, and ‘Kozohara’, while ‘Ozaki’ can-
2
CTAHR — June 2002
Figure 1. Damage to anthurium by anthurium thrips (top
to bottom): leaves; unfurled spathe; front of spathe,
‘Marian Seefurth’ cultivar; back of spathe.
IP- 9
not tolerate hot-water dipping except as whole stem pieces (gobo).
Figure 2. Adult
anthurium thrips.
Chemical control
Because pesticide registrations may
change, consult a chemical sales
representative, the Hawaii Depart
ment of Agriculture, or the CTAHR
Cooperative Extension Service for
information on insecticides cur
rently approved for use against
thrips in anthurium.
Remove infested flowers and
foliage from the field or shadehouse
to allow increased insecticide pen
etration and coverage. Because thrips prefer young,
growing plant tissue, good spray coverage at the base of
plants where spathe development occurs is essential to
contact any exposed thrips. Caution should be used if
applying insecticides on anthurium, because phytotox
icity can occur under hot, dry conditions. Granular con
tact insecticides are effective against the prepupal and
pupal stages of anthurium thrips that occur in the soil,
Figure 3. Life cycle of the anthurium thrips.
Eggs
Nymph I
Nymph II
Adult
(no wing pads)
These stages crawl and feed
Eggs are laid in the unfurled spathe
and young leaf tissue
Prepupa
Pupa
(longer wing pads)
(with wing pads)
Living under the soil or growth medium,
these stages do not feed
CTAHR — June 2002
medium, and plant debris near the base of the host plant,
but no granular insecticide is currently registered for use
in anthurium.
Generally, anthurium thrips populations increase
during the summer and decrease during the winter due
to fluctuations in temperature and rainfall. Consequently,
repeated spray applications may be needed only from
May through August. Depending on the insecticide used,
three to four applications at 2-week intervals may be
necessary to protect newly developed anthurium flow
ers from moderate to severe infestations.
When thrips injury is sustained during the bud stage,
injured anthurium flowers will be harvested for at least a
month following application of an effective insecticide.
References
Anathakrishnan, T.N. 1984. Bioecology of thrips. Indira
Publishing House, MI. pp. 77–78.
Hara, A.H., R.F.L. Mau, D.M. Sato, and B.C. Bushe.
1987. Effect of seasons and insecticides on orchid
thrips injury of anthuriums in Hawaii. HortScience
22(1):77–79.
Hara, A.H., K.T. Sewake, and T.Y. Hata. 1990. Anthu
rium thrips. HITAHR Brief no. 086. College of Tropi
cal Agriculture and Human Resources, University of
Hawaii at Manoa. 1 p.
Hata, T.Y., and A.H. Hara. 1992. Anthurium thrips, Chae
tanaphothrips orchidii (Moulton): biology and insec
ticidal control on Hawaiian anthuriums. Tropical Pest
Management 38(3):230–233.
continued, p. 4
Figure 4. Seasonal fluctuation of thrips injury (%) to an
thurium flowers at Mountain View, Hawaii (Hara et al., 1987).
100
90
80
70
60
50
40
30
20
10
0
CONTROL 1
CONTROL 2
A S O N D
J
F M A M J
J
A
S
Insect drawings from D. Schulz; plant photo from Higaki et al. (see References).
3
IP- 9
Higaki, T., J.S. Lichty, and D. Moniz (eds.). 1994. An
thurium culture in Hawaii. Research Extension Se
ries 152, College of Tropical Agriculture and Human
Resources, University of Hawaii at Manoa.
Jacot-Guillarmod, C.F. 1974. Catalogue of the Thysan
optera of the world. Annals of the Cape Provincial
Museums (Natural History) 7 (Part 3):634.
Pelikan, J. 1954. Remarks on the orchid thrips Chae
tanaphothrips orchidii (M.). Fulia Zoologica et Ento
mologica 3:3–12.
Pinese, B., and R. Piper. 1994. Bananas; insect and mite
management. Queensland Department of Primary
Industries, Australia. 67 pp.
Sakimura, K. 1975. Danothrips trifasciatus, new spe
cies, and collection notes on the Hawaiian species
Danothrips (Thysanoptera: Thripidae). Proc. Hawai
ian Entomol. Soc. 22:125–132.
Schulz, D. ca. 1950. Department of Entomology, Uni
versity of Illinois at Urbana-Champaign. <http://
www.life.uiuc.edu/Entomology/insectgifs/>.
4
CTAHR — June 2002
Insect Pests
June 2002
IP-10
Banana Rust Thrips
Damage to Banana and Ornamentals in Hawaii
Arnold H. Hara1, Ronald F. L. Mau1, Ronald Heu2, Christopher Jacobsen1, and Ruth Niino-DuPonte1
1
CTAHR Department of Plant and Environmental Protection Sciences, 2Hawaii Department of Agriculture
B
anana rust thrips, Chaetanaphothrips signipennis
(Bagnall) (Thysanoptera: Thripidae), was collected
once in 1954 from an outdoor planting of anthurium in
Manoa, Oahu, and was not seen again until 1996, when
it was collected from several commercial nurseries and
farms on the island of Hawaii, after causing severe dam
age to anthurmium, ti, dracaena, and banana.
Banana rust thrips are present in parts of Australia
(Queensland and New South Wales) and Central America
(Honduras, Panama), Brazil, Fiji, Sri Lanka, and India.
They are also established in Florida.
The banana rust thrips is similar in appearance to
two other introduced Chaetanaphothrips species, the an
thurium thrips, C. orchidii (Moulton) (see Hara et al.
2002), and C. leeuweni (Karny), which also share the
same hosts, including banana, ti, and anthurium. Banana
rust thrips can be differentiated from the other two spe
cies by clear differences in body features (specifically,
the presence in females of body hairs and glands that
are visible only with a microscope [Sakimura 1975]).
Hosts
The primary hosts of banana rust thrips are anthurium,
ti, dracaena, and banana. They also infest immature fruits
of orange, tangerine (mandarin), and tomatoes, as well
as green beans.
Damage
The appearance of feeding damage caused by banana
rust thrips varies with the host plant species. In most
cases, thrips prefer to feed on very young, succulent,
immature fruits, flowers, and foliage.
On dracaena and ti (Fig. 1a), thrips can be observed
feeding in the whorls of immature leaves, causing dis
coloration and silvering (characterized by long white
streaks) as well as random squiggles or curlicues near
the petiole end of developed, unfurled leaves. Also, par
ticularly on red ti varieties, the immature leaves may
fail to unfurl and thus appear as deformed leaf whorls
(Fig. 1b).
On anthurium, banana rust thrips damage appears
as white streaks or scarring on the front and back of the
spathe, deformed spathes, and, with age, bronzing of
injured tissues (Fig. 1c). In severe cases, mature anthu
rium spathes fail to open, plant growth may be reduced,
and the foliage may be affected by deformity, bronzing,
and streaking. Damage by banana rust thrips to certain
anthurium cultivars, such as ‘Kalapana’ and ‘Ozaki’, may
appear as curlicues rather than streaks.
On banana, feeding damage is observed on the
pseudostem, but it is the injury to the fruit that signifi
cantly affects marketability (Fig, 2). Thrips feeding in
leaf sheaths results in characteristic dark, V-shaped marks
on the outer surface of leaf petioles. Damaged tissue
becomes bronzed or rust-colored with age. Feeding dam
age to the fruit occurs on fingers soon after the flower
petals dry, initially typified by a water-soaked appear
ance. Young fruits may have dark, smokey-colored ran
dom squiggle or curlicue feeding tracks on the surface.
On mature fruit, oval-shaped, reddish “stains” may be
seen where the fingers touch. Extensive damage may
cover more of the fruit surface with reddish-brown or
black discoloration and superficial cracks. Though un
marketable, such fruits are still edible.
IP- 10
Banana Rust Thrips Damage to Banana and Ornamentals in Hawaii
Biology
Adult banana rust thrips reproduce sexually. After mat
ing, females lay kidney-shaped eggs that are invisible
to the naked eye by depositing them in plant tissues
where the thrips feed. Eggs hatch in 6–9 days; the newly
hatched yellow nymphs feed for a few days before molt
ing into the second nymphal stage, which is yellow or
orange and feeds for a few more days. After 8–10 days,
mature nymphs migrate off the host plant into the soil
or growth medium below and molt into prepupae that
look similar to nymphs but have wing pads. After 2– 5
days, prepupae enter the pupal stage, which has longer
wing pads. Both stages remain in the soil, medium, or
surface debris beneath the host plant and are capable of
crawling but do not feed. In 6–10 days, the adult emerges
from the pupal cells and may remain beneath the sur
face for up to 24 hours before making its way up to rein
fest the host plant.
Adult female banana rust thrips are slender, creamy
yellow to golden brown, and 1⁄16–1⁄25 inch long (about
the thickness of a dime; Fig. 3). Their wings have dark,
eye-like spots at the base and are fringed; when the wings
are folded, the adult appears to have a black line down
its back.
The entire life cycle (egg to adult, Fig. 4) is com
pleted in approximately 28 days, but it may take up to 3
CTAHR — June 2002
months during cooler seasons. Higher temperature and
humidity and new growth of host plants appear to be
favorable to thrips’ feeding and breeding, leading to
heavier infestations and greater damage during the sum
mer months.
Biological control
In Hawaii, anthocorid bugs (Orius tristicolor, O. perse
quens, and O. insidiosus), are general thrips predators,
but the extent of their effectiveness against banana rust
thrips is not known. Some lacewings, ladybird beetles,
and predacious mites may also exert some control on
nymph and adult thrips, while ants may prey on prepupae
and pupae in the soil, growth medium, or surface debris
near the base of the host plant. Several fungi, including
Paecilomyces spp. and Verticillium lecanii, have been
isolated from other thrips species and may infect ba
nana rust thrips as well.
Cultural control
Remove infested flowers and foliage from the field or
shadehouse to eliminate sources of thrips. Discard old
stock plants that may harbor thrips, and obtain thrips
free propagative material for restocking.
There are no reports of resistant or susceptible an
thurium cultivars, although injury is more noticeable on
Figure 1. Feeding damage by banana rust thrips on ti and anthurium: A. Streaks and curlicue markings on opened ti leaf.
B. Deformed leaf whorls on red ti that failed to unfurl. C. Deformed anthurium spathe.
A
2
B
C
IP- 10
Figure 2. Damage to banana fruit by banana rust thrips.
Biorational control
A hot-water dip at 120°F (49°C) for 10 minutes before
planting can disinfest anthurium propagative material
of banana rust thrips. Banana, dracaena, ti, and anthu
rium have all shown potential for heat treatment, al
though cultivar sensitivity has been observed to vary
with season. Tests indicated that some anthurium culti
vars tolerate hot-water treatment as top cuttings with
leaves, including ‘White Lady’, ‘Blushing Bride’, and
‘Kozohara’, while the ‘Ozaki’ cultivar cannot tolerate
the hot-water dip except as whole stem pieces (gobo).
The dracaena cultivar ‘Janet Craig’ was also tolerant of
hot-water treatment. Due to variations among cultivars
and growing conditions, small-scale phytotoxicity tests
should be conducted before a large amount of propaga
tive material is hot-water treated.
Chemical control
Figure 3. Adult
Because pesticide registrations banana rust thrips.
may change, consult a chemical
sales representative, the Hawaii
Department of Agriculture, or the
CTAHR Cooperative Extension
Service for information on insec
ticides currently approved for use
against thrips in a particular crop.
Remove infested flowers and
foliage from the field or green
house to allow increased insecti
cide penetration and coverage.
Growers have reported that banana rust thrips tends to
be more difficult to control than anthurium thrips, pos
sibly due to the former’s pesticide tolerance and greater
reproductive capacity. Growers are advised to consider
insect development of pesticide resistance in devising
their integrated pest management practices.
Generally, thrips populations increase during the
summer and decrease during the winter due to fluctua
tions in temperature and rainfall. Consequently, repeated
spray applications may be needed only from May
Figure 4. Life cycle of the banana rust thrips.
Eggs are laid in leaf
and fruit tissues
Eggs
Nymph I
Nymph II
Adult
(no wing pads)
These stages crawl and feed
pastel shaded cultivars such as ‘Marian Seefurth’.
In banana plantings, covering bunches with poly
ethylene bags during fruit development provides a physi
cal barrier to insect infestations, but bags cannot fully
protect the fruit when a thrips infestation is heavy.
CTAHR — June 2002
Prepupa
Pupa
(longer wing pads)
(with wing pads)
Living under the soil or growth medium,
these stages do not feed
Insect drawings from D. Schulz (see References).
3
IP- 10
through August. Foliar sprays are usually applied two
to three times at 2-week intervals for moderate to se
vere thrips infestations. Since thrips prefer young, grow
ing plant tissue, direct insecticide sprays to the area of
bud development or, in anthurium, to the base of the
plant, where the spathes develop. Use caution when ap
plying insecticides on anthurium, because phytotoxic
ity varies among cultivars and is more likely to occur
under hot, dry growing conditions. When thrips injury
is sustained during the bud stage, injured anthurium flow
ers will be harvested for at least a month following ap
plication of an effective insecticide.
In banana, spraying the immature bunches and the
surrounding soil can significantly reduce thrips damage
to the fruit; when bagging bunches, spray just before
bagging. A contact, granular insecticide applied in a 30
inch radius around each banana plant is effective against
the prepupal and pupal stages of banana rust thrips that
inhabit the soil. No granular insecticide is currently reg
istered for use on anthurium.
References
Caldwell, N.E.H. 1938. The control of banana rust thrips.
Bulletin 16, Department of Agriculture and Stock, Di
vision of Plant Industry (Research), Queensland,
Australia.
Denmark, H.A., and L.S. Osborne. 1985. Chaetanapho
thrips signipennis (Bagnall) in Florida (Thysan
optera: Thripidae). Ento. Circular no. 274, Sept. 1985,
Florida Department of Agriculture and Consumer
Service, Division of Plant Industry.
Hara, A.H., C. Jacobsen, and R. Niino-DuPonte. 2002.
Anthurium thrips damage to ornamentals in Hawaii.
4
CTAHR — June 2002
University of Hawaii at Manoa, College of Tropical
Agriculture and Human Resources, publication IP-9.
4 pp.
Jacot-Guillarmod, C.F. 1974. Catalogue of the Thysan
optera of the world (Part 3). Annals of the Cape Pro
vincial Museums—Natural History 7(3):517–976.
Lewis, T. (ed.) 1997. Thrips as crop pests. Institute of
Arable Crop Research, Rothamsted, Harpenden,
Hertfordshire, CABI Publishing, UK.
Pinese, B. 1987. Soil and bunch applications of insecti
cides for control of the banana rust thrips. Queensland
Journal of Agricultural and Animal Sciences 44(2):
107–111.
Pinese, B., and R. Piper. 1994. Bananas; insect and mite
management. Queensland Department of Primary
Industries, Australia. 67 pp.
Pinese, B., and R. Elder. 2000. DPI Notes; pest of plants;
bananas; banana rust thrips in bananas. Department
of Primary Industries, Queensland Horticulture In
stitute, Australia. 5 pp. <http://www.dpi.qld.gov.au/
horticulture/5528.html>.
Sakimura, K. 1975. Danothrips trifasciatus, new spe
cies, and collection notes on the Hawaiian species
Danothrips (Thysanoptera: Thripidae). Proc. Hawai
ian Entomol. Soc. 22:125–132.
Schulz, D. ca. 1950. Department of Entomology, Uni
versity of Illinois at Urbana-Champaign. <http://
www.life.uiuc.edu/Entomology/insectgifs/>.
Stover, R.H., and N.W. Simmons. 1987. Bananas (3rd
edition). Longman Scientific and Technical, Harlow,
UK. 468 pp.
Photo credits: Figure 3 by C. O’Donnell, University of California–
Davis; all others by A. Hara.
Insect Pests
June 2002
IP-11
Blossom Midge in Hawaii—
a Pest on Ornamentals and Vegetables
Arnold H. Hara and Ruth Y. Niino-DuPonte
B
lossom midge, Contarinia maculipennis Felt (Dip
tera: Cecidomyiidae), has been present in Hawaii
since the early 1900s and is thought to have originated
in Asia (the “West Indies”). Currently, the blossom midge
can be found on all of the major Hawaiian islands. Jensen
(1946) presented compelling evidence that C. macu
lipennis had been misidentified in earlier reports as C.
solani (Rübsaamen) or C. lycopersici Felt due to its di
verse range of hosts. Elsewhere in the USA, the blos
som midge was reported on dendrobium orchids in
Florida in 1992.
Hosts
The blossom midge has a wide host range spanning at
least six plant families, including the flower buds of or
chids, plumeria, hibiscus, pikake (jasmine), white mus
tard cabbage or pak choi, tomato, eggplant, pepper, po
tato, bittermelon, and other vegetables and ornamentals.
buds. They are white to cream colored, invisible to the
naked eye, and hatch within 24 hours into maggots that
move into the bud and feed on fluids from the damaged
plant tissue.
The maggots are white when newly hatched, becom
ing yellow with a pink tinge as they age (Fig. 2). As
they mature in 5–7 days, growing to 1⁄12 inch long (about
the thickness of a nickel), the maggots are capable of
flipping themselves several inches into the air to exit
the buds and burrow into the soil to pupate, like other
ground-pupating fly larvae such as the melon fly and
oriental fruit fly.
Pupation is most successful in soil that is moist but
not wet. The late-stage pupa turns from yellowish-white
to brown (Fig. 3) and burrows back up to the soil sur
face in preparation for emergence as an adult 14–21 days
after entering the soil. The pupa works itself partially
free of the soil, and the adult emerges, leaving the pupal
skin protruding from the soil.
The adult blossom midge is tiny, about the thickness
of a nickel in length; males are slightly smaller than fe
males. The adult is somewhat mosquito-like, with typical
fly features, and survives for only 4 days. It has relatively
large, multifaceted eyes and a single pair of spotted wings
about one to two times as long as its body (Fig. 4).
Biology
The blossom midge reproduces year-round in Hawaii.
The duration of its life cycle from egg to adult is ap
proximately 21–28 days. The eggs are deposited in
masses by the adult female into the open tips of flower
Behavior
Except for the adult, all stages of the blossom midge are
secluded within the bud (as maggots) or in the soil (as
pupae). Adult emergence from pupae in the soil usually
occurs in the early evening.
Damage
Blossom midge maggots feed inside unopened flower
buds, causing deformed, discolored buds and blossoms
and, in severe infestations, premature bud or blossom
drop (Fig. 1). As many as 30 maggots may be found
infesting a single dendrobium bud.
IP- 11
Blossom Midge in Hawaii—a Pest on Ornamentals and Vegetables
CTAHR — June 2002
Figure 1. Feeding damage to flower buds by blossom midge: left, plumeria buds; center, dendrobium buds; right,
dendrobium bud drop. (Photos: A. Hara, R. Mau)
When laying eggs, the adult female blossom midge
is unable to penetrate plant tissues but rather inserts its
ovipositor into the open end of a bud. To ensure an opti
mal food source and moist environment, the adult midge
avoids late-stage buds and prefers to lay eggs in young
buds whose growth to maturity will approximately par
allel that of the maggot.
If growing conditions become unsuitable for larval
development (for example, if the flower or bud on which
maggots are feeding begins to dry), immature maggots
may leave the flowers or buds to pupate in the soil; how
ever, their pupation may take a few weeks longer, and
the emerging adult midges are invariably smaller than
adults from fully mature maggots.
In Florida, blossom midge populations maintained
in greenhouses were observed to decrease rapidly dur
ing the winter, even though the temperature was main
tained at 65°F and the plants had sufficient numbers of
buds.
Cultural control
Sanitation is the most important management practice
for the blossom midge. Remove and destroy all dropped
buds and infested buds still on the plant. Place infested
flower buds in a plastic bag or a sealed container to pre
2
vent escape of maggots.
Due to the blossom midge’s wide range of hosts,
avoid planting possible alternate hosts around the crop
area.
A certain variety of tomato was observed to be more
susceptible to blossom midge infestation due to its flower
structure, which facilitates ovipositing. Host plant vari
eties in which petals remain tightly fitted until the bud
is almost ready to open may reduce susceptibility.
Biological control
To date, no parasites have been isolated or specifically
introduced to Hawaii to control the blossom midge. The
adults are vulnerable to general predators, such as web
spinning spiders and ants. Ants may also prey on pupae
in the soil.
Chemical control
Only the adult stage of the blossom midge is vulnerable
to contact insecticides, because the maggots are protected
within the bud and the pupae are burrowed in the soil.
Some insecticides can be applied as a foliar spray
against larvae as well as a soil treatment to target the
pupal stage. Translaminar insecticides (those that move
from the sprayed leaf surface to the lower surface) may
IP- 11
Blossom Midge in Hawaii—a Pest on Ornamentals and Vegetables
Figure 2. Blossom midge larvae in a dendrobium bud.
CTAHR — June 2002
Figure 3. Blossom midge pupae from hibiscus.
Photos in Figures 2 and 3 by Walter Nagamine, Hawaii Dept. of Agriculture;
Figure 4 photo by S. Chun.
The actual size of the larvae and pupae is 1–2 mm; the adult is about 2 mm long.
1 mm is just over 1⁄32 inch; the following lines are 1 and 2 mm long, respectively:
be capable of penetrating the bud to affect the maggots.
Trials of systemic insecticides (those that are spread from
the site of application throughout the rest of the plant)
on dendrobium have been disappointing, possibly be
cause the chemicals are not able to reach the flower buds
to affect the maggots.
Consult the Hawaii Department of Agriculture or
the CTAHR Cooperative Extension Service for regis
tered chemicals that are known to be effective against
the blossom midge.
References
Felt, E.P. 1933. A hibiscus bud midge new to Hawaii.
Proceedings, Hawaiian Entomological Society 8(2):
247–248.
Gagné, Raymond J. 1995. Contarinia maculipennis
(Diptera: Cecidomyiidae), a polyphagous pest newly
reported for North America. Bulletin of Entomologi
cal Research 85:209–214.
Jensen, D.D. 1946. The identity and host plants of blos
som midge in Hawaii (Diptera: Cecidomyiidae: Con
tarinia). Proceedings, Hawaiian Entomological So
ciety 12(3):525–534.
Jensen, D.D. 1950. Notes on the life history and ecol
ogy of blossom midge Contarinia lycopersici Felt
(Diptera: Cecidomyiidae). Proceedings, Hawaiian En
tomological Society 14(1):91–100.
Figure 4. Adult blossom midge.
Osborne, L.S., T.J. Weissling, J.E. Pena, and D.W.
Armstrong. 2001. A serious pest is causing signifi
cant problems for dendrobiums and hibiscus grow
ers. In: Felter, L., T. Higgins, and N. Rechcigl (eds.),
Proceedings, 17th Conference on Insect and Disease
Management on Ornamentals. February 25–27, 2001,
Orlando, FL. Society of American Florists, Alexan
dria, VA. p. 21.
3
Insect and Mite Pests of Macadamia Nuts
in Hawai‘i—A Quick Reference Guide
College of Tropical Agriculture and Human Resources • University of Hawai‘i at Manoa
This poster provides a quick reference guide
to CTAHR’s book Macadamia Integrated Pest
Management by Vincent P. Jones, 2002.
The most important arthropod pests of
macadamia are illustrated here, with brief
comments on their biology and the damage
caused. A page reference to the Jones book
is provided for each.
Compiled by Mark G. Wright, Department of Plant and Environmental
Protection Sciences. CTAHR publication IP-12, June 2003.
Photos from V. Jones, Macadamia Integrated Pest Management, 2002.
Tropical nut borer (TNB)
Where do they occur?—TNB are found in sticktight nuts, in nuts on the
orchard floor, and in alternative hosts, e.g., carob, asoka fruit, and castor bean.
Egg, larva, and pupa
Adult TNB
Damage to a kernel
by TNB
(Jones, p. 24)
What kills them?—The beetles shown here eat
the eggs and larvae of TNB.
Chemical control can be
achieved with endosulfan.
Predatory beetle larva
Southern green stinkbug (SGS)
Where do they occur?—SGS attacks macadamia nuts and various weed species. They
attack nuts both on the tree and the ground.
SGS causes pitting on
kernels, resulting in rejection
of nuts by processors.
Beetle adults
(Jones, p. 35)
What kills them?—SGS eggs are parasitized by a wasp; the adults are parasitized
by a fly. The flies lay eggs on the adult
SGS (arrow), and their larvae
burrow into and kill the bug.
Fly
(3⁄8″ long)
Eggs and different stages of SGS development
SGS damage to kernel
Wasp (1⁄16″ long)
Koa seedworm and litchi fruit moth
(Jones, p. 42)
What kills them?—No parasites
of their eggs are found here. Some
parasitic wasps attack and kill the
larvae. Chemical control of koa
seedworm is not recommended.
Where do they occur?—Koa seedworm moths lay their eggs on the husks of macadamia
nuts. The larvae then bore into the husk or kernel, if the shell has not yet hardened.
Adult koa seedworm moth
Eggs (3⁄100″ diameter)
Larvae may bore into
the kernel
Koa seedworm damage to
macadamia kernels
Wasps (four species) (bodies ~1⁄16″ long)
Some minor pests
Broad mites feed on macadamia flowers, leaves, and
fruit; damage to flowers may
be significant (Jones, p. 52).
Broad mite damage on husks
Flat mites rarely
cause economic
damage (Jones, p. 54).
Flat mite damage on husks
Redbanded thrips feed on husks and leaves. They may cause
malformation of leaves (Jones, p. 56).
Thrips damage to
leaves
Redbanded thrips
damage to husks
Redbanded thrips
juveniles have red
bands; adults are black
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in cooperation with the U.S. Department of Agriculture. Andrew G. Hashimoto, Director/Dean,
Cooperative Extension Service/CTAHR, University of Hawaii at Manoa, Honolulu, Hawaii 96822. An Equal Opportunity / Affirmative Action Institution providing programs and services to the people of Hawaii without regard to race, sex, age, religion, color, national origin, ancestry,
disability, marital status, arrest and court record, sexual orientation, or veteran status. CTAHR publications can be found on the Web site <http://www.ctahr.hawaii.edu> or ordered by calling 808-956-7046 or sending e-mail to [email protected].
Insect Pests
July 2003
IP-13
Integrated Pest Management for Home Gardens:
Insect Identification and Control
Richard Ebesu
I
ntensive, high-production agricultural systems have
traditionally used synthetic pesticides as the primary
tool to eliminate pests and sustain the least amount of
economic damage to the crop. Dependence on these pes
ticides has led to development of pest resistance to pes
ticides and increased risk to humans, other living or
ganisms, and the environment.
Integrated pest management (IPM) is a sustainable
approach to managing pests that combines biological,
cultural, physical, and chemical tools in a way that mini
mizes economic, health, and environmental risks.
The objective of IPM is to eliminate or reduce po
tentially harmful pesticide use by using a combination
of control methods that will reduce the pest to an ac
ceptable level. The control methods should be socially
acceptable, environmentally safe, and economically
practical. Many commercial agricultural systems use
IPM methods to manage pest problems, and home gar
deners can use similar methods to control pest problems
in their gardens.
The first key to IPM is to identify the pest. This
publication describes the major pests of home garden
crops in Hawaii and gives their identifying characteris
tics. The second key to IPM is to know which stages of
the pest cause damage and which are most susceptible
to management with the various possible control meth
ods. With an understanding of the pest life cycle and its
relationship to the susceptible host plant, and with knowl
edge of the types of control methods available, garden
ers can better utilize IPM to manage common insect pest
problems. The elimination or reduction in pesticide use
that can be achieved through thoughtful application of
IPM strategies will prevent misuse of pesticides and help
keep the environment healthy.
IPM components and practices
Integrated pest management strategies consist of site
preparation, monitoring the crop and pest population,
problem analysis, and selection of appropriate control
methods. Home gardeners can themselves participate in
IPM strategies and insect control methods with a little
knowledge and practice.
Preparation
What control strategies can you use before you plant?
You need to be aware of potential problems and give
your plants the best chance to grow in a healthy envi
ronment.
Soil preparation
Improve the physical properties of the soil including
texture and drainage to reduce waterlogging. Improve
soil fertility and soil organic matter by working well
rotted compost into the soil.
Prevent pest build-up with crop rotation, fallowing,
and using resistant crop varieties or crops less suscep
tible to pests.
Monitoring (scouting) for pests
Observe your garden and learn to identify the pest prob
lems, as well as beneficial organisms.
Problem analysis
Do you have a pest problem? Is it a pest such as an in
sect or plant disease? Is it a nutrient deficiency or a prob
lem with soil drainage? Is the pest problem major and
needs control or minor and can be tolerated?
IP- 13
IPM for Home Gardens—Insect ID and Control
Insect identification
If you have an insect problem, you need to know what
insect pest you are dealing with and what stage of the
insect’s life cycle is the most likely to cause damage, as
well as the stage most susceptible to control measures.
General insect information
Insects have lived on the planet Earth for about 350
million years. Insects have adapted to just about every
type of habitat, including plants, animals, soil, water,
snow, deserts, buildings, stored products, and people.
Most insects are not pests, and it is impractical to at
tempt to eliminate all the insects from our environment,
so insect pest management strategies should include a
variety of techniques. Integrated pest management (IPM)
of insects is designed to use these techniques to reduce
pesticide use, use less toxic pesticides, and use environ
mentally safe pesticides to keep insect populations be
low economically damaging levels.
Characteristics of insects
Insects are invertebrates (no backbone) with an exosk
eleton (outer skeleton). Their bodies are segmented with
three major body regions: the head, thorax, and abdo
men. Adults have a pair of antennae, a pair of compound
eyes, three pairs of legs, and zero, one, or two pairs of
wings. Their appendages and mouthparts come in a va
riety of shapes, sizes, and functions. They respire mostly
through holes in their body called spiracles (for terres
trial insects) and by diffusion through the body wall (in
aquatic insects). Insects are cold-blooded; their body
temperature closely follows the temperature of their sur
roundings. Insects differ from mites, ticks, and spiders,
which have two major body sections, four pairs of legs,
and lack antennae and compound eyes. Centipedes are
arthropods with one pair of legs on each body segment,
and millipedes have two pairs of legs on a body seg
ment. Sowbugs are crustaceans, usually with seven pairs
of legs.
Insect development
All insects develop from eggs. Most hatch after the egg
is laid, but some, like the aphids, hatch within the fe
male, and live young are produced. Metamorphosis is
the change in form from the egg to adult stage.
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CTAHR — July 2003
Simple or gradual metamorphosis
Eggs hatch and there is a gradual change as the imma
ture forms, called nymphs, mature to the adult stage.
Nymphs have compound eyes and antennae and re
semble the adults but are smaller, without fully devel
oped wings, and cannot reproduce. Wings of the adult
develop externally, and there is no resting stage, like a
pupa. Nymphs usually live in the same habitat as the
adults. Development is sometimes called ametabolous
in forms without wings, such as collembola and silver
fish. Insects with gradual metamorphosis include grass
hoppers, cockroaches, and aphids.
Some insects, such as dragonflies, have an incom
plete metamorphosis. Their nymphs live in water, have
gills, and differ in appearance from the adults; they
emerge from the water and molt into the adult form with
wings, without a resting stage.
Complete metamorphosis
Immature stages are normally worm-like and are called
larvae. Larvae do not have compound eyes, some may
have thoracic legs, and some have leg-like appendages
on the abdomen. The last larval stage is a resting stage
called the pupa. The pupa does not feed, usually is not
active, and often is covered by a silken cocoon. Wings
are developed internally, and upon emergence the adult
expands the wings. Immature and adult stages are usu
ally different in form and often live in different habitats.
Insects with complete metamorphosis include butterflies,
flies, wasps, and beetles.
Insects and their importance to people
Injury to plants
Many insects are agricultural pests; they
• chew leaves, stems, bark, or fruits of plants
• suck sap from leaves, buds, stem, or fruits
• bore and tunnel into bark, stems, twigs, wood, fruits,
nuts, and seeds
• cause galls and abnormal growth on plants
• attack the roots of plants in any of the above ways
• lay eggs in plant tissue
• take plant parts for nest or shelters
• carry other harmful insects to plants
• vector (transmit) plant diseases.
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CTAHR — July 2003
Types of pest activity and examples of organisms.
Activity in relation to plants
Examples of organisms
Chewing leaves, stem, fruit
Sucking plant sap
Boring, tunnels
Galls on plants
Egg-laying
Waste product contamination
Remove parts for nests or shelter
Carry or protect pests
Transmit plant disease
Grasshoppers, beetles, caterpillars, slugs
Aphids, leafhoppers, whiteflies, scales, thrips, mites
Leafminer, weevils, twig borers, root borers, caterpillars
Gall wasp, erinose mites
Katydids, fruit flies
Cockroaches, caterpillars, ants, aphids, whiteflies
Leaf-cutting bees, some ants, bagworms
Ants
Aphids, leafhoppers, thrips
Injury to animals or people
Annoyance and buzzing
Biting, stinging
Transmit disease
Infesting animals, people
Contamination
Flies, mosquitoes
Mosquitoes, fleas, wasps, bees, bed bugs
Mosquitoes, fleas, ticks
Bot fly, ticks, lice
Cockroaches, flies
Damage to products, structures
Wood structures
Stored products, food
Clothing, fiber
Termites, powderpost beetles
Flour beetle, meal moth, rice weevil, cigarette beetle
Clothes moth, carpet beetle
Beneficial qualities
Pollinate flowers
Products, honey, wax, silk, dye
Biological control
Food source (people, animals)
Decompose carcasses, dung
Soil improvement, excavation
Scientific research, medicine
Aesthetic value
Injury or annoyance to people and animals
Some insects are general annoyances; they
• cause annoyance by their presence, buzzing, foul
odors, and excretions on foods
• infest fruits
• bite
• enter the eyes, ears, nose
• lay eggs on skin, hair, feathers
Bees, flower flies
Honey bee, silkworm, mealybug
Lady beetle, praying mantis, wasps, flies
Beetles, flies, grubs
Maggots, beetles
Beetles, springtails
Vinegar fly, bees (stings)
Butterflies, beetles
•
•
•
•
•
•
apply venom by biting, stinging, or hairs
leave caustic body fluids or irritants when crushed
cause allergies
can be poisonous if swallowed
make their homes on or in the body as parasites, in
juring the host
transmit disease organisms or create unsanitary con
ditions.
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IP- 13
Damage to stored products, possessions,
buildings, and utilities
Insects are serious pests when they
• stored food, clothing, fiber, and paper may be eaten
or contaminated by excretions
• termites and wood-boring insects damage structures
and furniture
• termites may feed on wire insulation and cause elec
trical fires and damage gaskets and seals leading to
water loss.
Insects can be beneficial
Not all insects are pests; they
• pollinate flowers producing fruits, seeds, vegetables,
and flowers
• produce silk, beeswax, shellac, honey, and dyes
• are used in biological control as predators and para
sites to destroy pest insects and weeds
• are food sources for some people, fish, birds, and ani
mals
• scavenge to remove carcasses, dead plant material,
and dung
• help to improve the soil by burrowing and providing
organic matter
• are important in scientific research and genetics
• can be pleasing and entertaining—some butterflies
and beetles are colorful and are collected as a hobby
• have had some value in medicine (such as maggots
cleaned out wounds, honeybee stings for arthritis).
Insect orders important in gardens and homes
CTAHR — July 2003
order. The Pacific beetle cockroach is often a pest on
cypress and juniper trees; it girdles the twigs and limbs,
often killing the branches. Household pests include the
American cockroach, German cockroach, and brown
banded cockroach.
Praying mantises are general predators and feed on
other insects.
Thysanoptera
Thrips are small, slender insects with mouthparts modi
fied into a short beak used to suck the plant sap. Their
wings are slender, with fringed margins. Thrips are im
portant plant pests. Their feeding often causes a stipling
of leaf tissue accompanied by scarring, bronzing, or sil
vering. Some are major vectors of plant viruses.
Melon thrips are pale yellow, tend to be found on
flowers and young foliage. Damaging on a range of
plants including cucumber, watermelon, tomato, egg
plant and beans.
Western flower thrips are important vector of to
mato spotted wilt virus affecting a number of plants in
cluding tomato, pepper, lettuce and flowering plants.
Red-banded thrips adults are black, while the lar
vae are yellow with a red band on the abdomen; their
feeding damage often scars fruits.
Hemiptera or Heteroptera
In these “true bugs,” the basal portion of the front wings
are somewhat thickened and leathery; the tip portion is
membranous. The hind wings are membranous, and the
wings are held flat over the abdomen with the tips of the
front wing overlapping. They have piercing-sucking
Orthoptera
In grasshoppers, crickets, praying mantises, and cock
roaches, the forewings of the adults are usually long and
narrow and somewhat thickened. The hind wings are
membranous, broad, and folded beneath the forewings
at rest. Mouthparts are the chewing type; the antennae
are often long and slender.
Among the grasshoppers, the pink-winged grasshop
per is common. Its head is pointed, the antennae fairly
short, the body color is light green to brown. Others in
clude the longhorned grasshopper and occasionally the
aggravating grasshopper.
The mole cricket and the twospotted cricket feed on
the roots of plants and may be a problem in some cases.
Cockroaches can be classified in their own separate
4
Figure 1. Thrips feeding may cause silvering damage.
IP- 13
mouthparts formed into a slender beak. Some are plant
feeding, while others are predatory.
Southern green stinkbugs are pests on beans, tomato,
cabbage, and macadamia nut. Nymphal stages are dark
colored with whitish markings; adults are mostly light
green and shield-shaped.
Black stinkbugs are small, rounded, and shiny black
with pale stripes; they are an occasional pest on beans
and some other legumes.
Lace bugs cause stipling of leaves similar to other
sucking insects; they commonly infest azaleas and
rhododendron in Hawaii.
Seed bugs include the southern chinch bug, a pest
on St. Augustine grass lawns; others bore into seeds.
Assassin bugs are important predators of other in
sects.
CTAHR — July 2003
These include aphids, whitefly, scales, leafhoppers, and
mealybugs. They are plant-sucking, and many excrete
honeydew, a liquid high in sugar, which attracts ants
and is used as a substrate for sooty mold fungus, which
interferes with plant photosynthesis. Some are soft bod
ied, slow moving, or sedentary, forming colonies with
wingless forms. Others are active. Adults have wings
held roof-like over the body; the antennae are often short
and bristle-like (as with leafhoppers). With sucking
piercing mouthparts, many are vectors of plant viruses.
Some secrete molted skins or a waxy, powdery substance
that covers the body. Many are spread by the wind or
carried by ants that feed on the honeydew and protect
the insects from natural enemies.
Aphids are small, rounded or pear-shaped, soft bod
ied, most with a pair of tube-like cornicles on the poste
rior of the abdomen. Some are covered with a white pow
der. Aphids suck the plant sap from leaves, stems, and
roots, often causing stunting, wilting, and deformed
leaves. The group is very important as vectors transmit
ting plant viruses. Females are able to reproduce with
out mating, giving birth to live offspring. Most are wing
less but produce winged forms in crowded or poor con
ditions and are easily blown by the wind to other plants.
Their color ranges from bright yellow to red, green,
brown, and black. Important aphids include green peach
aphid, melon aphid, cabbage aphid, banana aphid, yel
low sugarcane aphid, black citrus aphid, and potato aphid.
Whiteflies are tiny; the adults resemble white moths;
the immature stages look like scale insects. Adults’ wings
are covered with a white, waxy powder, making them
difficult to wet. Some are vectors of plant viruses; others
cause various plant disorders such as silver-leaf. Impor
tant whiteflies include silverleaf whitefly, greenhouse
whitefly, spiraling whitefly, and anthurium whitefly.
Scales have adult females that are wingless, often
legless, and sedentary. Two groups are the soft scales
and the armored scales. Soft scales tend to be flattened,
oval, elongated, and covered with a waxy substance or
a smooth, hard outer covering. Armored scales are very
small, soft bodied, and concealed under a scaly cover
ing that is free from the body, formed by waxy secre
tions and the shed skins of its immature stages. Impor
tant soft scales include green scale and hemispherical
scale. Armored scales include oleander scale, magnolia
white scale, and Boisduval scale.
Figure 2. Aphids suck plant sap and spread plant diseases.
Figure 3. Whiteflies are covered with a waxy coating.
Homoptera
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CTAHR — July 2003
Mealybug females are oval and segmented with well
developed legs. The body is covered with a mealy or
waxy substance. Mealybugs can be found on almost any
part of the host plant including leaves, stems, roots, and
fruits. Important mealybugs include pineapple mealy
bug, gray pineapple mealybug, and citrus mealybug.
Leafhoppers are elongated, slender insects with
bristle-like antennae; the wings of adults are held roof
like over the body, and they often hop when disturbed.
They have one or two rows of spines on the hind legs.
Some are vectors of plant viruses; others cause a phyto
toxic reaction due to feeding called hopperburn. Impor
tant leafhoppers include twospotted leafhopper, Steven’s
leafhopper, and Southern garden leafhopper.
Planthoppers are similar to leafhoppers but have a
flattened spur on the hind tibia and lack the rows of spines
on hind legs. Many have reduced or shortened wings.
Important planthoppers include corn delphacid, taro
delphacid, and sugarcane delphacid.
Treehopper adults have a humpback appearance.
Solanceous treehopper nymphs are orange with black
spiny projections and can be found on tomato, eggplant,
and peppers.
Spittlebug nymphs produce white spittle, a froth
like covering, to conceal themselves. They are found on
rosemary, basil, mint, hibiscus and other plants.
Psyllids are small, jumping insects resembling
aphids. They are a nuisance pest on monkeypod and koa
haole. Native psyllids on ohia plants cause leafgalls.
Isoptera
Figure 4. Leafhopper feeding is often toxic to plants.
Figure 5. The Chinese rose beetle feeds at night.
6
The Formosan subterranean termite feeds on cellulose,
which is found in plant material. Although normally
found in wood, the termites can feed on live plant tissue
including roots and fruits.
Insects with complete metamorphosis
Coleoptera
The coleoptera (beetles and weevils) are the largest in
sect order, including pests and beneficial insects. The
adults have a hardened, sometimes horny outer skeleton,
usually with two pairs of wings, the outer pair thick
ened, leathery, or hard and brittle, usually meeting in a
straight line down the middle, and the inner pair mem
branous (mostly). Adults usually have a noticeable pair
of antennae, variously shaped. Both adults and larvae
have chewing mouthparts. Beetle larvae, also known as
grubs, have a head capsule, three pairs of legs on the
thorax, and no legs on the abdomen. Weevil larvae lack
legs on the thorax.
Foliage feeders, including Chinese rose beetles, feed
at night, and heavy infestations cause lace-like appear
ance of leaves. Rose beetles are common and damage
many different plants including rose, grapes, beans, egg
plant, corn, cucumber, ginger, and ornamentals.
Tobacco flea beetles are tiny brown beetles whose
feeding damage causes shot-hole appearance of leaves.
They are found on eggplant and tobacco.
Stem borers include long-horned beetles, whose
adults have long antennae and larvae bore into stems,
and wood; pinhole borers that leave pin-holes in
branches, and wood; orchid weevils, whose larvae bore
into orchid stems and tissue; black twig borers, whose
IP- 13
CTAHR — July 2003
adults bore through stems of coffee and other economi
cal and ornamental plants and whose larvae feed on fun
gus cultured by the adult female.
Root borers include banana root borer, whose grubs
bore into the banana corm causing damage and poor
growth, and sweetpotato weevil, whose grubs feed in
side the stems and tubers, often followed by decay or
ganisms.
Fruit weevils include pepper weevils, the adults and
grubs of which infest peppers and cause internal dam
age and premature drop, and mango seed weevil, whose
grubs bore into the seed, preventing fresh fruits to be
exportable.
Household pests include confused flour beetle, rice
weevil, cigarette beetle, and carpet beetle; they may in
fest stored grain products and other household belong
ings.
Beneficial beetles include ladybird beetles, also
called ladybugs, which feed on homopteran insects such
as aphids, scales, mealy bugs, whiteflies, and psyllids,
and scavenger beetles, which help to remove carcasses
from the environment.
leaves by leafmining or bore into stems and fruits. Some
lepidoptera have been successfully used to control
weeds, such as some cactus species. Some pupae forms
are distinctive of the species or family.
Noctuid moths include common pests such as lawn
armyworm, beet armyworm, corn earworm, cabbage
looper, black cutworm, and monkeypod-kiawe caterpil
lar. The adults are active at night and often are attracted
to lights.
Diamondback moth adult males have a diamond
pattern on the wings when folded over the back. Dia
mondback moth is a pest of cabbages, and the leek moth
attacks onions.
Hawk moth caterpillars are called hornworms for
the distinctive, hornlike protrusion at the rear of the ab
domen. They include sweetpotato hornworm and ole
ander hawk moth.
Other pests include citrus swallowtail, imported
cabbage worm, cabbage webworm, banana skipper, to
mato pinworm, and various leafrollers.
Household pests include Indian meal moth and
casemaking clothes moth.
Lepidoptera
Diptera
Lepidoptera (butterflies and moths) have a caterpillar
(larval) stage that causes the most damage by chewing
and boring, while the adult, fruit piercing moth may be
a pest on some ripe fruits. Most adult lepidoptera have
long, siphoning, tube-like mouthparts to feed on plant
nectar. Larval (caterpillar) stages have chewing mouth
parts; most have three pairs of thoracic legs and five or
less pairs of abdominal prolegs. Most larvae feed on
The diptera (flies, fruit flies, leafminers, and midges)
adults have only one pair of wings and have sucking
mouthparts that may be modified. Their larvae are called
maggots, are legless, and many lack a well defined head
capsule, with only hook-like mouthparts. The order is
important in medical and veterinary entomology and
includes fruit flies, mosquitoes, house flies, horse flies,
and blow flies.
Figure 6. Grubs are immature beetles or weevils.
Figure 7. Sweetpotato hornworm.
7
IP- 13
Tephritid fruit flies at present include four economi
cally important species in Hawaii: Mediterranean fruit
fly, Oriental fruit fly, melon fly and solanaceous fruit
fly. The maggots infest fruits and fruiting vegetables and
thus prevent many fruits and vegetables from being ex
portable without disinfestation treatment.
Leafminers are important agricultural pest. The
small adults lay eggs on plant tissues and the larvae bore
into the tissues and create tunnels or mines. Heavy in
festations can cause reduced photosynthesis and leaf
drop, interrupt the uptake of water and nutrients, and
cause wilting. The group includes bean fly, serpentine
leafminer, and vegetable leafminer.
Midge adults are small, delicate, gnat-like flies.
Midge pests include mango blossom midge, chrysan
themum gall midge, and a blossom midge on pikake,
plumeria, and orchids.
Beneficial flies includes parasitic flies like the ta
chinid flies and predators like the syrphid fly larvae and
aphid flies; others are important as scavengers.
CTAHR — July 2003
the abdomen is fused with the thorax and constricted to
form a narrow, waist-like connection. The Apocrita lar
vae are grub-like or maggot-like, legless, and often lack
well developed head capsules.
Plant pests include seed wasps, gall wasps, orchidfly,
leafcutting bees, and some ants. Ants usually do not feed
directly on plants, but their presence may be a nuisance.
In addition, ants that feed on honeydew excreted by
aphids and scale insects in turn protect those insects from
predators.
Household pests include ants, some wasps, carpen
ter bees and occasionally honeybees.
The most significant contribution is the parasitic and
predatory nature of the many wasps and the pollinating
of important fruit crops by bees.
Among the ants, bees, wasps, the suborder Symphyta is
an important group of plant feeders, but it is not com
mon in Hawaii. Here the suborder Apocrita is of rela
tively minor concern as plant pests but is an important
group that includes beneficial pollinators, parasitoids,
and predators used in biological control of insect pests.
The adults have membranous wings, the forewings be
ing larger than the hind wings, and many have a well
developed ovipositor modified into a sting. The base of
Mites
Mites are more closely related to spiders than insects,
but some are important plant pests. Like the spiders,
mites have two major body parts, four pairs of legs, and
the plant-feeding mites often have rasping mouthparts.
In addition, many are predators and help to control other
plant-feeding mites and some insects. Most mites are
very small and difficult to see without magnification.
Spider mites include carmine spider mite and
twospotted spider mite; their feeding damage includes
stippling of the leaves.
Broad mites are found on many plants including
papaya and pepper, where they feed on the young, grow
ing leaves, causing distortion and bronzing.
Erinose mites include tomato russet mite, hibiscus
Figure 8. Fruit fly maggot and pupae.
Figure 9 Leafminer maggots form tunnels on leaves.
Hymenoptera
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IP- 13
erineum mite, lychee erinose mite, and the papaya leaf
edgeroller mite.
Medically important mites and tick pests include the
house dust mite, itch mite, brown dog tick, Rocky Moun
tain tick, and chiggers.
Other pests
Slugs and snails feed mostly at night; they can feed on
bark and girdle stems, and chew leaves and fruits. Slugs
hide during the day under boards, rocks, potted plants,
and in the soil.
Birds tend to feed on fruits and young tissues like
the cotyledons of emerging seedlings and flower buds.
Rodents feed on fruits and may chew on the bark
and stems of some plants. Mice have been known to
spread plant diseases in nurseries by carrying the patho
gen on their feet from one plant to another. Rodents may
enter homes and other buildings and feed on stored prod
ucts.
IPM insect control methods
CTAHR — July 2003
tors such as birds. Deep plowing may bury some insects
so they cannot emerge on the surface.
Crop rotation and fallow eliminate the insect host
plant to disrupt the life cycle.
Sanitation removes crop residues and infested plants
to eliminate sources of insects.
Crop timing manipulation includes planting early
maturing varieties before the pest insect population
builds up.
Mixed cropping involves planting several species
of crops including cover crops in the same area to create
diversity, thereby eliminating a monoculture system.
Insects need to search for the host plant, while other
plants provide a habitat or food for beneficial insects.
Trap crops are crops planted for the pests so they
leave the desired crop alone. Pesticides can often be used
on the trap crop that cannot be applied on the desired
crop.
Proper use of fertilizer and water result in healthy
plants that normally are more tolerant of insects and dis
ease. Overhead watering may also disrupt diamond back
moth mating and egg laying in watercress fields.
Cultural controls
These methods are used in the process of cultivating the
crop. The techniques are used to disrupt the normal life
cycle of the pest. IPM strategies include changing the
environment by eliminating the host plant, attracting the
pest away from the host plant, and using mechanical
means to trap insect pests.
Tilling and plowing physically destroy soil insects
or expose them to adverse weather, temperature or preda-
Figure 10. Slugs feed on plants at night.
Mechanical and physical controls
These methods utilize machinery, manual operations, or
the physical environment in cultivation practices and
may be more practical for small gardens. For example,
remove insects, their eggs, and infested plant parts by
hand-picking, or hose off pests like aphids. Vacuums
also can remove some pests from plants.
Mechanical exclusion uses barriers such as screens,
netting, and row covers to keep pests off the plants.
Collars around seedlings prevent cutworms, sticky
coated tree trunks prevent access by crawling insects,
and copper barriers repel slugs.
Mechanical traps such as colored sticky traps can
be used to control or monitor insects. Many insects are
attracted to yellow, while other colors used include blue,
red, and white. Pheromone-baited traps can attract a
certain sex, usually males, of an insect species and can
help reduce the mating population in the area. Food baits
are also used in traps and usually attract both sexes.
Physical manipulation examples include tempera
ture extremes such as heat or cold to control pests. Solar
radiation helps to control soil insects and nematodes.
Water can be used to forcefully wash insects off plants
and also to disrupt their mating. Flood conditions force
9
IP- 13
soil insects to the surface where predators can feed on
them. Light can attract insects or confuse nocturnal in
sects such as the Chinese rose beetle. Aluminum mulches
reflect light to repel some aphids, whiteflies, and thrips.
Irradiation, heat, and cold temperatures are used in
postharvest treatments. Electricity is used in drywood
termite control.
Biological controls
Living organisms naturally compete for food and living
space. Biological control is the manipulation of one liv
ing organism to control another living organism. In Ha
waii, introductions of biological control agents are done
by government agencies; however, home gardeners can
help themselves by providing a favorable environment
for predators and parasites as well as using less harmful
pesticides and thus avoid killing beneficial insects.
Predators eat insect pests. Examples include lady
bugs, praying mantis, assassin bugs, lace wings, preda
tor mites, spiders, lizards, frogs, toads, and birds.
Parasites complete all or part of their life cycle in
the pest. Examples include wasps and certain flies.
Insect pathogens such as bacteria, fungi, viruses, and
nematodes can cause insect diseases. The bacteria Ba
cillus thuringiensis (Bt for short) is used in commercial
pesticide sprays.
Genetic controls
Genetic control methods utilize plant breeding for pest
resistance or insect sterilization to affect mating.
Figure 11. Immature and adult lady bug predators.
10
CTAHR — July 2003
Some insects including male fruit flies can be ster
ilized by radiation and released to mate with wild popu
lations. The resulting matings do not produce viable
young and can reduce the pest population.
Some plants are bred to resist insect infestations.
Plant characteristics can affect insect behavior; for ex
ample, trichomes (hairs) on the underside of leaves can
deter insects from feeding or laying eggs.
Plant resistance can affect the biology of the pest,
as when the Bacillus thuringiensis gene is implanted into
the corn genome to control European corn borer. Plant
resistance can also allow a host plant to tolerate the pest
below economic threshold levels.
Regulatory controls
These are usually government-imposed restrictions on
the movement of plants and pests to help prevent un
wanted infestations. Also included is quarantine, or hold
ing of plant material to determine that the material is
pest free. Home gardeners can help by not moving in
fested plants and having plant materials inspected be
fore moving them into pest-free areas.
Chemical controls
Insecticides can be a part of the integrated pest manage
ment system if other IPM methods are not sufficient for
pest control.
If pesticides are necessary, gardeners should use the
least toxic pesticide that will control the pest. New pes
ticides include more environmentally safe materials.
Measure and use only the amount of pesticide nec
essary to cover the targeted plants.
Calibrate your sprayer to determine the amount of
water necessary to apply the pesticide to plants.
Insect pest control questions and strategies
• Identify the insect pest you are dealing with.
• Learn the life cycle of the pest—what is the suscep
tible stage to best apply control measures?
• Learn the host plant or living conditions of the pest—
are there alternate host plants? Does the insect prefer
dry conditions or warm weather?
• Determine the extent of the problem—is the infesta
tion serious enough to cause significant damage? Are
control measures cost-effective?
• Determine which control measures are the most ef
fective—consider biological control, less toxic and
IP- 13
CTAHR — July 2003
environmentally safe pesticides, and applicator safety.
• Learn the proper use of pesticide application equip
ment.
• Avoid insect pest overexposure to pesticides, which
may reduce effectiveness and create resistance.
Gardeners can obtain more information from other
publications and resources of the Cooperative Exten
sion Service of University of Hawaii’s College of Tropi
cal Agriculture and Human Resources. The Web site
www.ctahr.hawaii.edu includes many publications at
www.ctahr.hawaii.edu/freepubs, as well as an insect pest
database, Knowledge Master, which can be found at
www.extento.hawaii.edu. The database includes more
information on insect life cycles and describes additional
nonchemical control methods.
11
Insect Pests
Jan. 2004
IP-14
Mangosteen Caterpillar
Mike A. Nagao1, Heather M. C. Leite1, Arnold H. Hara2, and Ruth Y. Niino-DuPonte2
Departments of 1Tropical Plant and Soil Sciences and 2Plant and Environmental Protection Sciences,
Beaumont Agricultural Research Center, Hilo
A
caterpillar that causes extensive damage to young
leaves of mangosteen trees in Hawaii has been
identified as Stictoptera cuculioides Guenee (Lepi
doptera: Noctuidae), formerly called S. subobliqua
(Walker). The mangosteen caterpillar was first recorded
in Hawaii in 1949 from larvae and adult specimens ob
tained in Honolulu in 1948.
Distribution
This noctuid moth was first described in Sri Lanka and
has been reported in India, Thailand, Singapore, Malay
sia, Papua New Guinea, and Guam. In Hawaii, the man
gosteen caterpillar is found on the islands of Oahu, Ha
waii, Maui, and Molokai.
Hosts
In addition to mangosteen (Garcinia mangostana), S.
cuculioides feeds on related latex-bearing plants of the
Guttiferae family including Garcinia cambogia, mammee
apple (Mammea americana), kamani (Calophyllum ino
phyllum), autograph tree (Clusia rosea), Ochrocarpus
obovalis, and O. excelsus (synonym, Mammea odorata).
Damage
The caterpillar feeds upon emerging leaves and shoot tips
of the host plant, causing extensive defoliation of new
flushes (Fig. 1), often leaving only the leaves’ midribs. A
single caterpillar as small as 1⁄4 inch (0.6 cm) long can
cause significant damage to tender, young leaves. Due to
their nocturnal feeding behavior, the caterpillars can be
inconspicuous until the damage is severe.
Behavior
Mangosteen caterpillars are active at night but can be
observed feeding on young leaves until early or mid
morning. During later daylight hours, they retreat into
the denser parts of the tree canopy, where they are not
easily detected. Under laboratory conditions, the cater
pillars hide during the day under mangosteen leaves left
in their cage, and they are most active during the early
evening. Prior to pupation, the caterpillars burrow into
the soil or hide under leaves in dark, shaded areas to
develop cocoons.
Figure 1. Damage to mangosteen foliage caused by Stictoptera cuculioides larvae: left, evidence of caterpillar feeding
on tender, new leaves; right, the remaining leaf midribs.
IP- 14
CTAHR — Jan. 2004
Figure 3. S. cuculioides pupa (actual size 1⁄2–5⁄8 inch [1.3–
1.6 cm] long and 1⁄4 inch [0.6 cm] wide).
Figure 4. S. cuculioides adults.
Figure 2. Color variations of the Stictoptera cuculioides
caterpillar (larva); actual size 1–2 inches (2.5–5.0 cm).
Life cycle
Few reports on the life cycle of the mangosteen cater
pillar have been published. Both the larval (caterpillar)
and adult stages of S. cuculioides are variable in size and
color. The caterpillar color ranges from light green with
black or maroon spots and white stripes to dark purple
with white stripes and dots just before pupation (Fig. 2),
at which time the last larval instar is 1–2 inches (2.5–5.0
cm) long.
2
Pupation occurs in the soil. The pupa (cocoon) is
dark brown, 1⁄2–5⁄8 inch (1.3–1.6 cm) long, and 1⁄4 inch
(0.6 cm) wide (Fig. 3).
The adult moth is brown but can vary in color tone
and pattern (Fig. 4). The adult male appears to have a
more ornate wing pattern and a larger abdomen com
pared to the female.
Previous reports indicate that the larval stage aver
ages 15 days and pupation lasts 10–12 days. Under labo
ratory conditions (69.6°F [20.9°C] minimum, 76.8°F
[24.9°C] maximum), the duration of the pupal stage can
extend to as long as 18–20 days. There are no reports on
the duration of the adult moth stage.
Management
Growers should monitor new flushes as they emerge for
evidence of feeding damage. Insecticides containing
Bacillus thuringiensis are effective in controlling leaf
eating caterpillars, including S. cuculioides. Azadirachtin
IP- 14
CTAHR — Jan. 2004
(neem extract) is reported to provide effective control
in Thailand. Consult product labels for information on
application rates and pre-harvest intervals. No biocontrol
agents have been detected on mangosteen caterpillar
infestations in Hawaii.
References
Ooi, P.A.C., A. Winotai, and J.E. Pena. 2002. Pests of
minor tropical fruits. In: J. Pena, J. Sharp, and M.
Wysoki (eds), Tropical fruit pests and pollinators:
biology, economic importance, natural enemies and
control. CAB International Publishing, Wallingford,
Oxfordshire, UK. pp. 315–330.
Hawaii Department of Agriculture. 2001. Heu, R. (ed).
Distribution and host records of agricultural pests and
other organisms in Hawaii. Survey Program, Plant
Pest Control Branch, Plant Industry Division. p. 61
Zimmerman, E.C. 1958. Insects of Hawai’i. Vol. 7,
Macrolepidoptera. University of Hawai’i Press, Ho
nolulu. pp. 345-347.
Acknowledgements
The authors would like to thank Shin Matayoshi, Ha
waii Department of Agriculture (retired); Dick Tsuda,
UH CTAHR; and Dr. Surmsuk Salakpetch, Chantaburi
Horticultural Research Center, for their contributions to
this publication.
3
Insect Pests
Jan. 2004
IP-15
Hopper Burn on Papaya
Caused by the Stevens Leafhopper
Richard H. Ebesu, Department of Plant and Environmental Protection Sciences
S
tevens leafhopper, Empoasca stevensi, can be a
serious pest of papaya. The leafhoppers are found
mostly on the underside of the leaves. They feed on the
plant sap, causing a drying of the leaf tissue called “hop
per burn.” The leafhopper releases saliva into the plant
tissue as it inserts its needle-like stylet mouthparts. The
saliva is toxic to the plant; the leaves turn yellow, their
edges dry and their tissue dies, and the plant becomes
stunted (Figure 1). Young plants are more susceptible,
yet plants of all ages are attacked. The red-fleshed com
mercial papaya cultivars like ‘Sunrise’ are more suscep
tible to hopper burn than the yellow-fleshed cultivars
like ‘Waimanalo Low-Bearing’ and ‘Kapoho’, although
some yellow-fleshed cultivars (notably ‘Line 8’) may
be susceptible. Common symptoms of leafhopper feed
ing are puncture marks along the leaf veins and petiole
and the resulting bleeding of milky white latex from the
plant. The plant usually recovers after removal of the
leafhoppers, but large populations of leafhoppers can
severely damage the plant.
The winged adult leafhopper is about 1⁄8 inch long
and slender, less than 1⁄32 inch wide. It is light yellowish
green with two longitudinal white stripes on top of its
thorax, just behind the head (Figure 2). The immature
stages (nymphs) are light green and look like the adults
only they are smaller and without wings. The leafhop
pers normally run quickly or jump when the leaf is turned
over to observe the underside.
The female lays her eggs singly, mostly in the veins
on the underside of the leaf. Usually, only the puncture
wound where the female laid the egg can be seen. On
average, the eggs take 10 days to hatch and the imma
ture leafhoppers take 12–15 days to complete five growth
stages before turning into adults. On becoming an adult,
the female lays her first egg after 7 days. Females live
for an average of 6 weeks, producing an average of seven
Figure 1. Severe damage to a papaya plant caused by
Stevens leafhopper feeding.
Figure 2. Stevens leafhopper adult, about 1⁄8 inch long.
eggs per week. The complete life cycle of a female will
take about 26 days.
The Stevens leafhopper is very similar to the south
ern garden leafhopper (Empoasca solana), which is also
light green but is slightly longer than the Stevens leaf
hopper. The southern garden leafhopper is found on
many plants including green beans, spiny amaranth, and
IP- 15
Hopper Burn on Papaya Caused by the Stevens Leafhopper
the weed black nightshade (or popolo, Solanum nigrum).
It has also been reported on papaya but it does not seem
to be the major cause of hopper burn on papaya.
Host plants
The Stevens leafhopper may be found on papaya
throughout the year but appear to be most damaging in
the warm summer months when the populations are at
their highest. Besides papaya, it has been reported on
cowpea, plumeria, lima bean, and the Mexican fire plant
(Euphorbia heterophylla). It has been known to rest on
the weed Sigesbeckia orientalis.
Control
There are no known biological control agents for the
Stevens leafhopper. General predators such as spiders
and small wasps may eat them, and a fungal disease can
infect them if conditions are right. Insecticides regis
tered for papaya may help to reduce leafhopper popula
tions provided that spray coverage is adequate. Papaya
2
CTAHR — Jan. 2004
plants are sensitive to many pesticides and the spreader
stickers used with them, and users should test products
for potential damage before proceeding with wide-scale
applications. Control of leafhoppers when their popula
tions are small is easier than after waiting until they are
present in large numbers. Leafhopper populations can
be monitored with yellow sticky traps spaced among
the plants.
References
Ebesu, R.H. 1985. The biology of the leafhopper
Empoasca stevensi Young (Homoptera: Cicadellidae)
and its toxicity to papaya. M.S. Thesis, Entomology,
University of Hawaii at Manoa, August 1985.
Mau, R.F.L., L. Gusukuma-Minuto, R. Ebesu, and R.
Hamasaki. 1994. Control of the Stevens leafhopper
on papaya. In: Proceedings, 30th Annual Hawaii Pa
paya Industry Association Conference. College of
Tropical Agriculture and Human Resources, Univer
sity of Hawaii.
Insect Pests
May 2004
IP-16
ALIEN
DRAFT
PEST
ALERT!
Identifying the Little Fire Ant
A New Invasive Species on Kaua‘i
Hawai‘i Ant Group; U.S. Fish and Wildlife Service; Hawai‘i Department of Agriculture (HDOA), Plant Pest Control
Branch; University of Hawai‘i, Pacific Cooperative Studies Unit and Department of Plant and Environmental Protec
tion Sciences, College of Tropical Agriculture and Human Resources; Kaua‘i Invasive Species Committee (KISC)
W
e are in the process of eradicating an infestation
of the little fire ant (LFA) in the Kalihiwai area
of Kaua‘i. We need the help of everyone on Kaua‘i to
report any ants they find that match this ant’s descrip
tion. With your help, we can keep Kaua‘i LFA-free.
Background
Since 1999 when it was first collected at Hawaiian Para
dise Park in the Puna area on Hawai‘i, over 30 LFA in
festations have been found on the Big Island. Contain
ment actions are being taken, but limited resources and
personnel, and pesticide label use restrictions, have made
it difficult to eradicate all the infestations there.
Beginning in 1999, HDOA has enforced quarantine
regulations to prevent shipment of infested potted plants
from the Big Island. However, at least one infestation at
Kalihiwai on Kaua‘i apparently was started from such a
shipment before the quarantine, and there may be oth
ers that have yet to be reported. No infestations are
known on any other islands in the state.
Identification and distribution
Little fire ant’s scientific name is Wasmannia auropunc
tata (Roger) (Hymenoptera: Formicidae).
• Little fire ants are tiny—1⁄16 inch long—pale orange,
and slow moving.
• They are found in South America, the West Indies,
warmer regions of Mexico, West Africa, Galapagos
Islands, New Caledonia, and the Solomon Islands.
• In the USA, in addition to its presence in Hawai‘i,
the little fire ant is common in southern Florida.
Actual length, 1⁄16 inch
Head
Little fire ant
worker
• Its sting produces large, painful, raised, red welts.
• Irritation from the sting lasts several days, aching pain
fully at first and later itching intensely in spells.
•
Although not quick to sting when handled, the LFA will
do so if trapped beneath clothing.
• LFA may also sting animals (livestock, pets, wildlife).
Not to be confused with the tropical fire ant
The tropical fire ant, Solenopsis geminata, is a stinging
red ant common in Hawai‘i.
• Tropical fire ants are 2– Tropical fire ant (“red ant”)
3 times longer than LFA:
Head
1
⁄8–1⁄4 inch (3–6 mm).
• Tropical fire ants will
have a few larger work
ers with large, square
1
⁄8 inch
shaped heads.
• LFA workers are all the
same size, about as long
Little fire ant
as a penny is thick.
UH–CTAHR, May 2004
Identifying the Little Fire Ant on Kaua‘i
ALIEN
PEST
ALERT!
Help keep Kaua‘i free of little fire ants!
If you think you have LFA on your property on Kaua‘i,
please call HDOA in Lïhu‘e at 274-3069. Do not apply any toxic baits or insecticides until HDOA or KISC person
nel survey the infestation. They can advise you on how best to control or eradicate the ant infestation. Hydramethylnon
based granular ant bait has been successfully used to contain or eradicate some LFA infestations on the Big Island.
Checking for presence of the little fire ant
Pick up the chopstick very carefully
1. Smear a very thin 2. Place the chopstick with peanut butter 3.
to avoid dislodging ants, and examine the
coat of peanut butter on
one end of a wooden
chopstick (which can be
painted day-glo orange
for ease of locating).
in an area where you see ants, preferably in
the shade, at the base of a tree, etc.; leave it
out for about 1 hour.
In a pot
Near a shadehouse
4.
Drop off or mail the
ants for identification to:
Hawai‘i Dept. of Agriculture
Plant Pest Control Branch
(Attn: Craig Kaneshige)
4398A Pua Loke St.
Lïhu‘e, HI 96766-1673
Phone number: 274-3069.
ants on the peanut butter.
• Are they orange or red?
• Are they no bigger than 1⁄16 inch?
If you can answer Yes to both questions,
then you may have little fire ants.
Put the chopstick with the ants into a zip
top bag. Write your name, location, and
phone number on the bag. Place the bag
into the freezer overnight to kill the ants.
5. It is very important that
you do not apply any toxic ant
bait or spray at the site until
the location is mapped and the
ant is identified by HDOA.
Doing so will suppress the
ants and make it more diffi
cult to map them before con
trol efforts are started.
For more information, please visit these websites:
Hawai‘i Department of Agriculture—http://www.hawaiiag.org/hdoa/npa/npa99-02-lfireant.pdf
Hawai‘i Ant Group—http://hbs.bishopmuseum.org/ants
Kaua‘i Invasive Species Committee—http://www.hear.org/KISC/index.html
Photo credits: p. 1, top, W. Nagamine, HDOA; p. 1, bottom, C. Hirayama, HDOA, Hilo; p. 2, C. Hirayama and P. Conant, HDOA, Hilo, except for step 4 photo, K. Gundersen.
2
Quarantine Pests
Commonly Found in Shipments from Hawaii
Presence of the insects and related organisms shown below will result in rejection of plant shipments from Hawaii to the U.S. mainland.
ACTUAL SIZE
ACTUAL SIZE < 1 MM
ACTUAL SIZE
ACTUAL SIZE
ACTUAL SIZE
longlegged ant
bigheaded ant
tiny yellow house ant
whitefooted ant
little fire ant
Anoplolepis gracilipes
Pheidole megacephala
Tapinoma melanocephalum
Technomyrmex albipes
Wasmannia auropunctata
ACTUAL SIZE
ACTUAL SIZE
ACTUAL SIZE
ACTUAL SIZE
ACTUAL SIZE
green scale
nigra scale
hemispherical scale
coconut scale
mining scale
Coccus viridis
Parasaissetia nigra
Saissetia coffeae
Aspidiotus destructor
Howardia biclavis
Cover removed to reveal female and eggs
Scale cover
ACTUAL SIZE
ACTUAL SIZE
ACTUAL SIZE < 1 MM
ACTUAL SIZE
ACTUAL SIZE (OF COVER)
black thread scale
an armored scale
ti scale
hibiscus scale
magnolia white scale
Ischnaspis longirostris
Lopholeucaspis cockerelli
Pinnaspis buxi
Pinnaspis strachani
Pseudaulacaspis cockerelli
ACTUAL SIZE
ACTUAL SIZE
ACTUAL SIZE
ACTUAL SIZE
ACTUAL SIZE
white peach scale
pink pineapple mealybug
coconut mealybug
palm mealybug
longtailed mealybug
Pseudaulacaspis pentagona
Dysmicoccus brevipes
Nipaecoccus nipae
Palmicultor palmarum
Pseudococcus longispinus
Egg mass
ACTUAL SIZE
ACTUAL SIZE
ACTUAL SIZE
ACTUAL SIZE (ADULT)
ACTUAL SIZE
rhizoecus root mealybug
pink hibiscus mealybug
southern garden leafhopper
a planthopper
torpedo bug
Rhizoecus hibisci
Maconellicoccus hirsutus
Empoasca solana
Kallitaxila granulata
Siphanta acuta
Adult
Pupa
Adult
ACTUAL SIZE (ADULT)
ACTUAL SIZE
a katydid egg in a leaf
ACTUAL SIZE ABOUT AS SHOWN
ACTUAL SIZE (ADULT)
ACTUAL SIZE (ADULT)
ACTUAL SIZE
spiraling whitefly
anthurium whitefly
nettle caterpillar
green garden looper
Aleurodicus dispersus
Aleurotulus anthuricola
Darna pallivitta
Chrysodexis eriosoma
ACTUAL SIZE
marsh slug
a slug
semi slug
cuban slug
a native snail
Deroceras species
Meghimatium striatum
Parmarion martensi
Veronicella cubensis
Tornatellides species
S. Chun, R. Niino-DuPonte, A.H. Hara, and C. Jacobsen
Eggs
Adult Male
Photos by S. Chun, A.H. Hara, W. Nagamine, R.F.L. Mau, B.C. Bushe, V.L. Tenbrink, T.Y. Hata,
P. Conant, and R. Heu; whitefooted ants by R.H. Scheffrahn, Univ. of Florida; southern garden
leafhopper from Department of Entomology, Texas A&M University; semi slug by R.G.
Hollingsworth, USDA-ARSP-BARC
CTAHR publication IP-18, Revised August 2011
coqui frog
Eleutherodactylus coqui
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in cooperation with the U.S. Department of
Agriculture, under the Director/Dean, Cooperative Extension Service/CTAHR, University of Hawaii at Manoa, Honolulu, Hawaii 96822. An Equal Opportunity / Affirmative Action Institution providing programs and services to
the people of Hawaii without regard to race, sex, age, religion, color, national origin, ancestry, disability, marital status, arrest and court record, sexual orientation, or veteran status. CTAHR publications can be found on the
Web site <http://www.ctahr.hawaii.edu> or ordered by calling 808-956-7046 or sending e-mail to [email protected].
How to Recognize Symptoms
of Aster Yellows in Watercress
Healthy
Infected
Insect Vector
Yellowing in watercress
due to an uncertain cause was first
reported by a farmer in September, 2000. After extensive efforts, including laboratory
analyses and greenhouse and field tests, CTAHR’s virology laboratory identified a
phytoplasma in watercress in October, 2001. This pathogen appears to be closely related
to two other phytoplasmas, western North American aster yellows and onion yellows
from Asia. Phytoplasmas are a group of microscopic organisms that cause over 700
diseases in plants. Phytoplasmas grow and multiply within host plants and insect vec
tors. In host plants, phytoplasmas are found only in the phloem tissue of leaves, stems,
and roots. When the concentration of phytoplasmas within the plant reaches a certain
level, it is believed to cause hormonal imbalance, resulting in the development of symp
toms such as chlorotic leaves, stunting, flower petals changing to a green color (phyllody
or virescence), and witches-broom (shoot proliferation). This is the second phytoplasma
to be identified in Hawai‘i; the first was on a native forest tree, ‘a‘ali‘i, Dodonaea viscosa.
In October 2001, the Hawaii Department of Agriculture confirmed the presence of a
recently introduced leafhopper vector of phytoplasma in watercress. This leafhopper is
known locally as the watercress leafhopper. It has not been formally identified but
appears to be closely related to the aster yellows leafhopper, Macrosteles fascifons.
The leafhopper feeds by inserting its mouthparts into the watercress phloem tissue.
After a noninfected leafhopper feeds on a phytoplasma-infected watercress plant, it takes
about 2–4 weeks for the insect to become a persistent vector. Then this leafhopper can
infect other noninfected watercress plants. It may take several weeks or longer before
plant symptoms such as chlorosis or shoot proliferation appear on a newly infected
plant, and during this time, noninfected leafhoppers can acquire the phytoplasma by
feeding on the symptomless infected plant. Because watercress plants can be infected
without showing symptoms, watercress from the Aiea-Waipahu production areas should
not be used as planting material for other areas on Oahu or the Neighbor Islands. Also,
these plants can carry leafhopper eggs within the leaves, petioles, and stems.
Phytoplasmas can spread via (1) watercress leafhoppers, (2) using infected plant
ing material, (3) grafting, and (4) parasitic plants (e.g., dodder). Phytoplasmas cannot
be transmitted by rubbing sap from infected plants onto healthy plants or by cutting
tools used in farming practices. Phytoplasmas are not known to be transmitted by seeds.
Best Management Practices
1) Start with noninfected planting material.
2) Manage and completely control the watercress leafhopper in the watercress
and in borders surrounding the field.
3) Aggressively rogue infected watercress plants.
4) Control all known weed hosts of the phytoplasma both within and around the
borders of the farm (see weed host photos, far right).
5) Fertilize periodically with a high-nitrogen, slow-release fertilizer.
6) Do not transport watercress planting material outside of the Aiea–Waipahu
watercress production area.
7) Backyard gardeners and new growers should not plant watercress unless they
know that the planting material is free of the phytoplasma and the leafhopper.
watercress leafhopper
(side view)
actual size: 2 mm
watercress leafhopper
(top view)
Weed Hosts
Leaves
parrot’s feather
Myriophyllum brasiliense
Flora’s paintbrush
Emilia sonchifolia
sow thistle
Sonchus species
broadleaved plantain
Plantago major
amaranth
Amaranthhus species
false daisy
Eclipta prostrata
Roots
Shoots
Written by Steve Fukuda1, Wayne Borth2, Rodrigo Almeida2, Randy
Hamasaki2, John McHugh3, Ron Hew4, Bernarr Kumashiro4, Mike
Kawate2, Desmond Ogata5, and Jari Sugano2
CTAHR Departments of 1Tropical Plant and Soil Sciences, 2Plant and Environmental
Protection Sciences; 3Crop Care Hawaii; 4Hawai‘i Department of Agriculture; 5CTAHR
Agricultural Diagnostic Services Center
Field
Leafhopper photos:
Walter Nagamine and Ron Heu, Hawai‘i Department of Agriculture
CTAHR Insect Pests publication IP-20, October, 2004
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in cooperation with the U.S. Department of Agriculture. Andrew G.
Hashimoto, Director/Dean, Cooperative Extension Service/CTAHR, University of Hawaii at Manoa, Honolulu, Hawaii 96822. An Equal Opportunity / Affirmative Action Institution providing programs and services to the people of Hawaii without regard
to race, sex, age, religion, color, national origin, ancestry, disability, marital status, arrest and court record, sexual orientation, or veteran status. CTAHR publications can be found on the Web site <http://www.ctahr.hawaii.edu> or ordered by calling 808956-7046 or sending e-mail to [email protected].
Stinging Nettle Caterpillar
Watch the release of wasps to help
control the stinging nettle caterpillar in
Hawai‘i! Scan with a QR code smartphone app or visit:
http://www.bigislandvideonews.com/2010/06/17/video-waspreleased-to-help-stinging-caterpillar-fight/ Video: David Corrigan
Report new infestations in areas other than Hilo and
Puna districts to the State Pest Hotline: 643-PEST
(643-7378)
For more information, contact:
Hawai‘i Department of Agriculture
Hilo: (16 E. Lanikaula St.) 974-4146
(or UH-CTAHR, 875 Komohana St., 981-5199)
Kahului: (635 Mua St.) 873-3962
Honolulu: (1428 S. King St.) 973-9525
Lïhu‘e: (4398 Pua Loke St.) 274-3072
Report any new infestations of stinging nettle
caterpillar in areas other than Hilo and Puna
to the State Pest Hotline, 643-PEST (6437378), or contact the Hawai‘i Department
of Agriculture (see the back panel of this
brochure for the branch nearest you).
Stinging Nettle
Caterpillar
Darna pallivitta
What to do if you are stung
• Avoid further contact with the caterpillar’s spines.
• Wash the area immediately with soap and water
Authors
Darna pallivitta
Stacey Chun, Arnold Hara,Ruth Niino-DuPonte
UH-CTAHR Komohana Research and Extension
Complex, Hilo
The stinging nettle caterpillar is of major
concern because of its painful sting, voracious appetite, lengthy larval feeding stage
(2 months), high fecundity (480 eggs per female),
and wide host range. A heavy infestation can
defoliate a potted plant in just a few days.
Walter Nagamine,1 Patrick Conant,2
Clyde Hirayama2
Hawai‘i Department of Agriculture, Plant Pest Control
Branch, 1Honolulu, 2Hilo.
Photos of larva and predator wasps by W. Nagamine; other photos
by S. Chun and A. Hara.
Caution: Pesticide use is governed by state and federal regulations.
Read the pesticide label to ensure that the intended use is included
on it, and follow all label directions.
References
Cock, M.J.W., H.C.J. Godfray, and J.D. Holloway (eds). 1987. Slug
and nettle caterpillars. CAB International, Wallingford, UK.
Conant P., A.H. Hara, L.M.Nakahara, R.A. Heu. Nettle caterpillar.
New Pest Advisory no. 01-03, March 2002 (revision). Hawai‘i
Department of Agriculture.
Nagamine, W.T. and M.E. Epstein. 2007. Chronicles of Darna
pallivitta (Moore 1887) (Lepidoptera: Limacodidae): biology
and larval morphology of a new pest in Hawaii. The Pan-Pacific
Entomologist 83(2): 120-135.
PEST
ALERT
College of Tropical Agriculture
and Human Resources
University of Hawai‘i at Mänoa
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and
issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914,
in cooperation with the U.S. Department of Agriculture. Andrew G. Hashimoto, Director/
Dean, Cooperative Extension Service/CTAHR, University of Hawai‘i at Mänoa, Honolulu,
Hawai‘i 96822. An equal opportunity/affirmative action institution providing programs and
services to the people of Hawai‘i without regard to race, sex, age, religion, color, national
origin, ancestry, disability, marital status, arrest and court record, sexual orientation, or
status as a covered veteran. CTAHR publications can be found on the Web site <http://
www.ctahr.hawaii.edu/freepubs>. Publication IP-22, Sept. 2005. Revised July 2011.
to reduce initial pain.
oral antihistamine may stop itching and
swelling.
• Hydrocortisone cream may also stop itching and
swelling.
• Get medical attention immediately if you
experience difficulty breathing or are stung
in the eye.
• Skin reactions vary from a red welt to severe
swelling lasting a couple of days.
• An
College of Tropical Agriculture
and Human Resources
University of Hawai‘i at Mänoa
S
tinging nettle caterpillar was first discovered in
Hawai‘i in September 2001, at a foliage nursery
in Pana‘ewa on the island of Hawai‘i. Nursery
workers there experienced an unusual burning and
itching sensation on their skin after handling rhapis
palms. Specimens sent to the Smithsonian Institution
were identified as Darna pallivitta Moore. The insect
probably arrived from Taiwan and is also found in
China, Thailand, Malaysia, and Indonesia.
Currently, nettle caterpillar infestations have been
reported from Volcano to Ninole in East Hawai‘i, with
most of lower Puna and nearly all of south Hilo district
having infestations. In West Hawai‘i, D. pallivitta has
been detected in Kailua-Kona, Ke‘ähole, Ka‘üpülehu,
and Köhala. Infestations on O‘ahu include central
O‘ahu (Mililani, Mililani Mauka, Waipi‘o-Gentry,
Waikele), Makakilo, and Waimänalo. On Maui, infested
areas include Ha‘ikü, Pä‘ia, Makawao, Wailuku, and
Kïhei. During 2010, the nettle caterpillar was detected
on Kaua‘i (Lïhu‘e, Kapa‘a, and Kïlauea areas), but has
yet to be recorded on Moloka‘i or Läna‘i.
The caterpillar’s spiny
hairs release an irritant
on contact
Harm to humans
The nettle caterpillar’s stinging, spiny hairs have a physical
effect on human skin similar to that of fiberglass. In addition,
the spines release an irritant (a mixture of histamines)
produced by a poison gland. The irritant causes the skin to
burn and itch. If spines get into the eyes, the irritation can
be acute; seek medical attention quickly.
Identifying and Managing Stinging Nettle Caterpillars
Host Plants
In Hawai‘i, the nettle caterpillar has been found on more
than 30 plants including palms, pasture and ornamental
grasses, weeds, and foliage plants. The nursery industry
has a very low tolerance for the nettle caterpillar—any
feeding by the larvae significantly damages and reduces
the value of ornamental and landscape plants. Many of
the host plants are of high economic value for export and
are common in residential and commercial landscaping.
The pest has been observed to complete its life cycle
on palms, including areca, fishtail, manila, rhapis, phoenix,
and coconut; it also reproduces on dracaena (cultivars
‘Lisa,’ ‘Compacta,’ and ‘Massangeana’) and on starfruit,
ti, iris, coffee, honohono grass, the beach pea (indigenous
Vigna marina), and the endemic mamaki.
The caterpillar has been observed feeding (but not reproducing) on many other plants, including bamboo orchid,
banana, ‘Pink Quill’ bromeliad, chickweed, Chinese star
jasmine, cigar plant, rabbitsfoot fern, gardenia, glory bush
(Tibouchina), ‘Golden Glory’ perennial peanut, californiagrass, hilograss, mondograss, napiergrass, vaseygrass,
wainakugrass, guava, Koster’s curse, macadamia, maunaloa
vine, monstera, ponytail palm, red and shampoo gingers,
sleeping grass, Spanish clover (silverleaf desmodium),
walking iris, wedelia, whaleback, and the endemics maile
and wiliwili (data gathered by UH-CTAHR and HDOA).
Damage by feeding of large larvae on (clockwise from upper
left) rhapis, coconut, dracaena, mondograss, and ti.
LIfe Cycle
Life cycle
The nettle caterpillar’s life span from egg to adult is 75–99
days, depending on the number of larval stages (instars),
which ranges from 8 to 11 (45–72 days total). Adult
female and male moths live for approximately 10 and 11
days, respectively. As the larvae develop over the 7-day
incubation period, the C-shaped embryos are clearly
visible. When the larvae are ready to pupate, they migrate
toward the base of the host plant to find protected crevices
in dried leaves and overlapping plant parts, and they often
pupate in clusters. The larva’s underside darkens to orange
just before pupation. The prepupa spins brown silk around
itself, eventually forming a hardened outer shell. The round
cocoons are 5⁄8 inch (16 mm) long, and pupation occurs
within the cocoon after 5 days.
Eggs
The female adult moth deposits eggs in small clusters, a
line, or singly, usually on the undersides of older leaves.
Eggs are flattened, transparent ovals, 1⁄32 inch (0.8 mm)
wide and 1⁄16 inch (1.6 mm) long, appearing as a glassy
sheen on the leaf surface that can easily be overlooked.
Eggs
Control Methods
Newly hatched
larvae
Incubating
larvae
Fully developed larva,
about 1 inch long
Prepupa
An early larva
on a quarter
Pupa
Larva
The larva can be up to 1 inch (25 mm) long and is covered
with many rows of stinging spines. Larvae vary from white
to light gray, with a dark longitudinal stripe down the back.
Prepupa, pupa
Larvae begin feeding 2 days after hatching. Onset of
pupation depends on food availability and environmental
conditions. The pupal period ranges from 17 to 21 days.
Smaller larvae cause
damage by feeding on the
leaf surface, creating a
“windowpane” effect.
Larvae often pupate in clusters in sheltered spots at the
base of the host plant.
Adult
The adult moth is approximately ½ inch (12.7 mm)
long. The forewing is divided
by a white diagonal marking,
with the upper portion rustcolored and the lower portion
lighter brown; the hind wings
are uniform light brown.
These nocturnal moths have
not been observed feeding.
Mating begins about two
days after emergence. During
the day they are inactive
and retreat into vegetation,
usually in an upside-down,
perching position.
Adult moth
Adults, resting
Male
Cultural control
Control weeds and modify landscape plantings to limit
caterpillar food availability. To avoid transporting the
eggs, which are difficult to detect, to new areas, don’t
bring in known host plants from any infested area. Ti
leaf, mondograss and related Liriope groundcover, and
palms are preferred host plants of nettle caterpillar. Peak
caterpillar populations occur in late summer, so trim these
host plants before then and avoid contact during that time.
Chemical control
Some pesticides (pyrethroid, organophosphate, carbamate, and microbial types including Bacillus thuringiensis, or Bt) are effective against the larval stage of
the nettle caterpillar. Consult a UH-CTAHR Cooperative
Extension Service agent or an agricultural products
professional for help in choosing an insecticide.
Biological control
HDOA staff discovered a locally established trichogrammatid wasp depositing its eggs into D. pallivitta eggs,
which provide a food source for the wasp larvae, which
eventually emerge as adults. This wasp, however, has had
only limited effect on the nettle caterpillar population on
Hawai‘i. Therefore, HDOA has worked with researchers
in Indonesia and Taiwan to
identify other biological control
agents of D. pallivitta.
Larvae of a wasp (Aroplectrus
dimerus) from Taiwan feed and
develop on the nettle caterpillar,
killing it. These wasps were evaluated and found safe for Trichogrammatid
release in Hawai‘i, and they
are now established on several
islands. A naturally occurring
cytoplasmic polyhedrosis virus
found infecting D. pallivitta
larvae helps control heavy
infestations.
Aroplectrus
Physical control
The adult moth is instinctively attracted to light, so
minimize outdoor lighting at night and use bug-zappers
with ultraviolet bulbs to reduce the numbers of this pest.
Position the unit away from any potential host plants and
under protected eaves, and place a bucket of soapy water
directly beneath it to capture fallen moths.
Female
Insect Pests
May 2007
IP-25
Evaluating Spiders for Their Potential
To Control Cabbage White Butterflies (Pieris rapae)
Cerruti R2 Hooks,a Raju R. Pandey,b and Marshall W. Johnsonc
a
CTAHR Department of Plant and Environmental Protection Sciences; bHimalayan College of Agricultural Sciences and Technology, Kathmandu, Nepal; cDepartment of Entomology, University of California, Riverside
Summary
A field experiment was conducted three times during
two seasons (twice in winter and once in spring) to
evaluate the impact of spiders on the survival of the cabbage white butterfly, Pieris rapae (= Artogeia rapae).
The proportion of P. rapae eggs surviving to the first
caterpillar stage was significantly reduced on spider
treatment plants compared to check treatment plants.
During the three experiments, the percentage of P. rapae
eggs surviving to the fifth caterpillar stage was increased
1.7-, 2.7-, and 1.3-fold, respectively, on check plants
compared to spider plants. Additionally, by completion
of the the experiment, above-ground plant biomass of
spider-“protected” plants was increased by 80, 121, and
28 percent compared to check plants.
Introduction
Although several studies have shown that spiders can
significantly reduce insect pest populations and the associated crop damage (Agnew and Smith 1989, Hooks
et al. 2003), their ability to suppress insect pest populations and enhance plant productivity has received limited
attention in cropping systems. On several occasions,
spiders were observed feeding on eggs and caterpillars of
lepidoptera prey inhabiting broccoli (Brassica olearacea
L.) plants, and although their densities were recorded,
no attempt was made to quantify their predatory impact
(Hooks and Johnson 2002). Hooks et al. (2003) found
significantly fewer large P. rapae caterpillars on plants
where spiders were allowed to forage freely, compared
to control plants in which spiders were removed daily.
However, during that study the amount of mortality
spiders inflicted upon P. rapae was not estimated. Therefore, field experiments reported here were conducted to
quantify the impact of spiders on P. rapae’s survivorship and broccoli plant biomass. The objective of this
study was to address two questions: (1) Does caterpillar survival differ on plants containing spiders? (2) Do
spiders indirectly increase broccoli plant size through
suppression of caterpillars?
Procedures
Experimental design
Three field trials were conducted to assess the impact
of spiders on P. rapae’s survival. Experiments were
conducted during 2003 and 2004 at the University of
Hawai‘i at Mänoa’s Poamoho Research Station. For each
trial, 5-week-old greenhouse-grown broccoli plantlets
were transplanted and randomly assigned to two treatments: (1) spiders present, and (2) a check (spiders removed). Twelve plants were assigned to each treatment
during each trial period.
Spiders were removed daily from check treatment
plants at 10:00 and 13:30 during the duration of each
trial. Immediately after the 13:30 removal, a sleeve cage
constructed of a transparent fabric was gently placed
over each check plant to prevent spiders from foraging
them. During the initial 16 days after planting (DAP),
the cages were removed from the plants daily from 09:00
until 13:30 to allow oviposition on the test plants by P.
rapae.
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in cooperation with the U.S. Department of Agriculture. Andrew G. Hashimoto, Director/Dean, Cooperative Extension Service/CTAHR, University of Hawai‘i at Mänoa, Honolulu, Hawai‘i 96822.
An equal opportunity/affirmative action institution providing programs and services to the people of Hawai‘i without regard to race, sex, age, religion, color, national origin, ancestry, disability, marital status, arrest and court record, sexual orientation, or status as a covered veteran. CTAHR publications can be found on the Web site <http://www.ctahr.hawaii.edu/freepubs>.
UH–CTAHR
IP-25 — May 2007
Sampling
Pieris rapae
The proportion of P. rapae eggs reaching the first
caterpillar stage was assessed during spring 2003 and
winter 2004. Twelve plants were randomly selected and
assigned to the check or spider treatment for each trial.
All P. rapae eggs found on these plants during the initial
16 DAP were counted and recorded, and their location
was marked with a permanent marker. Each egg was
checked daily to determine if it reached the caterpillar
stage. At the experiment’s completion, the percentages
of the 16-day egg cohort reaching the caterpillar stage
were calculated for each treatment.
The percentage of P. rapae eggs reaching their final
(fifth) caterpillar stage was measured on 12 additional
randomly selected check and spider plants during three
field trials (i.e., winter and spring 2003, and winter 2004),
respectively. Similarly, as mentioned above, P. rapae
eggs laid during the initial 16 DAP were monitored to
determine the percentage reaching their final caterpillar
stage.
To determine if whole-plant biomass differed between
check and spider plants, upon the experiment’s completion test plants from each treatment were cut at soil level,
transported to the laboratory, oven-dried, and weighed
to measure above-ground dry vegetative biomass.
During the spring egg mortality experiment, the proportion of P. rapae eggs reaching their first caterpillar
stage was significantly higher on check (83%) compared to spider (56.5%) treatment plants. However, no
significant differences were found during the winter
experiment (Fig. 1). For all three trials, the percentage
of P. rapae eggs reaching their final caterpillar stage
was significantly reduced on spider plants compared to
check plants (Fig. 2).
Statistical analysis
Treatment effects on the number of eggs oviposited,
plant weight, and percentage of eggs reaching the first
and final caterpillar stages were assessed using analysis
of variance (Proc GLM, SAS Institute, Cary, NC 1990).
To fulfill assumptions regarding normality and equal
variances, data were transformed when necessary.
Results
Spiders
Over the course of the trial, spiders removed from the
check plants included Nesticodes rufipes, Oxyopes sp.,
Cheiracanthium mordax Koch, Neoscona oaxacensis
Keyserling, and an unidentified linyphiid. The average
number of spiders found per broccoli leaf during the
three trials ranged from 0.25 to 0.69, 0.17 to 0.60, and
0.17 to 0.46 for the winter 2003, spring 2003, and winter
2004 trials, respectively. During each trial the number
of spiders found per plant increased during the broccoli
growth cycle.
2
Whole plant biomass
During the 2003 trials, the average plant weight was significantly greater for spider plants than for check plants
(Fig. 3). During the 2004 winter experiment, spider
plants were larger than check plants, but the difference
was not statistically significant.
Discussion
During one of the two field trials, the percentage of
eggs reaching the first caterpillar stage was significantly
lower on spider treatment plants compared to check
plants. Furthermore, the proportion of eggs reaching
the fifth instar stage was significantly lower on spider
plants compared to check plants during all three field
trials. Broccoli whole-plant biomass of spider plants was
significantly greater than that of check plants during the
first two field experiments.
Spiders were rarely observed feeding on P. rapae
eggs, but the results suggest that spiders had a significant
impact on P. rapae egg mortality. A number of cabbage
looper (Trichoplusia ni) eggs were also encountered
on spider plants during the spring, but no larva of this
species was observed during the trial, suggesting that
spiders also fed on T. ni eggs. The wandering spider
(Oxyopes sp.) appears to be the most important spider
for suppressing populations of P. rapae and T. ni. Suppression of P. rapae was greatest when populations of
Oxyopes sp. were high and least when they were low.
Acknowledgment
The authors wish to thank the crew at the Poamoho Research Station for their valuable help in the field. This
research was partially funded by the USDA/CSREES,
Special Grant for Tropical and Subtropical Agriculture
Research (T-STAR).
UH–CTAHR
IP-25 — May 2007
Fig. 1.
Percentage
rapae
Percentage
ofof
A.A.
rapae
Percentage
of P.
rapae
st eggs
st1 instar
st
eggs
to
reach
to to
reach
1 instar
eggs
reach
1 instar
Figure 1. Percentage
(± S.E.) of Pieris rapae eggs reaching the first caterpillar stage on spider-removed and spiderFig. 1.
present treatment plants during two field trials. Different letters above a bar for each trial indicate that treatments are
significantly different at the 5% level (P < 0.05).
100
100
Spider removal
Spider
removal
Sp
ider p
resent
Spider present
a
a
80
80
b
b
60
60
83.0
83.0
40
40
89.1
89.1
79.3
79.3
56.5
56.5
20
20
0
0
Spring 2003
Spring 2003
Winter 2004
Winter 2004
Percentage
of
A.rapae
rapae
Percentage
Percentage
ofof
P. A.
rapae
eggs
th
th instar
th 5
eggs
to
reach
5 5instar
instar
eggstotoreach
reach
Figure 2. Percentage (± S.E.) of Pieris rapae eggs to reach the fifth caterpillar stage on spider-removed and spiderFig. 2. plants during three field trials. Different letters above a bar for each trial indicate that treatments are
present treatment
Fig. 2.
significantly different at the 5% level (P < 0.05).
80
80
Spider removal
Spider removal
Spider present
Spider present
a
a
a
a
60
60
40
40
a
a
b
b
bb
72.3
72.3
66.3
66.3
20
20
38.8
38.8
b
b
52.4
52.4
56.2
56.2
19.2
19.2
00
Winter
Winter 2003
2003
S
ring
03
3
Sp
prin
g2
20
00
Winter 2004
2004
Winter
Dry weight/broccoli plant (g)
Brocolli dry weight per plant (g)
Fig.(±
3.S.E.) dry whole-plant biomass on spider-removed and spider-present treatment plants during three
Figure 3. Average
field trials. Different letters above a bar indicate that treatments are significantly different at the 5% level (P < 0.05).
70
Spider removal
Spider present
b
60
50
40
a
30
56.3
b
20
31.2
a
10
21.0
15.3
9.5
11.9
Spring 2003
Winter 2004
0
Winter 2003
3
UH–CTAHR
References and further readings
Agnew, C.W., and J.W. Smith Jr. 1989. Ecology of spiders (Araneae) in a peanut agroecosystem. Environ.
Entomol. 7: 402–404.
Hooks, C.R.R., and M.W. Johnson. 2002. Lepidopteran
pest populations and crop yields in row intercropped
broccoli. Agric. For. Entomol. 4: 117–125.
Hooks, C.R.R., R.R. Pandey, and M.W. Johnson. 2003.
Impact of avian and arthropod predation on lepidopteran caterpillar densities and plant productivity
in an ephemeral agroecosystem. Ecol. Entomol. 28:
522–532.
Hooks, C.R.R., R.R. Pandey, and M.W. Johnson.2006.
Effects of spider presence on Artogeia rapae and host
plant biomass. Agri. Ecosys. Environ. 112: 73–77.
4
IP-25 — May 2007
Insect Pests
May 2007
IP-26
Unlikely Guardians of Cropping Systems:
Can Birds and Spiders Protect Broccoli from Caterpillar Pests?
a
Summary
A field experiment was conducted to examine the impact
of bird and spider predation on lepidopteran caterpillar
densities and broccoli productivity. Densities of Pieris
rapae and Trichoplusia ni large caterpillars and their
post-caterpillar stages were reduced significantly by bird
predation. The abundance of large caterpillars was also
reduced on plants where spiders were allowed to forage
freely. Further, plants foraged by birds, spiders, or birds
plus spiders sustained less feeding damage attributable
to leaf-chewing caterpillars than plants without birds or
spiders (the check). Plants foraged by bird and/or spiders
were also larger than check plants.
Introduction
On the island of O‘ahu, birds and spiders have been casually observed preying on insect pests of Brassica plants.
Their impact on insect pests, especially caterpillars, is
potentially significant but had not been investigated in
Hawai‘i. Thus this study was designed to examine the
impact of bird and spider predation on caterpillars commonly found feeding on Brassica crops and to determine
whether their presence on broccoli, Brassica oleracea L.,
would result in reduced plant consumption by caterpillars
and an associated increase in broccoli plant biomass.
Several studies have shown that birds that feed on
insects, commonly called insectivorous birds, can significantly reduce insect population size (Bock et al. 1992,
Greenberg et al. 2000) and the amount of plant damage
they cause (Sanz 2001). These studies were mainly conducted in perennial plant communities such as temperate
forests and grasslands where birds are likely the top
predators of insect prey. However, insectivorous birds
have a diverse diet and when feeding do not discriminate
between pest and beneficial insects. Thus, it is not safe
to assume that their presence within a cropping system
will result in greater insect pest suppression.
Similar to insectivorous birds, spiders are not preyspecific and may fulfill their dietary needs by feeding
on natural insect pest enemies such as parasitoids and
predators. In some instances, this may cause an increase
in insect pest numbers by reducing natural enemies that
would normally keep their population in check. Further,
spiders have a long generation time compared to most
insect pests. As such, insects can produce offspring at a
much faster pace than spiders. Thus, theoretically speaking, it is believe that spiders are not capable of reducing
insect pests to noticeable levels in cropping systems.
Nevertheless, we were interested in knowing the extent
to which birds and spiders may influence the number of
caterpillar pests on broccoli plants.
It was our belief that birds and spiders found on broccoli plants in Hawai‘i are capable of reducing caterpillar
pests to levels that will result in significantly less plant
damage and an increase in plant size. As such, using a
four-level system inclusive of birds, spiders, lepidopteran
caterpillars, and broccoli plants, two questions were
addressed: (1) Do bird and spider predation reduce lepidopteran caterpillar densities directly and subsequently
increase plant productivity? and (2) Does an assemblage
of birds and spiders reduce insect herbivore densities
more than birds or spiders alone?
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in coopera
tion with the U.S. Department of Agriculture. Andrew G. Hashimoto, Director/Dean, Cooperative Extension Service/CTAHR, University of Hawai‘i at Mänoa, Honolulu, Hawai‘i 96822.
An equal opportunity/affirmative action institution providing programs and services to the people of Hawai‘i without regard to race, sex, age, religion, color, national origin, ancestry, dis
ability, marital status, arrest and court record, sexual orientation, or status as a covered veteran. CTAHR publications can be found on the Web site <http://www.ctahr.hawaii.edu/freepubs>.
UH–CTAHR
Procedures
Study system
The insectivorous bird community within the study
site consisted of the red-crested (Brazilian) cardinal,
Paroaria coronata, and the northern cardinal, Cardinalis cardinalis. Four spider species frequently found on
Brassica plants at the study site, in order of abundance,
include (1) Nesticodes (Theridion) rufipes (Theridiidae),
(2) Oxyopes sp. (Oxyopidae), (3) Neoscona oaxacensis
(Araneidae), and (4) Cheiracanthium mordax (Clubionidae). The tangle web spider, N. rufipes, which spins
a sparse web on the leaf surface, is the most abundant
spider encountered on Brassica plants at the study site,
consistently comprising 90 percent or more of the total
spider fauna.
Brassica plants found at the study site are affected
by a complex of caterpillar pests. Listed in order of
importance, these are the imported cabbage worm,
Pieris rapae =(Artogeia rapae) (Lepidoptera: Pieridae);
the cabbage looper, Trichoplusia ni Hübner (Lepidoptera: Noctuidae); and the diamondback moth, Plutella
xylostella L. (Lepidoptera: Plutellidae). The imported
cabbage worm causes the greatest feeding damage. The
adults are frequently seen searching for food or egg laying sites within agricultural fields or neighboring areas
that contain flowering plants.
Experimental design
The experiment was conducted at the University of
Hawai‘i at Mänoa’s Poamoho Research Station on
O‘ahu during May 2002. Greenhouse-grown broccoli
plants were transplanted and randomly assigned to four
treatments: (1) bird-accessible plants (with daily spider
removal); (2) both bird plus spiders present (no manipulation, plants accessible to birds and spiders); (3) spideraccessible plants (plants enclosed in a cage that allowed
access to plants by spiders but not birds); (4) no birds or
spiders (check plants enclosed in a cage that prevented
access by birds with daily spider removal). Spiders were
removed from the bird-accessible and check plants daily
at ~ 3.5-h intervals beginning at 09:30 and completed
everyday by 19:00 hour. After the final spider removal
task of the day, a sleeve cage was placed over the bird
and check treatment plants to prevent spiders from foraging them at night.
2
IP-26 — May 2007
Sampling
For each treatment plant, all spiders, moth, and butterfly
stages (e.g., eggs, caterpillars, pupae) were identified and
counted to species. Insects and spiders were sampled at
5-day intervals initiating 10 days after planting (DAP).
The caterpillar counts were divided into four size categories (bantam < 0.5 cm, small >0.5 but < 1.0 cm,
medium > 1.0 but <1.5 cm, large > 1.5 cm). However,
because overall counts of cabbage looper caterpillars
were notably low, especially size categories medium
and large, these latter size categories of cabbage loopers were pooled to one size group. Empty cocoons from
which adult moth and butterflies had recently emerged
were also counted and removed to provide an indication
of the number of caterpillars that made it to the adult
stage. Final counts of caterpillars were made during
the harvest period by dissecting the broccoli crown and
counting all caterpillars and cocoons present. Additionally, the remaining plant parts (e.g., leaves, stems, etc.)
were surveyed from top to bottom, and all late-stage
caterpillars and pupae were counted.
All insect and spider count data were analyzed by ANOVA (PROC GLM, SAS Institute, 1990). When the overall
ANOVA was significant, differences among treatment
categories were determined using Fisher’s protected
least significant difference (LSD). Plant damage data
were analyzed using 2 x 2 contingency tables (PROC
FREQ). Treatments were considered significantly different at P < 0.05.
Results
Spider and insect density
On foliage during broccoli growth cycle. The composition of spiders found on broccoli plants during the
study included N. rufipes (91%), Oxyopes sp. (7%), N.
oaxaencis (1.5%), and C. mordax and an unidentified
lyniphiid, which each made up approximately 0.7%.
The abundance of spiders found on bird plus spider,
and spider treatment plants were similar throughout the
experiment (Fig. 1).
The abundance of bantam, small, and medium sized
imported cabbage worm caterpillars were similar among
treatments on most dates. However, beginning 20 DAP,
the density of large caterpillars (> 1.5 cm) was greater
on check plants than plants where birds were allowed to
UH–CTAHR
IP-26 — May 2007
FigureFig.
1. Mean
1. number of spiders per broccoli leaf in four experimental treatments. * indicates when birds were first
observed foraging the study area (18 days after planting). Graph symbols within a period without letters indicate that
Fig.
1. is insignificant at P > 0.05; Fisher’s protected LSD).
the overall
ANOVA
a
Bird
Bird
+ Spider
Bird
Spider
Bird + Spider
Check
Spider
3.0
1
Numb
Number
er of spiders le-af
Numberer
of o
spiders
1
Numb
Number
f spidper
ersleaf
le-af
3.0
2.5
2.5
2.0
a
a
a
Check
2.0
1.5
a
a
a
b
0.5
0.0
0
0.0
b
b
10
10
0
a
a
a
a
1.0
0.5
a
a
a
1.5
1.0
a
a
b
*
b
b
b
b
*
20
20
b
b
b
b
b
30
30
Days After Planting
Days after planting
Days After Planting
b
b
b
50
50
40
40
Fig. 2.
of large
rapae larvae
per leaf
leaf -1
larvae
No.Number
of large
A.P.rapae
No. of large A. rapae larvae leaf -1
Figure 2. Mean number (+ SE) of Pieris rapae large caterpillars (> 1.5 cm) per broccoli leaf in four experimental treatments.
Fig.
2. birds were first observed foraging the study area (18 days after planting). Graph symbols within a period
* indicates
when
without letters indicate the overall ANOVA is insignificant at P > 0.05; Fisher’s protected LSD).
0.18
0.18
0.16
a
Bird
Bird
Bird
+ Spider
Bird + Spider
Spider
Spider
Check
a
Check
0.16
0.14
0.14
0.12
0.12
0.10
0.10
0.08
a
a
a
a
a
a
a
a
0.08
0.06
0.06
0.04
b
0.04
0.02
0.02
0.00
b
*
0
0.00
0
10
10
ab
ab
b
b
b
b
b
b
*20
30
20
30
Days after planting
Days After Planting
Days After Planting
freely forage, and beyond 25 DAP large caterpillar numbers were similar on check and spider treatment plants
(Fig. 2). Additionally, a greater number of medium/large
size category of cabbage looper caterpillars were found
on spider compared to bird plus spider treatment plants
(Fig. 3). The check and bird treatment plants were cov-
b
40
40
b
50
50
ered at night to prevent spiders from accessing these
plants. However, cabbage looper moths mostly lay eggs
at night. As such, their initial egg counts were lower on
these plants compared to plants left uncovered at night.
Whole plant count at end of growth cycle. At harvest
time, a significantly higher number of imported cabbage
3
UH–CTAHR
IP-26 — May 2007
Mean number of T. ni per plant day -1 -1
Mean
number of T. ni per plant day
Mean number of large T. ni per plant per day
Fig.
3.3. Mean number (+ SE) of Trichoplusia ni medium/large caterpillars (> 1.0 cm) per broccoli leaf in four experimental
Figure
Fig.
3.
treatments.
Bars with same letters are not significantly different (P > 0.05, Fisher’s protected LSD).
medium/large
medium/large
a
0.35
0.35
0.3
0.3
0.25
0.25
0.2
0.2
0.15
0.15
0.1
0.1
0.05
0.05
0
0
a
b
b
b
b
b
b
Bird
Bird
Bird + Spider
Bird + Spider
Spider
Spider
Check
Check
Figure 4. Proportion of broccoli plants displaying extensive chewing damage in four treatments. Numbers within and
Fig.
4. of bars indicate the proportion of plants showing greater than 50 to 75% of leaf area (four terminal leaves)
outside
consumed
respectively, in each of four treatments. * indicates that no plants within that treatment sustained extensive
Fig.
4.
defoliation. A significantly higher proportion of check treatment plants sustained extensive damage compared with
other treatments on each date listed (P < 0.05, Fisher’s exact test).
75
75
Bird + Spider
Bird + Spider
Check
36.4
Check
45.5
45.5
36.4
27.3
27.3
50
50
25
25
20
20
25
25
*
*
*
*
30
30
Days
After
Planting
Days after
planting
Days After Planting
*
*
9.1
9.1
18.2
18.2
36.4
36.4
*
* *
* *
9.1
81.8
81.8
*
9.1
0
9.1
9.1
9.1
9.1
72.7
72.7
0
4
Bird
Bird
Spider
Spider
9.1
9.1
18.2
18.2
63.6
63.6
Proportion
of plants
showing
Proportion of plants
showingshowing
Proportion
of plants
extensive
defoliation
extensive
defoliation
extensive defoliation
100
100
35
35
UH–CTAHR
IP-26 — May 2007
Table 1. Mean number of late-stage lepidopterans per brocolli whole plant at plant maturity (inflorescence fully developed)
after exposure to four experimental treatments.
Pieris rapae1
Treatment2
Large caterpillar3
Pre-pupae
Pupae
Empty cocoon
Bird
0.18 + 0.18 b
0.09 + 0.09
0.0 b
0.0
Bird + spider
0.44 + 0.24 b
0.11 + 0.11
0.0 b
0.0
Spider
3.62 + 1.1 a
0.90 + 0.38
4.2 + 0.8 a
0. + 0.22 a
Check
4.70 + 0.46 a
0.0 + 0.22
.0 + 0.22 a
0.9 + 0.28 a
Same letter denotes no significant differences among treatments (P > 0.0), and means followed by columns with no letters indicates the overall
ANOVA is insignificant (Fisher’s Protected LSD).
2
Bird = plants accessible by birds with daily spider removal; Bird + spider = no manipulation, plants accessible to birds and spiders; Spider =
plants enclosed in a cage that allowed access to plants by spiders but not birds; Check = plants enclosed in a cage that prevented access by
birds, with daily spider removal.
3
large = >1. cm
1
worm caterpillars, pupae, and empty cocoons, respectively, were found on spider and check plants compared
to those plants where birds or birds plus spiders were
allowed to forage. Further, pupae and empty cocoons of
the imported cabbage worm were only found on spider
and check plants (Table 1).
Number in broccoli crown at maturity. At harvest, the
imported cabbage worm was the most abundant caterpillar found in the broccoli crowns. The lowest numbers
were found in the crowns of plants were both bird and
spiders were allowed to forage. Large cabbage looper
caterpillars were only found in the heads of spider treatment plants (Table 2).
Plant damage
No plants in which birds were allowed to forage displayed significant insect chewing damage (50 percent
or more of the leaf area missing) in the plants’ terminal
growth area throughout the experiment. The highest
proportion of plants displaying extensive damage was
observed among check plants. The amount of chewing
damage sustained by the other treatments was similar
throughout the experiment (Fig. 4).
Plant biomass
Broccoli head size was greatest on plants where birds
were allowed to forage and smallest on check plants
(Table 3). Significant differences were also found among
treatment plants with respect to whole-plant biomass.
Table 2. Mean numbers of Pieris rapae per broccoli crown
exposed to four experimental treatments.
Pieris rapae1
Treatment
Large caterpillar2
Pre-pupae
Pupae
Bird
4. + 1.34 b
0.2 + 0.12
0.3 + 0.19
Bird + spider
1.7 + 0.3 c
0.0
0.1+ 0.11
Spider
8.2 + 1.69 a
0.3 + 0.1
0. + 0.27
Check
6.9 + 0.3 ab
0.1 + 0.10
0.8 + 0.1
1
Same letter denotes no significant differences among treatments
at the % level (P > 0.0), and means followed by columns with no
letters indicates the overall ANOVA is insignificant (Fisher’s Protected
LSD).
2
large = >1. cm
Similar to head size, the smallest and largest plants by
weight were check and bird plus spider plants, respectively.
Discussion
This field experiment used a natural colonization of
moth and butterfly pests on broccoli plants to determine
the direct effect of birds and spiders on caterpillar pest
densities and their indirect impact on plant productivity.
Both bird and spiders were found to suppress caterpillar
UH–CTAHR
IP-26 — May 2007
Table 3. Mean fresh head weight and dry whole-plant
biomass of brocolli plants exposed to four experimental
treatments.
Weight (± SE)1
Treatment
Head (kg)2
Whole plant (g)3
Bird
0.182 + 0.01
a
13. + 6.7
ab
Bird + spider
0.161 + 0.01
ab
166.2 + 9.7
a
Spider
0.177 + 0.02
a
139.2 + 9.
b
Check
0.112 + 0.02
b
91.2 + 7.7
c
1
Same letter denotes no significant differences among treatments at the % level (P > 0.05; Fisher’s Protected LSD).
2
Multiply kg by 2.2 to obtain pounds and g by 0.03 to obtain ounces.
3
Whole-plant biomass excludes the weight of the crown and plant parts below the soil surface.
numbers, thereby significantly reducing the level of plant
damage. Additionally, plant productivity was greatest for
plants where birds and/or spiders were allowed to freely
forage; however, despite the negative effect of birds and
spiders on caterpillar populations, the combination of
birds and spiders did not suppress caterpillar densities
on the broccoli foliage more significantly than either
predator alone.
In conclusion, several studies have shown that bird
predation can significantly reduce insect herbivore densities in forest ecosystems (Sipura 1999 and reference
therein). The impact of bird predation on insect herbivores and their interaction with other natural enemies in
agricultural systems is potentially great but has received
limited attention (Greenberg et al. 2000). Clearly, insect
6
pathogens, predators, and parasitoids and spiders may
not be the only natural enemies inflicting mortality
among insect pests in cropping systems. Therefore, more
integrated research studies that evaluate the relationship
arthropod natural enemies have with vertebrate predators
such as birds are needed.
Acknowledgments
The authors wish to thank the crew at the Poamoho
Experiment Station for their valuable help in the field.
This research was funded by the USDA/CSREES,
Special Grant for Tropical and Subtropical Agriculture
Research (TSTAR).
References and further reading
Bock, C.E., J.H. Bock, and M.C. Grant. 1992. Effects of
bird predation on grasshopper densities in an Arizona
grassland. Ecology 73 1706–1717.
Hooks, C.R.R., R.R. Pandey, and M.W. Johnson. (2003).
Impact of avian and arthropod predation on lepidopteran caterpillar densities and plant productivity
in an ephemeral agroecosystem. Ecol. Entomol. 28
522–532.
Greenberg, R., P. Bichier, A.C. Angon, C. MacVean, R.,
Perez, and E. Cano. 2000. The impact of avian insectivory on arthropods and leaf damage in some Guatemalan coffee plantations. Ecology 81: 1750–1755.
Sanz, J.J. 2001. Experimentally increased insectivorous bird density results in a reduction of caterpillar
density and leaf damage to Pyrenean oak. Ecol. Res.
16: 387–394.
SAS Institute, 1990. SAS User’s Guide: Statistics. Cary,
NC.
Sipura, M. 1999. Tritrophic interactions: Willows, herbivorous insects and insectivorous birds. Oecol. 121:
537–545.
Insect Pests
May 2007
IP-27
Using Clovers as Living Mulches
To Boost Yields, Suppress Pests, and Augment Spiders
in a Broccoli Agroecosystem
a
Summary
A field study was conducted to examine the influence of
intercropping broccoli (Brassica oleracea L.) with three
living mulches on caterpillar pest and spider densities
and crop yield. Broccoli was grown in bare ground or
intercropped with strawberry clover (Trifolium fragiferum L.), white clover (Trifolium repens L.), or yel
low sweetclover (Melilotus officinalis L.). Lepidopteran
(butterfly and moth) eggs and caterpillar densities were
significantly greater on broccoli in bare-ground plots
compared with broccoli intercropped with clover dur
ing the late broccoli growth cycle. More spiders were
found on bare-ground broccoli during early crop growth;
however, during the later growth period, spider counts
were significantly higher on broccoli in intercropped
plots. The number of insect contaminants found in
harvested broccoli crowns were significantly less in
intercropped than in bare-ground broccoli plots. The
weight of harvested crowns was similar in intercropped
and bare-ground habitats.
Introduction
An established cover crop that is interplanted and grown
with an annual row crop is known as a living mulch.
Living mulches can provide many benefits to a crop
ping habitat, including weed control, reduced erosion,
enhanced fertility, and improved soil quality (Lanini et
al. 1989). However, recent studies have shown that when
living mulches are undersown with a vegetable crop
they can also help reduce injury imposed by insect pests
(Costello and Altieri 1995, Hooks et al. 1998, Hooks and
Johnson 2001). Undersowing is the intercropping of an
economically important crop with an undersown plant
species that has no direct market value but is used to di
versify the agroecosystem or influence the main crop.
The impact on insect pest densities of undersowing
vegetable crops with living mulches has been examined
mainly in Brassica crops (Asman et al. 2001; Hooks
and Johnson 2001, 2002). In most of these studies,
fewer insect pests were found on interplanted vegetable
crops (Hooks and Johnson 2003 and references therein).
However, Brassica crops are typically slow growing
and do not compete well with background vegetation.
Therefore, most of these studies reported significant
yield reductions, possibly caused by interspecies com
petition.
Several strategies may be implemented to limit com
petition among Brassica crops and their companion
plants. These strategies may include
• proper fertilization and irrigation of the main crop
• use of vigorous or rapidly growing crop cultivars
• optimal spacing between the main crop and compan
ion plants
• use of less competitive background plants (e.g., low
canopy height)
• timely planting of the main crop and companion plant
• planting the intercrop at a lower seed rate or in nar
rower strip
• suppression of the intercrop (e.g., mowing, reduced
fertilization and irrigation) at critical times
• use of a self-suppressing companion plant (i.e., one
that dies during the critical period of crop growth)
• use of non-crop borders surrounding the field crop.
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in coopera
tion with the U.S. Department of Agriculture. Andrew G. Hashimoto, Director/Dean, Cooperative Extension Service/CTAHR, University of Hawai‘i at Mänoa, Honolulu, Hawai‘i 96822.
An equal opportunity/affirmative action institution providing programs and services to the people of Hawai‘i without regard to race, sex, age, religion, color, national origin, ancestry, dis
ability, marital status, arrest and court record, sexual orientation, or status as a covered veteran. CTAHR publications can be found on the Web site <http://www.ctahr.hawaii.edu/freepubs>.
UH–CTAHR
Each approach should be viewed with caution and
weighed against its potential to negate any positive
benefits of pest suppression.
This publication describes a field experiment that
was part of a continuing effort to refine management of
lepidopteran pests in broccoli by mixed cropping and
undersowing it with other plant species. The primary pur
pose of this study was to determine if specific undersown
living mulches would have the ability to suppress lepi
dopteran pest densities without reducing crop yields to
economically unacceptable levels. We focused on cover
crops in the genus Trifolium partially because of their
low-growing structural similarities to yellow sweetclover.
Additionally, because spiders were recurrently observed
preying on various lepidopteran caterpillars (Hooks and
Johnson 2002), their densities were compared among
undersown and bare-ground broccoli.
Materials and methods
Experiment layout
The field experiment was conducted during the summer
2000 at the University of Hawai‘i at Mänoa’s Poamoho
Research Station on O‘ahu. The four cropping systems
examined were broccoli plants undersown with: (1) yel
low sweetclover (Melilotus officinalis L.) seeded at 72 g
per row, (2) strawberry clover cv. O’Conners (Trifolium
fragiferum L. seeded at 58 g per row, (3) white clover
cv. New Zealand (Trifolium repens L.) seeded at 54 g
per row, and (4) broccoli monoculture used as a check
treatment. Experimental plots were 11 m x 11 m with
each treatment replicated four times and arranged in a
randomized complete block design.
The living mulches were sown on 10 July 2000 in all of
the undersown plots. Undersown plots contained 10 rows
of broccoli with 11 rows of the living mulch species and
monoculture plots contained 10 rows of broccoli. Broccoli
seedlings, cv. Liberty (Petoseed Co., Saticoy, California),
were grown for 5 weeks in the greenhouse before being
manually transplanted on September 18 and 19.
Clover growth
The spread of the clover canopies was monitored weekly
beginning 16 days after broccoli planting (DAP) until
broccoli harvest. Five areas between two adjacent clover
rows in each plot were randomly selected, excluding
border rows, and the distance of exposed soil surface (not
covered by clover canopy) between rows was measured.
2
IP-27 — May 2007
Arthropod census
Caterpillar pest stages and predators on broccoli plants
were sampled at 7-day intervals until the harvest period.
Sampling was stratified according to plant structure (5
upper, 10 middle, and 10 lower positioned leaves). Egg,
caterpillar, and pupa stages of the diamondback moth,
Plutella xylostella L., imported cabbageworm, Pieris
rapae L, and cabbage looper, Trichoplusia ni Hübner,
were counted separately.
Broccoli yield
The diameter and weight of 16 broccoli crowns, chosen
at random from the interior rows of each plot, were mea
sured at harvest time. The heads were then completely
dissected and examined for insects, insect parts, and as
sociated contaminants (e.g., frass, webbing, cocoon).
The effects of habitat type on the experimental param
eters were analyzed using analysis of variance (Proc
GLM, SAS Institute, Cary, NC 1990) and predetermined
orthogonal comparisons to separate mean differences.
Within the model, the following predetermined contrasts
were conducted: broccoli-clovers vs. monoculture; true
clovers (white and strawberry) vs. yellow sweetclover;
and strawberry clover vs. white clover. Because eggs of
moths and butterflies were acutely low, all species were
pooled together by stage (e.g., egg, larvae) prior to final
analysis. The criteria for significance was P < 0.05.
Results
Clover growth
The yellow sweetclover canopy expanded over the soil
surface faster than the other undersown living mulches.
Subsequently, the amount of exposed soil surface area
between broccoli rows was significantly less in broc
coli–yellow sweetclover contrasted with broccoli under
sown with strawberry clover and white clover on each
sampling date. Strawberry clover canopy development
occurred at the slowest rate and was significantly less
than that of white clover on most dates.
Arthropod census
Effect of clovers on leidopteran pest densities. Lepi
dopteran populations were low during the experiment.
Approximately 85, 10, and 5 percent of the lepidopteran
fauna observed were P. rapae, T. ni, and P. xylostella,
UH–CTAHR
IP-27 — May 2007
Figure 1. Mean population densities of lepidopteran (a) eggs and (b) larvae in bare-ground broccoli (BG); broccoli–
strawberry clover intercrop (SC); broccoli–white clover intercrop (WC); and broccoli–yellow sweetclover intercrop (YSC).
Figure 1.
*** indicates intercrops significantly less than bare ground; s indicates (SC) significantly less than (WC); w indicates (WC)
significantly less than (SC); y indicates (YSC) significantly less than (SC + WC); and ns means no significant differences
exist (P > 0.05).
0.7
BG
SC
WC
YSC
0.6
Number
of lepidopteran
per leaf
Number
of lepidopteran
per leaf
0.5
0.4
0.3
***
s
ns
ns
0.2
A
eggs
***
ns
ns
0.1
0
0.8
larvae
ns
B
0.7
0.6
0.5
0.4
w
0.3
y
*** y
***
30
37
44
***
0.2
0.1
***
0
16
23
51
58
Days
planting
Daysafter
After
Planting
respectively. From early to mid-season, no significant
differences were detected in the abundance of eggs
among broccoli habitats (Figure 1a). However, during
the late broccoli growth cycle, more eggs were found
in monoculture than in undersown broccoli. Similarly,
more caterpillars were recorded in monoculture com
pared with undersown broccoli from mid- to late season
(Figure 1b).
Impact of clovers on spider abundance. At the ex
perimental site, four spider species frequently inhabit
Brassica plants. Nesticodes (= Theridion) rufipes Lucas
(Theridiidae), Neoscona oaxacensis Keyserling (Ara
neidae), Oxyopes sp. (Oxyopidae), and Cheiracanthium
mordax L. Koch (Clubionidae) composed approximately
77, 13, 7, and 3 percent of the spider fauna, respectively.
Significantly fewer spiders were encountered on broccoli
plants with clovers compared with bare-ground broccoli
during the initial three sampling dates. However, during
the later part of the broccoli growth cycle, this trend re
versed, and more spiders were found on broccoli plants
in clover than in bare-ground plots (Figure 2).
Additionally, fewer spiders were found on broccoli
grown with the true clovers (e.g., strawberry clover,
white clover) compared with plants grown with yellow
sweetclover during mid-season. Fewer spiders were also
observed on broccoli plants undersown in strawberry
3
UH–CTAHR
IP-27 — May 2007
Numberof
of spiders
spiders per
leaf
Number
per
leaf
Figure 2. Mean population
Figure 2. densities of spiders in bare-ground broccoli (BG); broccoli–strawberry clover intercrop (SC);
broccoli–white clover intercrop (WC); broccoli–yellow sweetclover intercrop (YSC). * indicates (BG) significantly less
than intercrops; *** indicates intercrops significantly less than (BG); c indicates (SC + WC) significantly less than (YSC);
s indicates (SC) significantly less than (WC); and ns means no significant differences exist (P ≥ 0.05).
1
BG
SC
WC
YSC
0.8
0.6
*cs
***
c
***
0.4
*
s
51
58
***
0.2
0
16
23
30
37
44
DaysAfter
after Planting
planting
Days
clover compared with white clover from mid- to late
season.
Crown contamination
At harvest, broccoli crowns were infested with various
stages of Pieris rapae, Trichoplusia ni, and Plutella
xylostela (Table 1). T. ni caterpillars and pupae were the
most abundant lepidopteran contaminants encountered in
broccoli heads. However, significantly more individuals
of all three species were found in crowns harvested from
bare-ground broccoli compared to undersown broccoli.
Crop yield
The largest crowns by diameter and mass were harvested
from broccoli undersown with strawberry clover or
white clover (Table 2). These crowns weighed signifi
cantly more than those harvested from broccoli–yellow
sweetclover plots. Yellow sweetclover plots contained
the smallest crowns, by weight.
Discussion
The purpose of this study was to determine if undersown
clovers could reduce lepidopteran pest densities without
4
reducing crop yields to unacceptable levels. We found
that the number of insect contaminants per broccoli
crown was significantly reduced on plants undersown
with strawberry clover and white clover compared to
bare-ground broccoli without causing any yield reduc
tions. Additionally, spider densities found on broccoli
plants seemed to be influenced by the amount of clover
canopy. As the clover canopies expanded and approached
the broccoli plants, more spiders were found on the
broccoli foliage.
Impact on spider abundance
During the early part of the season, the living mulches
may have negatively influenced biological control activ
ity on broccoli plants by serving as a “sink” for spiders.
Spiders may have preferred the micro-environment
2
and prey selection within the clovers and thus did not
colonize neighboring broccoli plants. Similar observa
tions were made in previous field experiments in which
fewer spiders were found on broccoli plants intercropped
with peppers or yellow sweetclover, but as the season
progressed these differences diminished.
UH–CTAHR
IP-27 — May 2007
Table 1. Mean number of lepidopterans per broccoli head in four broccoli habitats during summer 2000 (mean ± SE).
Habitat
Pieris rapae
Trichoplusia ni
Plutella xylostella
Total
Bare ground
0.1 + 0.0
0.8 + 0.10
0.13 + 0.0
0.87 + 0.0
Broccoli-SC
0.08 + 0.04
0.10 + 0.04
0.00 + 0.00
0.18 + 0.06
Broccoli-WC
0.0 + 0.03
0.00 + 0.00
0.02 + 0.02
0.07 + 0.03
Broccoli-YSC
0.0 + 0.03
0.12 + 0.0
0.03 + 0.02
0.20 + 0.06
P-value
Contrast1
BG vs. LMs
TCs vs. YSC
SC vs. WC
0.03
0.07
0.0
0.01
0.4
0.62
0.006
0.4
0.67
< 0.0001
0.39
0.20
SC (strawberry clover), WC (white clover), YSC (yellow sweetclover), BG (bare ground, broccoli monoculture); LMs (living mulches) includes
broccoli-SC, broccoli-WC, and broccoli-YSC; TCs (true clovers) includes broccoli-SC and broccoli-WC.
1
Conclusion
Using undersown living mulches seems to be promising
in reducing lepidopteran pest densities and increasing
the activity of spiders in broccoli plantings. In this study,
white clover appeared to be more suited for the broccoli
system. White clover expands over the soil surface faster
than strawberry clover and may therefore be a better
weed suppressor. For those farmers looking to create
more sustainable cropping practices or practicing organic
farming, undersowing may be a valuable addition to their
crop production practices.
This field trial showed that it is possible to lower insect
pest density while maintaining crop quality and yield.
However, insect pest management is only one potential
benefit of using leguminous living mulches. Other po
tential benefits not examined during this study include
nematode and weed suppression and enhancement of
soil nitrogen. However, before an undersown compan
ion plant is chosen for insect suppression purposes, its
impact on other important organisms associated with the
crop should be considered.
Acknowledgements
The authors wish to thank the crew at the Poamoho
Research Station for assisting in the field and Dr. Raju
Pandey for his notable contributions to this study. This
project was funded by the USDA/CSREES Special
Grant for Tropical and Subtropical Agriculture Research
(T-STAR).
Table 2. Mean head size per broccoli plant in four habitats
during summer 2000.
Broccoli parameters1
(mean ± SE)
Habitat
Diameter (cm)
Weight (kg)
Bare ground
13.0 + 0.3
0.3 + 0.01
Broccoli-strawberry clover
14.7 + 0.3
0.40 + 0.02
Broccoli-white clover
14.4 + 0.24
0.39 + 0.01
Broccoli-yellow sweetclover 13.7 + 0.28
0.33 + 0.01
Effect2
Planned contrast
BG vs. LMs
TCs vs. YSC
SC vs. WC
P -Values
0.0002
0.03
0.71
0.11
0.000
0.1
1
Multiply cm by 0.394 to obtain inches, and multiply kg by 2.2 to obtain pounds.
2
SC (strawberry clover), WC (white clover), YSC (yellow sweetclover), BG (bare ground, broccoli monoculture); LMs (living mulches) include broccoli-SC, broccoli-WC, and broccoli-YSC; TCs (true clovers) include broccoli-SC and broccoli and broccoli-WC.
References and further reading
Asman, K., B. Ekbom, and B. Rämert. 2001. Effect of
intercropping on oviposition and emigration behavior
of the leek moth (Lepidoptera, Acrolepiidae) and the
diamondback moth (Lepidoptera, Plutellidae). Envi
ronmental Entomology 30: 288–294.
UH–CTAHR
Costello, M.J., and M.A. Altieri. 1995. Abundance,
growth rate and parasitism of Brevicoryne brassicae
and Myzus persicae (Homoptera, Aphididae) on broc
coli grown in living mulches. Agriculture, Ecosystems
and Environment 52: 187–196.
Hooks, C.R.R., H.R. Valenzuela, and J. Defrank. 1998.
Incidence of pests and arthropod natural enemies in
zucchini grown with living mulches. Agriculture,
Ecosystems and Environment 69: 217–231.
Hooks, C.R.R., and M.W. Johnson. 2001. Broccoli
growth parameters and level of head infestations
in simple and mixed plantings: Impact of increased
6
IP-27 — May 2007
flora diversification. Annals of Applied Biology 138:
269–280.
Hooks, C.R.R., and M.W. Johnson. 2002. Lepidopteran
pest populations and crop yields in row intercropped
broccoli. Agriculture and Forest Entomology 4:
117–126.
Lanini, W.T., D.R. Pittenger, W.L. Graves, F. Munoz, and
H.S. Agamalian. 1989. Subclovers as living mulches
for managing weeds in vegetables. California Agri
culture. Nov.–Dec., p. 25–27.
SAS Institute. 1990. SAS User’s Guide: Statistics. Cary,
NC.
Insect Pests
Jan. 2008
IP-28
Guide to Insect and Mite Pests
of Tea (Camellia sinensis) in Hawai‘i
Randall T. Hamasaki1, Robin Shimabuku2, and Stuart T. Nakamoto3
1,2
Department of Plant and Environmental Protection Sciences, 1Kamuela Extension Office, 2Kahului Extension Office;
3
Department of Human Nutrition, Food and Animal Sciences
T
his guide provides photographs and general informa
tion about insect and mite pests associated with tea
in Hawai‘i. Details on pest identification, crop damage,
crop hosts, pest life cycle, and pest distribution are given.
Accurate identification of the pest is essential for mak
ing sound pest management decisions. Early detection is
often critical to eventual success in managing pests and
reducing economic losses.
The pests were selected based on surveys of tea plants
growing at the UH-CTAHR Mealani Research Station at
Type of damage
2800 feet elevation in Waimea on Hawai‘i. In addition,
pests were collected from cooperating growers in other
locations on Hawai‘i. Pest samples were identified by
the UH-CTAHR Agricultural Diagnostic Service Center
(ADSC).
If you suspect pest problems but cannot determine the
cause, we suggest that you submit plant samples to the
ADSC for identification. The samples may be taken to
the nearest UH-CTAHR Cooperative Extension Service
office.
Page
Pests with damage caused by chewing
Chinese rose beetle, Adoretus sinicus (Burmeister) .......................................................................................2
Mexican leafroller, Amorbia emigratella (Busck)..........................................................................................3
Pests that feed on plant sap
Red and black flat mite, Brevipalpus phoenicis (Geijskes) ............................................................................4
A spider mite (unidentified) ...........................................................................................................................5
Broad mite or yellow tea mite, Polyphagotarsonemus latus (Banks) ............................................................6
Mining scale, Howardia biclavis (Comstock)................................................................................................7
Avocado scale, Fiorinia fioriniae (Targioni-Tozzetti) ....................................................................................8
Florida red scale, Chrysomphalus aonidum (L.) ...........................................................................................9
Brown soft scale, Coccus hersperidum (Linnaeus)......................................................................................10
Melon aphid or cotton aphid, Aphis gossypii (Glover) ................................................................................. 11
Spiraling whitefly, Aleurodicus dispersus (Russell).....................................................................................12
Twospotted leafhopper, Sophonia rufofascia (Kuoh & Kuoh).....................................................................13
Transparentwinged plant bug, Hyalopeplus pellucidus (Stål)...................................................................... 14
Greenhouse thrips, Heliothrips haemorrhoidalis (Bouche).........................................................................15
Acknowledgments
Project support was received from Extension Integrated Pest Management (IPM), Dr. Arnold H. Hara, IPM Coordinator for Hawai‘i. The
authors thank Milton Yamasaki and the staff of the Mealani Research Station and Brian Bushe and Dick M. Tsuda of the CTAHR Agricul
tural Diagnostic Service Center.
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in cooperation with the U.S. Department of Agriculture. Andrew G. Hashimoto, Director/Dean, Cooperative Extension Service/CTAHR, University of Hawai‘i at Mänoa, Honolulu, Hawai‘i 96822.
An equal opportunity/affirmative action institution providing programs and services to the people of Hawai‘i without regard to race, sex, age, religion, color, national origin, ancestry, disability, marital status, arrest and court record, sexual orientation, or status as a covered veteran. CTAHR publications can be found on the Web site <http://www.ctahr.hawaii.edu/freepubs>.
UH–CTAHR
Insect and Mite Pests of Tea
IP-28 — Jan. 2008
Chinese rose beetle
Adoretus sinicus Burmeister, Coleoptera: Scarabaeidae
Damage
Life cycle
Adult Chinese rose beetles are nocturnal and chew plant
leaves. Recently transplanted and young plants appear
to be most susceptible, although established plants may
also be attacked. Serious defoliation can occur when pest
numbers are high, and this may kill young plants. Only
the adult stage of the insect will damage crops.
The larval stages are commonly found in the soil of lawns
and gardens where organic matter is present. The grubs
are thought to feed on organic matter and do not attack
plants. Eggs are laid in soil about 11 ⁄2 inches deep. They
hatch in about 7–16 days. There are three larval stages.
The grubs are whitish with a conspicuous brown head and
short legs. When still, they tend to be C-shaped. The lar
val stage lasts for 3–4 weeks. The pupa is yellowish-white
when initially formed and then turn brown. Pupation is
completed in 1–2 weeks. Development from egg to adult
takes 7–16 weeks, depending on temperature. The life
cycle from egg to adult is completed in 6–7 weeks.
Identification
Holes in leaves and chewing of all but the leaf veins are
signs of feeding damage. These beetles are active at night
and will not be present during they day. Look for beetles
beginning about 30 minutes after sunset. The beetles are
sturdy, pale reddish brown, and about 1 ⁄2 inch long. The
body is densely covered with minute hairs, which may
give it a grayish appearance.
Hosts
The plant host range for this species comprises over 250
plants from a wide variety of ornamental and cultivated
crops, including asparagus, beans, broccoli, cabbage,
cacao, Chinese broccoli, Chinese cabbage, chiso, corn,
cotton, cucumber, eggplant, flowering white cabbage,
ginger, grape, green bean, okra, rose, soybean, straw
berry, sweetpotato, and tea.
Chinese rose beetle
2
Distribution
Originally from Japan and Taiwan, this beetle is widely
distributed throughout Southeast Asia and many Pacific
islands. Introduced to Hawai‘i before 1896, it is now a
common pest on all major islands in the state.
Reference
Mau, R.F.L., and J.L. Martin. Adoretus sinicus (Burmeis
ter). Crop Knowledge Master. www.extento.hawaii.
edu/ Kbase/crop/type/adoretus.htm
Feeding damage on young tea plant
Grub in soil
UH–CTAHR
IP-28 — Jan. 2008
Mexican leafroller
Amorbia emigratella Busck, Lepidoptera: Tortricidae
Damage
This caterpillar rolls the young leaves at the shoot tips
and lives and feeds within. Leaves from damaged shoots
have holes and may be distorted. In tea, insect parts
may contaminate the harvested product. In addition to
damage in the field, this insect can be a pest of cuttings
in the nursery.
Identification
macadamia, orchids, papaya, passion fruit, potato, rose,
sweetpotato, tea, and tomato. It also attacks many other
shrubs, fruit trees, and indigenous Hawaiian plants in
the mountains.
Life cycle
Eggs are laid in clusters of 65–120 on the upper surfaces
of leaves. There are three or four molts in the larval
stage, which is completed in 28–35 days. Pupation oc
curs within the folded leaf. The adult emerges in about
10 days. The life cycle from egg to adult takes from
48–55 days.
Examine the shoot tips for rolled leaves and look for
the caterpillar inside. Newly hatched caterpillars are
1
⁄8 inch long, growing to 1 inch long when fully grown.
They have a brownish-yellow head, a light-green body,
and a black stripe on the sides behind the eyes. The adult
moths are yellowish-brown with a small pointed head.
The wingspan of female moths is 1–11 ⁄6 inches. Males
are slightly smaller and paler.
This caterpillar has been in Hawai‘i since 1900 and has
been reported from all major islands except Läna‘i. It
also occurs in Mexico and Costa Rica.
Hosts
Reference
This pest has a wide host range. It is commonly found
on ornamental plants and some fruit trees, but vegetables
are not common hosts. Hosts include avocado, broc
coli, cacao, citrus, cotton, eggplant, green beans, guava,
Mau, R.F.L., and J.L. Martin Kessing. 1992. Amorbia
emigratella (Busck). Crop Knowledge Master. www.
extento.hawaii.edu/kbase/crop/type/amorbia.htm
Distribution
Mexican leafroller
caterpillar
Damaged tea shoot tip
Adult moth
3
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Red and black flat mite
Brevipalpus phoenicis (Geijskes), Acari: Tenuipalpidae
Damage
Red and black flat mites feed on plant sap and cause
bronzing and/or browning of the leaves. These mites
favor the upper leaf surface of mature leaves, and the
damage progresses from the lower leaves to the younger
leaves. Young plants that are not yet fully established
appear to be highly susceptible.
feet, and has not been recorded in areas above 2500
feet elevation. It is usually not considered to be a pest of
economic importance above 1000 feet.
Reference
Martin Kessing, J.L., and R.F.L. Mau. 1992. Brevipalpus
phoenicis (Geijskes). Crop Knowledge Master: www.
extento.hawaii.edu/Kbase/crop/Type/b_phoeni.htm
Identification
Clusters of bright reddish orange eggs are more easily
seen with the naked eye than any other life stage. Note
that other mites found on tea also have reddish eggs.
These mites are microscopic—the adult female mite
is about a hundredth of an inch long. Populations are
primarily composed of females, with males less than
1 percent of the population. A feature distinguishing
these mites from other mites is that the body is flattened.
Coloration ranges from light to dark green or reddish
orange. There are four legs extending forward and four
legs extending behind. Depending on temperature, adult
females may have a black mark in the shape of an H. The
adult male is flat, reddish, more wedge-shaped than the
female, and lacks black markings.
Hosts
The red and black flat mite has been recorded on over
65 hosts. In Hawai‘i, the red and black flat mite has been
reported on anthurium, banana, hemigraphis, lemon,
macadamia, orchid, papaya, and passion fruit. In other
parts of the world it is common on tea and citrus.
Red and black flat mite damage
Life cycle
Reproduction primarily occurs without mating. The
life stages are egg, larva (six-legged), protonymph,
deutonymph, and adult. As observed under laboratory
conditions, egg-to-adult timespan has been observed to
be as short as 10.6 days at 86°F and as long as 27.3 days
at 68°F.
Distribution
This mite was first found in Hawai‘i on O‘ahu in 1955
and has subsequently been reported on Kaua‘i, the Big
Island, and Maui. The mite is abundant in areas between
sea level and 1000 feet, scarce between 1000 and 2500
4
Red and black flat mite viewed under microscope
UH–CTAHR
IP-28 — Jan. 2008
A spider mite
(identification pending)
Acari: Tetranychidae
Damage
Leaves of severely damaged plants turn reddish and
drop prematurely. Plants may be totally defoliated when
populations are very high. The damage progresses from
older leaves upward to the younger growth. Although
mites and their eggs are present on both leaf surfaces,
they appear to prefer the upper leaf surface. Plants in
the establishment phase appear to be the most prone to
severe damage from this mite.
Spider mites and their eggs
Identification
Mites and their eggs are reddish.
Hosts
Tea; other hosts unknown.
Life cycle
As yet unknown.
Distribution
As yet unknown.
A tea plant severely damaged by spider mites
Plants recovering from defoliation
5
UH–CTAHR
Broad mite, yellow tea mite
Polyphagotarsonemus latus (Banks),
Acari: Tarsonemidae
Damage
Broad mites feed on plant sap and cause scarring and
distortion of the leaves and stems. The scarred tissue may
appear to be a greasy darkened discoloration that may
later turn to a brown, corky surface on the undersides of
leaves. Broad mites appear to favor young growth. Plants
in the greenhouse or nursery are highly susceptible.
Identification
Although broad mite eggs are microscopic (0.08 mm
long), they are distinct and helpful in identifying broad
mite infestations. The clear eggs are oval and have five
to six rows of whitish bumps. A good hand lens (at least
10x) is needed to see the eggs.
Hosts
The broad mite attacks many plants including bit
termelon, Chinese waxgourd, chiso, chrysanthemum,
cucumber, edible gourds, eggplant, green beans, guava,
hyotan, macadamia, mango, papaya, passion fruit, pep
per, pikake, plumeria, poha, pumpkin, Spanish needle,
tomato, watercress, winged bean, and yardlong bean. In
temperate and subtropical areas, the broad mite is a pest
of greenhouse plants.
IP-28 — Jan. 2008
Life cycle
The life cycle, from egg to adult, is completed in about
4–6 days. The number of eggs laid per female and the
population growth are affected by temperature and rela
tive humidity.
Distribution
This mite has a worldwide distribution. It is known to
occur in Australia, Asia, Africa, North America, South
America, and the Pacific. Countries included in this
mite’s distribution include American and Western Sa
moa, Bermuda, Brazil, China, Cook Islands, Guyana,
Fiji, India, Japan, Kiribati, Malaysia, Marianas, New
Caledonia, Pakistan, Papua New Guinea, Philippines,
Sri Lanka, Taiwan, Tonga, Vanuatu, and Wallis. It is
present on all the major islands of Hawai‘i.
Reference
Martin Kessing, J.L., and R.F.L. Mau, Polyphagotarsonemus latus (Banks). 1993. Crop Knowledge Master:
www.extento.hawaii.edu/kbase/crop/Type/p_latus.htm
Broad mites and egg
Damaged shoot
6
Scarring on leaf undersides
UH–CTAHR
IP-28 — Jan. 2008
Mining scale
Howardia biclavis (Comstock),
Homoptera: Diaspididae
Damage
References
In general, feeding by these insects on the juices of its
host plant causes loss of vigor, deformation of infested
plant parts, and even death of the plant. In tea plants, this
scale has been mostly found on the bark of the trunk.
Tenbrink, V.L., and A.H. Hara. Howardia biclavis
(Comstock), Crop Knowledge Master. www. extento.
hawaii.edu/Kbase/Crop/Type/h_biclav.htm
Watson, G.W. 2005. Arthropods of economic importance
—Diaspididae of the world. http://ip30.eti.uva.nl/bis/
diaspididae.php?selected=beschrijving&menuentry=
soorten&id=102
Identification
On tea plants, these scales appear to favor living on the
bark of the trunk and stems. The scales are round and
slightly dome-shaped and may measure up to 1 ⁄8 inch
in diameter. The color is variable from white to gray or
yellow. A reddish spot is located at or near to the edge
of the margin of the armor. This species is called the
mining scale because it may burrow beneath the host
plant’s epidermis and be partially concealed by it. They
are a transparent light or yellowish brown.
Hosts
In Hawai‘i it has been recorded on acacia, allamanda,
bougainvillea, cassia, ficus, ebony, gardenia, hibiscus,
ixora, jasmine, lantana, lychee, mango, papaya, plumeria,
poinsettia, pulasan, sapodilla, sapote, Sterculia foetida,
and tea. Among its many other hosts are albizia, kukui,
annona, camellia, citrus, coffee, tomato, and macada
mia.
Life cycle
Males have not been observed, and parthenogenesis (fe
males producing females) is suspected. The first nymphal
stage is commonly called the crawler stage, and it is at
this early stage that the insect is mobile on the plant and
can be transported to other plants by people, animals,
birds, ants, and wind currents. The life cycle is probably
about 30 days, based on the generalized life history of
other tropical armored scale species.
Distribution
The mining scale was first reported from Kona on
Hawai‘i in 1895. It has since been recorded from Ni‘ihau,
O‘ahu, and Maui. Worldwide, the mining scale is found
in the tropics and in glasshouses in temperate areas.
Mining scales on bark of tea plant
7
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IP-28 — Jan. 2008
Avocado scale
Fiorinia fioriniae (Targioni-Tozzetti),
Damage
This scale often causes yellow spotting on the leaves
where it feeds on plant sap, due to its toxic saliva. This in
sect is a pest of tea both in the nursery and in the field.
Identification
Look for the scales on the leaves, especially along the
veins. The scales are small: 1–1.5 mm long. They are
transparent and light or yellowish brown.
Hosts
In Hawai‘i this scale was first reported on tea on Maui in
1997. Elsewhere, it has been recorded on tea, avocado,
anthurium, Araucaria, Buchanania, Callistemon lanceolatus, Cinnamomum, Citrus spp., coconut, Cupressus, Cycas, Decaspermum, Dictyosperma, Eucalyptus,
Eugenia, Ficus spp., Hedera, Howea, Lauris nobilis,
Livistona, mango, Myristica, olive, Phoenix, Pinus,
Podocarpus, Salix, Santalum, Sida, Taxus, Ulmus, and
others.
Reference
Watson, G.W. 2005. Arthropods of economic impor
tance—Diaspididae of the world. http://ip30.eti.uva.
nl/bis/diaspididae.php?selected=beschrijving&menu
entry=soorten&id=102
Yellowing caused by feeding damage
Avocado scales on tea leaf
8
UH–CTAHR
IP-28 — Jan. 2008
Florida red scale
Chrysomphalus aonidum (Linnaeus),
Damage
Life cycle
The Florida red scale mainly infests leaves, where it
feeds on plant sap, but it may spread to other plant parts
when its population is very high. Severely infested leaves
may drop prematurely. Dry weather conditions favor
infestation.
Reproduction is sexual. Each adult female lays about
50–150 eggs. The eggs hatch under the female scale,
and these crawlers seek a suitable feeding site to settle.
Development from egg to adult takes 7–16 weeks, de
pending on temperature.
Identification
Distribution
From a distance, the scales appear as dark circular spots
on leaves, especially on the lower leaf surfaces. Closer
examination with a hand lens will reveal more detail.
Adult female scales are conical and up to 2 mm in diam
eter. The area near the tip of the cone may appear pale.
Immature male scales are smaller and paler than female
scales. They are elongate-oval and half the size of adult
females. Adult male scales are winged insects that look
very different from adult female scales.
References
Hosts
Hosts recorded in Hawai‘i include citrus, coconut, anthur
ium, bougainvillea, dendrobium, dracaena, eucalyptus,
ficus, hibiscus, palm, plumeria, podocarpus, bird of
paradise (Strelitzia), ginger (Zingiber officinale), Citrus
spp. (lime, lemon, pummelo, grapefruit), asparagus, tea,
apple, mango, banana and plantain, palms, and pines.
Florida red scales on tea leaf
In Hawai‘i, the Florida red scale was first reported from
Oahu in 1907. It has since been recorded from the Big
Island, Läna‘i, and Kaua‘i. The Florida red scale is very
widely distributed in the tropics and subtropics. It is
present in Europe, Asia, Africa, South America, parts
of North America such as Florida, Maryland, Texas, and
Virginia, and on many Pacific islands.
Heu, R.A. 2005. Agricultural pests, related organisms
and purposely introduced natural enemies in Hawaii.
Biological Control Section, Hawai‘i Department of
Agriculture.
Watson, G.W. 2005. Arthropods of economic impor
tance—Diaspididae of the world. http://ip30.eti.uva.
nl/bis/diaspididae.php?selected=beschrijving&menu
entry=soorten&id=102
Close up of Florida red scales
(note the yellow crawlers)
9
UH–CTAHR
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Brown soft scale
Coccus hesperidum Linnaeus, Homoptera: Coccidae
Damage
Soft scales feed on plant sap and excrete honeydew, a
sugar-rich substance that is fed on by ants and is a sub
strate for the sooty mold fungus.
Identification
Adult female scales are pale yellowish brown to greenish.
The color may darken with age. They are 1 ⁄8–1 ⁄6 inch long.
Male scales have not been recorded for this species.
Hosts
The brown soft scale attacks a variety of field, orna
mental, and greenhouse crops. Host plants reported in
Hawai‘i include citrus, iliahi, loquat, orchids, papaya,
and tea.
Distribution
The brown soft scale was first recorded in Hawai‘i in 1896
and is found on all the main islands. It has been reported
in Algeria, Australia, Austria, British Guiana, Canada,
Chile, Cuba, Dutch East Indies, Ecuador, England, Eu
rope, Haiti, Japan, Mauritius, Mexico, Morocco, New
Zealand, Seychelles, South Africa, and West Indies.
Reference
Tenbrink, V.L., and A.H. Hara. 1994. Howardia biclavis
(Comstock). Crop Knowledge Master. www.extento.
hawaii.edu/Kbase/Crop/Type/h_biclav.htm
Life cycle
Brown soft scales reproduce primarily by parthenogen
esis (females producing females without mating) and
live birth. It makes up for its relatively slow growth by
producing large numbers of offspring (80–250 eggs
per female). The first stage is tiny crawlers, which are
the dispersive stage. The nymphs undergo three molts
before they become adults. The adult female scales are
immobile.
Brown soft scales on a tea leaf
10
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Melon aphid (cotton aphid)
Aphis gossypii Glover, Homoptera: Aphididae
Damage
Life cycle
Melon aphids have piercing-sucking mouthparts that en
able them to feed on plant sap. These aphids excrete hon
eydew, which is a sweet, sticky substance that can become
deposited on infested plants. Honeydew is attractive to
ants and is a substrate for sooty mold fungus. Sooty mold
blackens the leaf surface and my decrease photosynthesis.
Infested leaves may become cupped and distorted. Melon
aphids commonly infest the tea shoot, and their body parts
may end up in the finished product.
In Hawai‘i, melon aphids are females that reproduce
without mating. They do not lay eggs, but instead produce
live nymphs. There are four nymphal stages separated by
molts. The nymphs become adults in 4–12 days, depend
ing on temperature. Adult aphids are generally wingless,
but overcrowding or decline of the host plant can trigger
production of winged forms. Adult aphids may live for
3–4 weeks and produce about 85 offspring each.
Identification
Melon aphids occur in tropical and temperate regions
throughout the world, except for the northernmost re
gions. In Hawai‘i it was first reported on Oahu in 1909
and is now present on all islands.
Melon aphids are soft-bodied insects that vary in color
from black to dark brown to brownish green to yellowish
green. They are usually 1 ⁄16 inch or less in size. Adults
may be winged or wingless. On tea, they often live in
groups on the underside of leaves at the shoot tips.
Hosts
The melon aphid attacks a wide variety of plants includ
ing many cucurbit vegetables, eggplant, guava, hibiscus,
orchids, peppers, taro, and weeds such as lamb’s quarters,
cheeseweed, and Spanish needle.
Distribution
Reference
Martin-Kessing, J.L., and R.F.L. Mau. 1991. Aphis gossyppii (Glover). Crop Knowledge Master. http://www.
extento.hawaii.edu/Kbase/crop/Type/aphis_g.htm
Melon aphids on tea shoot
11
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Spiraling whitefly
Aleurodicus dispersus Russell,
Homoptera: Aleyrodidae
Damage
It is common to find the waxy spirals made by whiteflies
on the undersides of tea leaves. However, because the
amount of whiteflies is usually kept at low levels, this
insect is likely to be only a minor pest of tea in Hawai‘i.
Whiteflies feed on plant sap and secrete honeydew and a
white, waxy material. Honeydew can serve as a substrate
for the sooty mold fungus.
Identification
Eggs are laid in waxy spirals that give this whitefly their
common name (see photo). When magnified, whitefly
adults somewhat resemble tiny moths. Adult spiraling
whiteflies are relatively large compared to other common
whiteflies and measure 2–3 mm in length. Larval and
pupal stages secrete a waxy material that may be in the
form of rod-like projections that appear fluffy. Labora
tory identification is often based on taxonomic characters
found on the pupal stage.
Distribution
The spiraling whitefly is native to the Caribbean region
and has spread to Africa, Australia, Bahamas, Barbados,
Brazil, Canary Islands, Costa Rica, Cuba, Dominica,
Ecuador, Haiti, India, Martinique, Panama, Peru, Phil
ippines, Republic of Maldives, Singapore, Sri Lanka,
Thailand, USA, Vietnam, and the West Indies. In the
Pacific it is present in American Samoa, Cook Islands,
Fiji, Hawai‘i, Kiribati, Majuro, Mariana Islands, Nauru,
Palau, Papua New Guinea, Pohnpei, Tokelau, Tonga,
and Western Samoa. This whitefly was first reported in
Hawai‘i in 1978 on O‘ahu and had spread to all the major
islands by 1981. It is most abundant in coastal areas and
elevations below 1000 feet.
Reference
Martin Kessing, J.L., and R.F.L. Mau. 1993. Aleurodicus
dispersus (Russell). Crop Knowledge Master. www.
extento.hawaii.edu/kbase/Crop/Type/a_disper.htm
Hosts
The spiraling whitefly has a wide host range and has
been recorded from over 100 plant species. It is common
to find this whitefly on various ornamental, fruit, and
shade tree crops in Hawai‘i. Some common host plants
include Annona sp., avocado, banana, bird-of-paradise,
breadfruit, citrus, coconut, eggplant, guava, kamani,
Indian banyan, macadamia, mango, palm, paperbark,
papaya, pepper, pikake, plumeria, poinsettia, rose, sea
grape, tī, and tropical almond.
Life cycle
The life stages are egg, three larval stages, pupal stage,
and adult. Eggs are elliptical, yellow to tan, and are laid
in groups of a few to several dozen in spiraling, waxy
lines on the leaf underside. Eggs hatch in 9–11 days. The
first larval stage is sometimes called the crawler stage
and is the only immature stage with functional legs that
enable mobility. The second and third larval stages are
sedentary, and waxy material is secreted. The third lar
val stage molts into the pupal stage. Pupae are white to
yellowish, nearly oval
12
Spiraling whitefly
adults, larvae, and
pupae
Whitefly eggs in waxy
“spiral”
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Twospotted leafhopper
Sophonia rufofascia (Kuoh & Kuoh),
Homoptera: Cicadellidae
Damage
References
The twospotted leafhopper uses piercing-sucking type
mouthparts to feed on plant sap. Although both nymphs
and adults are often associated with tea plants in Hawai‘i,
the importance of this insect to tea crops is yet unknown.
In some other plants, this insect is known to cause injury
by injecting saliva into the plant while feeding. Symp
toms of plant reaction to the saliva include leaf yellowing,
formation of brown or black patches on the leaves, leaf
distortion, and stunting of the plant.
Jones, V.P., M.T. Fukada, D.E. Ullman, J.S. Hu, and
W.B. Borth. Sophonia rufofascia The two spotted
leafhopper. Crop Knowledge Master. www.extento.
hawaii.edu/kbase/Crop/Type/s_rufofa.htm
Duan, J. and R. Messing. Biological control of the two
spotted leafhopper. www2.ctahr.hawaii.edu/t-star/
leafhopper.htm
Identification
Adult leafhoppers are about 3⁄16 inch long and are yel
lowish, with a dark stripe with red markings through the
middle of the body and two dark spots on the posterior
end. Nymphs are smaller, do not have wings, and cannot
fly. Nymphs are translucent yellow and have two dark
spots on the posterior end. The skin that is cast upon
molting has the dark spots also. These cast skins tend
to remain on the leaves for some time and can useful for
determining the presence of the pest.
Twospotted leafhopper adult
Hosts
This leafhopper attacks over 300 species of plants in
cluding many fruit, vegetable, and ornamental crops as
well as endemic plants. A few examples include avocado,
guava, chili peppers, sweetpotato, ti, octopus plant, uluhe
fern, and mamaki.
Life cycle
The eggs are laid in plant tissue and are very difficult to
detect. Eggs take about 4 weeks to hatch. There are four
nymphal stages which last a total of about 7–8 weeks.
Cast skin (note spots)
Distribution
This species was originally described in southern China.
It was first discovered in the state on O‘ahu in 1987 and
has since spread to all the major Hawaiian islands.
Nymphal stage
13
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IP-28 — Jan. 2008
Transparentwinged plant bug
Hyalopeplus pellucidus (Stål), Heteroptera: Miridae
Damage
Hosts
Transparentwinged plant bugs are frequently associated
with tea plants in Hawai‘i, but it is not known if this insect
is a pest of tea. It is a serious pest of guava in Hawai‘i,
where its feeding and egg-laying into flower buds causes
bud drop. On guava, this insect prefers to feed on the co
rolla region of the flower bud, where it results in a necrotic
blackening of the anthers within the bud. It is thought that
salivary enzymes are involved in the plant damage.
This insect has been collected from Acacia koa, avocado,
coffee, Coprosoma, Dodonaea, guava, Hibiscus sp., rose
flowered jathropha, Metrosideros, Pipturis, Psidium
cattleianum, Sida, Straussia, and Trema orientalis
(charcoal tree).
Identification
Adults are 1 ⁄3 –2 ⁄5 inch long. The transparentwinged
plant bug is perhaps the largest species from the family
Miridae in Hawai‘i. The adult has smoky colored wings
that are folded over the back when at rest. Nymphal
stages are pale, translucent green with purplish-red or
pinkish-red specks on the abdomen and heads shaped
similar to that of the adults, one-half wider than long,
and with the vertex being wider than the eyes together.
Black bristly hairs over an undercoat of golden yellow
hairs cover the head and antennae. The second antennal
segment is three times the length of the first and twice
as long as the third.
Adult transparentwinged plant bug
14
Life cycle
The life stages are egg, five nymphal stages, and the
adult stage. The eggs hatch in 6–8 days after being laid
(inserted into plant tissue). The duration of the nymphal
stages is about 14 days.
Distribution
The transparentwinged plant bug was first reported in
Hawai‘i in 1902 and occurs on all of the major islands
from sea level to the mountains. This insect might be
endemic to Hawai‘i.
Reference
Mau, R.F.L., and J.L. Martin. 1992. Hyalopeplus pellucidus (Stål). Crop Knowledge Master. www.extento.
hawaii.edu/Kbase/Crop/Type/h_pelluc.htm
UH–CTAHR
Greenhouse thrips
Heliothrips haemorrhoidalis (Bouche’),
Thysanoptera: Thripidae
Damage
This insect appears to be only a minor pest of tea in
Hawai‘i. Greenhouse thrips feed on plant sap, and the
damage causes a silvering of the leaf. These thrips appear
to prefer living and feeding on the undersides of the older
leaves of a tea plant. They cause a characteristic fecal
spotting, which appears as dark specks on the leaf. These
insects prefer to live in the shady areas of the tea tree
canopy and do not appear to damage the tea shoot.
Identification
IP-28 — Jan. 2008
is found in Europe in Germany, England, France, Italy,
Vienna, Finland, Palestine, and North Africa. This spe
cies is thought to be found throughout the world because
of its habit of living in greenhouses.
References
Denmark, H.A., and T.R. Fasulo. 2004. Greenhouse
thrips, Heliothrips haemorrhoidalis (Bouche).
University of Florida, IFAS Extension. EENY-075.
http://edis.ifas.ufl.edu/in232
Heu, R.A. 2005. Agricultural pests, related organisms
and purposely introduced natural enemies in Hawaii.
Biological Control Section, Hawai‘i Department of
Agriculture.
On the plant, check for silvering and fecal spotting, es
pecially on the undersides of older leaves. Mature larvae
and adult thrips are about 1 mm in length. Larvae are
yellowish, and adults are mostly black with light yellow
legs. Definitive identification can be done by an insect
diagnostic laboratory.
Hosts
In Hawai‘i the greenhouse thrips has been reported on
various ornamentals and conifers. Elsewhere, it has
been recorded on plants such as ardisia, Aspidium sp.,
avocado, azalea, Coleus sp., Crinum sp., croton, dahlia,
dogwood, ferns, guava, hibiscus, magnolia, mango, natal
plum, orange, phlox, and viburnum.
Life cycle
Thrips damage on leaf underside
The greenhouse thrips is parthenogenetic (females re
produce without mating). Eggs are laid singly in plant
tissue. There are two larval instars, which are the feeding
stages. The larval stage is then followed by a prepupal
and a pupal stage, during which the insect does not feed.
The pupal stage molts into the adult stage. The adult stage
has fully formed wings and is capable of flight.
Distribution
The greenhouse thrips was first reported in Hawai‘i
in 1910 from O‘ahu and has since been found on all
the major islands except Lāna‘i. It is thought to have
originated in tropical America. It is found in Brazil, the
West Indies, and Central America. It occurs in the U.S.
mainland outdoors in Florida and southern California.
It is found in greenhouses throughout the mainland. It
Adult greenhouse thrips
15
Insect Pests
June 2012
IP-29
Ant Damage to Banana Fruits by Abdominal Secretions
Scot Nelson and Glenn Taniguchi
S
HCO2H. This subfamily of ants
ome ants can directly damuses formic acid, which they
age plants and agricultural
eject or spray from an acidopore
commodities (Peng and Chrislocated at the end of the abdotian 2007); at least two species
men, to attack other animals and
of ants in Hawai‘i damage the
for self-defense. Formic acid
skin of banana fruits with their
is the simplest carboxylic acid
abdominal secretions. These
and one of the strongest acids
ants spray their secretions to
known, with a pH between 2
protect sap-feeding insects,
and 3. It can produce painful
from which they derive sweet,
injuries to human skin, causing
nutritious honeydew. The foragskin burns and eye irritation of
ing ants may also enter a self-defieldworkers. In Hawai‘i, the
fense mode and spray secretions
ant species that produce formic
if disturbed by banana cultivaacid are Anoplolepis gracilipes;
tion practices that jar the banana
Paratrechina longicornis; Plaplant, or if they are startled
giolepis allaudi; Nylanderia
when pesticide sprays impact
vaga; Nylanderia bourbonica;
the banana bunches. The marks
Lepisiota hi01; Camponotus
and scars caused by their secrevariegatus, and Brachymyrmex
tions, although they are cosmetic
obscurior.
and do not affect the fruit pulp,
Hawaiian apple banana (Dwarf Brazilian ‘Santa
On the east side of the Big
can make the fruits unmarketCatarina’ variety) fruits with the typical sympable. Here we discuss the ants toms of formic acid injury caused by ants. Also Island, the ant most commonly
and the damage they cause to shown along the upper edge of the center fruit associated with banana dambananas, and suggest integrated is a slight “corky scab” injury caused by the age is the yellow crazy ant, A.
management practices to reduce feeding of flower thrips (Thrips hawaiiensis). The gracilipes. In Hawai‘i, this ant
thrips injury, although similar in color, is raised
is also known as the longlegged
or avoid costly injury.
and corky in texture, not smooth and sunken as
ant. Crazy ants have a broad
Ants belong to the family with formic acid injury.
diet. They prey on a variety of
Formicidae, from the Latin word
arthropods, reptiles, birds, and
formica, meaning ant. They
mammals at soil level and within plant canopies. These
are arthropods in the order Hymenoptera, an order that
sweet-loving pests also feed on plant nectars, and they
also includes sawflies, bees, and wasps. Ant species in
farm and protect sap-feeding insects, including aphids,
the subfamily Formicinae produce formic acid (methascales, and mealybugs (Abbott et al. 2012). They infest
noic acid), which has the chemical formula HCOOH or
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in cooperation with the U.S. Department of Agriculture, under the Director/Dean, Cooperative Extension Service/CTAHR, University of Hawai‘i at Mānoa, Honolulu, Hawai‘i 96822.
Copyright 2011, University of Hawai‘i. For reproduction and use permission, contact the CTAHR Office of Communication Services, [email protected], 808-956-7036. The university is
an equal opportunity/affirmative action institution providing programs and services to the people of Hawai‘i without regard to race, sex, gender identity and expression, age, religion, color,
national origin, ancestry, disability, marital status, arrest and court record, sexual orientation, or status as a covered veteran. Find CTAHR publications at www.ctahr.hawaii.edu/freepubs.
UH–CTAHR
IP-29 — Jun. 2012
Ants that produce formic acid are attracted to flower
nectaries and to sap-feeding insects that secrete
honeydew. They climb the plants to feed and, when
startled, eject formic acid from their abdomens, causing
blackened spots and trails.
Entire bunches may be damaged by formic acid injury.
Startled and disturbed ants scatter over the banana
fingers, spraying formic acid and leaving burnt, sunken
trails.
Severe formic acid injury to a hand of bananas in East
Hawai‘i caused by Anoplolepis gracilipes.
2
UH–CTAHR
banana bunches to feed at flower nectaries and on honeydew secreted by sap-feeding arthropods.
When they eject formic acid for offensive or defensive purposes, damage to agricultural crops can also occur. A. gracilipes can cause cosmetic, or superficial, but
nonetheless significant damage to banana fruit bunches
when their colonies are disturbed or agitated by bunch
spraying with pesticides or when banana pseudostems receive vibrations through contact with humans or tractors.
The startled, disturbed ants scramble around, spraying
formic acid as they run, which burns the banana skins,
leaving irregularly shaped, sunken, blackened lesions.
Another ant species—the white-footed ant, Technomyrmex albipes—caused similar fruit damage at a farm
in Waimänalo. However, this canopy-nesting ant belongs
to a different subfamily. These ants are dolichoderines,
known for producing odiferous defensive compounds,
some of which may be either acidic or damaging to plant
tissues. This ant species mainly secretes benzaldehyde
(Hayashi and Komae 1980), which presumably also damaged the banana skins.
Other ant species in the subfamily Formicinae in
Hawai‘i may also damage banana fruits. For example,
Plagiolepis alluaudi is a very common ant in plant
canopies, tending various Hemiptera. Paratrechina
longicornis and possibly two Nylanderia species may
also be associated with banana plants. The latter three
species, however, are often outcompeted by a few of the
more dominant ant species. They may, therefore, not
be the main ant species tending sap-feeding insects or
feeding at banana flower nectaries and thus not the species causing the primary damage. A further ant species,
Pheidole megacephala, also tends various Hemipterous
insects attracted to banana plants and nectaries, but it has
not been observed damaging banana fruit.
Symptoms and Damage
The secretions of these ants create dark brown to
charcoal-black trails and spots on the skins of banana
fruits. The trails are irregular in shape and may be linear,
curved, serpentine, or semi-circular, coinciding with
the movement of the running, spraying ants. Spots may
vary in size from 2 to 6 mm in diameter. The trails can
be several millimeters wide and up to 12 mm long. All
banana varieties are susceptible.
IP-29 — Jun. 2012
Affected fruits, and often whole bunches, produced
at commercial banana farms in Hawai‘i cannot be sold.
They are destroyed in the field, resulting in an economic
loss for each affected bunch. Since the edibility of the
fruit is not affected, however, ant-damaged fruits may be
found at farmers’ markets in Hawai‘i, though even here
the value is reduced if injury is severe.
Management
When managing ant pests, suppression is preferable
to eradication. Another ant species, perhaps an even
worse pest, will usually colonize the niche vacated by
an eradicated ant species. Suppression of an ant species
can reduce pest injury to acceptable levels while allowing
the ant to fill the ecological niche.
The following techniques are suggested for managing ants that damage banana fruits in Hawai‘i. Also included is some of the rationale behind these approaches.
• Always identify the ant species before starting an antmanagement program. Ant behavior, biology, ecology,
and susceptibility to insecticides vary among species.
For example, T. albipes can spread from plant to
plant without contacting the ground, so groundbased treatments are not effective. Photographs and
species descriptions are available at www.antweb.
org (AntWeb 2012) or the Hawaii Ant Lab, www.
littlefireants.com/index_files/ant_key.htm
• Confirm that ants are causing the observed damage.
Some other insect pests of banana, such as thrips
or moths, may cause feeding injuries that resemble
the symptoms of formic acid injury caused by ants.
• Avoid disturbing the foraging ants within a banana
bunch. Do not jar the ant colony by bumping into the
banana pseudostem with your body or with tools or
equipment. The ants perceive the jarring vibrations
as a threat, causing them to disperse rapidly and
spray trails of formic acid on fruits as they scatter.
Reduce banana pesticide spraying operations where
possible, as forceful sprays near bunches can disturb
the ants.
• Control sap-feeding insects in the banana canopy,
including aphids, scales, and mealybugs. This may
include the application of insecticides.
3
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IP-29 — Jun. 2012
Affected bunches are left in the field and are not
harvested.
De-flowering the fingers on a banana bunch by plucking
them off and severing the male flowers (the hanging
“bell”) will remove the sweet flower nectaries that attract
sugar-loving ants in the subfamily Formicinae.
Fruits in bunches should be de-flowered to make them
less attractive to foraging ants.
The flowers shown here may attract sweet-loving ants
that may produce formic acid secretions when disturbed.
This “bell” should be severed from the bunch.
4
UH–CTAHR
• Use insecticidal baits as appropriate. For instance,
they may not be effective in reducing or destroying
colonies of A. gracilipes, as ant baits registered for
banana in Hawai‘i do not attract A. gracilipes. Test
a small amount of your intended insecticidal bait to
see if the ants will carry it back to their nest before
applying it to a large area. Some organic and backyard growers may prefer to use plastic bait stations
containing a boric acid solution, but this pesticide is
not registered by the Hawai‘i Department of Agriculture. T. albipes is difficult to control chemically
because the workers do not carry food (or bait) back
to the colony. Hence, baits must be very appealing
to the ants so that large numbers of them will leave
the nest and feed on the poisoned bait. • Destroy nesting habitats for ants that produce formic
acid. Pick up, remove, and compost plant litter such
as banana leaves and fallen pseudostems, as ants
such as A. gracilipes form nests beneath the litter
(O’Dowd 2012). Periodically replace the old boric
acid or other mixture with fresh bait.
• Scout areas around banana plants regularly for signs
of ants. Smaller and more localized ant colonies are
easier to control than larger infestations.
• Regularly de-flower young banana fingers and sever
the male inflorescence (the “bell”). This will remove
the flower nectaries and thereby make the young
fruits less attractive to the sugar-loving ants that forage on the bunch. A ladder may be needed to reach
the developing bunch.
• For canopy-nesting ants such as T. albipes, practice
field sanitation (Tenbrink and Hara 1992). The removal of touching banana leaves between plants may
slow the spread of the ant in the plantation.
IP-29 — Jun. 2012
Acknowledgements
The authors thank Fred Brooks and Paul Krushelnycky
of UH-CTAHR for their thoughtful reviews of this
manuscript.
References
Abbott, K, Harris, R, and Lester, P. 2012. Invasive Risk
Assessment: Anoplolepis gracilipes. Biosecurity
New Zealand. http://www.biosecurity.govt.nz/files/
pests/invasive-ants/yellow-crazy-ants/yellow-crazyant-risk-assessment.pdf
Ant Web. Subfamily: Formicinae. The California Academy of Sciences. http://www.antweb.org/description.
do?name=formicinae&rank=subfamily&project=ha
waiiants (accessed 8 May 2012)
Hawaii Ant Lab, www.littlefireants.com/index_files/
ant_key.htm
Hayashi, N, and Komae, H. 1980. Components of the ant
secretions. Biochemical Systematics and Ecology
8:293–295.
O’Dowd, D. Global Invasive Species Database: Anoplolepis gracilipes. http://www.issg.org/database/
species/ecology.asp?si=110. Centre for Analysis and
Management of Biological Invasions, Australia &
IUCN/SSC Invasive Species Specialist Group (ISSG)
(accessed 8 May 2012).
Peng, RK, and Christian, K. 2007. Integrated pest management in mango orchards in the Northern Territory Australia, using the weaver ant, Oecophylla
smaragdina, (Hymenoptera: Formicidae) as a key
element. International Journal of Pest Management
51:149–155.
Tenbrink, VL, and Hara, AH. 1992. Technomyrmex albipes (Fr. Smith). Crop Knowledge Master. University
of Hawai‘i at Manoa http://www.extento.hawaii.edu/
kbase/crop/Type/technomy.htm
• Another, potentially less harmful ant species that
competes for the same ecological niche may naturally displace an injurious ant species over time.
However, it may be unwise for growers to attempt to
introduce a competing ant species to a farm or site
without proper training and sufficient understanding
of the potential ecological or social consequences,
which could be dire.
5
Insect Pests
July 2005
IP-21
Banana Moth as a Pest of Coffee
Scot Nelson,1 Virginia Easton Smith,2 and Mark Wright1
Departments of Plant and Environmental Protection Sciences and 2Tropical Plant and Soil Sciences
1
B
anana moth, Opogona sacchari (Bojer), is a sig
nificant pest of coffee bark tissues and young ver
tical branches in Hawaii. The moth’s larvae feed upon
the cambium, vascular system, and pith within the green
verticals and on the cambium and phloem beneath the
exfoliating bark of the main trunk.
The banana moth is a threat to coffee in Hawaii be
cause its feeding can cause the death or weakening of
large numbers of young coffee verticals and can disin
tegrate large patches of coffee stem bark. Substantial
losses in crop yield and overall reductions in the health
of coffee plant populations may result.
Significant damage occurred in recent years at some
coffee farms in the Kona districts, located from approxi
mately 1200 to 2400 feet elevation. The damage was
locally severe and patchy, associated mainly with plants
recovering from pruning. Here we describe and docu
ment the damage to coffee. We also suggest some inte
grated pest management practices for coffee farmers to
adopt to control the banana moth.
pillars bore into the plant, eventually producing the char
acteristic frass deposits shown in Figure 2. Fully devel
oped caterpillars removed from their tunnels will be 3⁄4 –
11⁄8 inches (2–3 cm) long and somewhat transparent—it
is even possible to see some of their internal organs.
A distinguishing characteristic of the larvae is the
presence of brown patches on the top of the caterpillar
and dark brown “breathing pores” along the sides of the
body. The caterpillars pupate within the plant. Follow
ing the emergence of the adult moths from the pupae,
empty pupal cases may be observed protruding from the
1. Adult banana moth with empty pupal case. (photos: A. Hara)
Banana moth biology and ecology
Opogona sacchari has a wide distribution, occurring in
the Americas, Africa, and many islands throughout the
world. It was accidentally introduced to Hawaii and is
known to occur on Oahu and Hawaii. The larvae of O.
sacchari are generally considered to be scavengers, feed
ing in dead plant material. In Hawaii they are best known
as pests of sugarcane, where they damage the “eyes”
(buds) of the plants, but they also attack many orna
mental plants. The adult moths are 3⁄4 –1⁄2 (10–15 mm)
long and have grayish-brown wings, each with two small
but prominent black spots (Figure 1). When the moths
are at rest, their antennae point forward, rather than back
ward over the wings or next to the abdomen.
The moths lay their tiny eggs into crevices on plants;
they hatch after about a week. The newly hatched cater
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in cooperation
with the U.S. Department of Agriculture. Andrew G. Hashimoto, Director/Dean, Cooperative Extension Service/CTAHR, University of Hawai‘i at Mänoa, Honolulu, Hawai‘i 96822.
An equal opportunity/affirmative action institution providing programs and services to the people of Hawai‘i without regard to race, sex, age, religion, color, national origin, ancestry, disability,
marital status, arrest and court record, sexual orientation, or status as a covered veteran. CTAHR publications can be found on the Web site <http://www.ctahr.hawaii.edu/freepubs>.
UH–CTAHR
stems of the infested plants. The life cycle is likely to
take about 40–45 days under summer conditions in Ha
waii. A considerable number of generations per year can
be produced.
Early damage by O. sacchari is hard to detect be
cause little frass will have been pushed out of the tunnel
in the early stages of attack. However, as the hatched
caterpillars continue to feed inside the stems, they hol
low them out. Thereafter, evidence of their presence will
become clear, both because of the presence of frass (Fig
ure 2) and the wilting of young coffee stems.
Where banana moth populations are large and in
creasing aggressively, attacks can kill coffee verticals
and partially disintegrate the bark tissues of the main
trunk. Severely affected vertical branches can wilt, col
lapse, and detach from the trunk (Figure 3). Infested
verticals are prone to snapping off during strong winds.
Less severely affected branches or branches in early
stages of infestation may grow poorly and be structur
ally weakened.
Foliar re-growth of coffee plants that have been
pruned or stumped is particularly susceptible to damage
caused by colonization and feeding injury (Figure 4).
Egg-bearing female moths are attracted to the wounded
coffee stumps. Such plants are weakened by stress and
have moist, dead, or dying tissues preferred by the moth
for egg-laying. The female banana moth prefers to lay
her eggs in necrotic plant tissues and will lay them in
wide range of plant species.
Moths are attracted to natural openings in the bark
of trunks of pruned coffee. Secondary branches emerge
from the primary stem after pruning, and the bark
“erupts” to allow the emergence of verticals; a natural
opening or hole occurs in the wood to allow the mer
istematic tissue beneath the bark to emerge and grow
through.
Miniscule chambers of decomposing, sloughing
bark occur next to the emerging branches. The cham
bers of necrotic tissue are perfectly suited to protect and
nurture the laid and hatching Opogona eggs. Females
prefer to deposit eggs within these decomposing wounds,
or within natural openings in the bark.
The most dangerous egg-laying site for coffee is at
the emergence junction between a secondary green stem
and an older, woody stem. The hatching larvae are very
close to the tender new branch and can easily enter in
side. They tunnel up from underneath the emerging
branch and into the pith, never having been exposed to
predators on the surface of the plant.
The hatching moth larvae are whitish caterpillars
that feed on the tender green tissues just beneath the
2. Left: Granular, light-brown frass produced by the banana moth caterpillar larvae at
the base of young coffee vertical branches. Right: The damaged branch is easily
detached. The black circle of dead tissue is evident around the perimeter of the branch;
it indicates where banana moth larvae were feeding. In this case, larvae did not
penetrate the center of the stem and did not create a tunnel; the larvae caused
structural damage that weakened but did not kill the vertical.
2
IP-21 — July 2005
3. Wilting and collapse of newly emerged
vertical branch on a recently pruned coffee
plant. A banana moth larva was feeding
within a tunnel in the affected vertical,
having hatched from an egg laid near the
base of the stem where it emerged through
the woody tissue of the trunk.
UH–CTAHR
woody surface of coffee stems. They also feed within
young, non-woody coffee verticals (Figure 6). Larvae
feed and create tunnels up to three inches long within
the young stem. This effectively severs the vascular sys
tem and interrupts the flow of water to the branch. Wilt
ing and collapse quickly follow. Although banana moths
can attack plants at all stages of development, signifi
cant damage to bearing or desired coffee verticals oc
curs during their first year of re-growth after pruning.
Secondary damage occurs to coffee plants as other in
vasive organisms, including insects and fungi, replace
the maturing banana moth larvae.
A reliable indicator of banana moth populations is
the presence of the characteristic piles and elongated
mounds of light-brown frass that accumulate copiously
on coffee stumps and on debris (Figure 7).
Banana moth integrated pest management
Manage the banana moth on coffee with on-time, integrated
management actions. Following are suggested tactics.
IP-21 — July 2005
Accurate diagnosis and assessment
Coffee verticals are susceptible to a number of signifi
cant pests, and an accurate diagnosis of the cause is es
sential. Contact the UH-CTAHR Cooperative Extension
Service for assistance in pest identification and for an
evaluation of damage. Learn to recognize the frass and
damage caused by this insect. Learn to recognize the
larval stage of the banana moth. Scout coffee fields on a
regular basis for damage, and keep systematic records
of your observations. To detect the banana moth, look
for signs of frass being pushed out of stems, and gently
bend stems by hand (severely infested stems will tend
to collapse under pressure and break, rather than bend
ing evenly).
Field sanitation
Remove pruned coffee branches and trimmings from
fields and destroy them immediately. Chipping the
woody material, for example, can destroy larvae em
bedded within and remove the material as a source of
4. Granular, light-brown frass of the banana moth larvae on the coffee bark surface indicates the location of their feeding sites
under the bark on stumped coffee plants. The newly emerging vertical branches are highly susceptible to damage, and those in
the photo at left are under attack. The wounds created by pruning can attract the gravid moth females to lay eggs.
3
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IP-21 — July 2005
5. Banana moth frass at the intersection of vertical branch and the main coffee trunk. The base of the vertical branch shows a
blackening from larval feeding. When the lateral branch is pulled away it separates from the trunk easily, and the burrowing hole
of the moth becomes visible. Banana moth larvae tunnel inside the lateral branch to about three inches or more. This is sufficient
damage to cause the wilting, collapse and death of the vertical branch.
attraction for egg-laying banana moths (a beneficial
byproduct is mulch). Banana moth populations can de
velop on pruned coffee materials that are left on the
ground and in the field within coffee rows (Figure 6).
Spray pruned plants with Bt or pyrethrin
Drench the bark and the newly emerging verticals with
applications of Bacillus thuringiensis (Bt) shortly after
pruning and periodically thereafter as needed to achieve
economic control. Proper spray timing is important to
achieve best results. It probably is not necessary to spray
Bt on plants which have not been pruned recently. For
established infestations of the banana moth, supplement
the use of Bt with pyrethrin sprays, which are useful as
contact insecticides.
Selection of pruning method
More damage has been reported at farms using the Beau
mont-Fukunaga pruning method than using the Kona style
of pruning. However, more information on this is needed.
6. Banana moth caterpillar in tunnel within young coffee vertical.
4
Minimize plant stress (maintain plant vigor)
Plants that suffer from nutritional deficiency, root prob
lems, nematodes, drought, or physical or chemical inju
ries may recover slowly after severe pruning; vertical
branches that do not re-grow vigorously are not as tol
erant of banana moth feeding injury.
Remove suckers
Side branches emerging from coffee-bearing verticals can
harbor larvae of the banana moth. Populations of the moth
can be reduced by timely, periodic removal and destruc
tion of unwanted, infested suckers from these plants.
7. Rows of coffee plants were pruned at a coffee farm in 2002
with the Beaumont-Fukunaga method, stumping. The severed
coffee foliage was discarded on the ground, between plants
within rows. In the following months, banana moths fed on the
discarded materials, as evidenced by the large amounts of
the characteristic frass which accumulated on them and the
presence of banana moth larvae embedded within them.
Insect Pests
Sept. 2005
IP-24
Banana Moth—A Potentially Fatal Pest
of Pritchardia and Other Palms
Scot Nelson and Mark Wright
P
ritchardia species, some endemic to the Hawaiian
Islands, are among the most valued and cherished
palms. A few species are quite rare. Growers expend
significant resources to acquire these plants and grow
and maintain them in tropical landscapes. Therefore, any
significant Pritchardia pest or disease problem must be
dealt with effectively to protect the investment of time,
human resources, and capital.
Plant stress arising from nutritional deficiencies,
especially deficiencies of potassium and magnesium, and
other factors is a ubiquitous problem for Pritchardia
species in some Hawaiian landscapes. Other stresses,
including herbicide injury, drought, shallow soil, plant
ing in blue-rock, flooding, and mechanical wounding of
stems, can place the palm’s physiology under great strain
during establishment in landscapes after outplanting.
Plants so weakened can become targets for the ba
nana moth, Opogona sacchari (Bojer), perhaps the most
important insect pest of Pritchardia in Hawaii. Adult
females lay eggs in wounded or compromised
Pritchardia tissues. The caterpillar larvae hatch and feed
voraciously on the living and decaying tissues of the host,
and this can cause extensive damage. Where enough eggs
are laid in the youngest leaves of a plant, a fatal heart rot
disease caused by caterpillar feeding may ensue. This
condition and its management are described here.
The banana moth’s morphology and life cycle
The banana moth is a significant pest of many plants in
Hawaii, including sugarcane, banana, and pineapple.
Substantial losses in crop yield, overall reductions in
plant health, and even plant death may result from ba
nana moth infestations.
O. sacchari has a wide distribution, occurring in
many island locations throughout the world and in the
Americas and Africa. It was accidentally introduced to
A young Pritchardia hillebrandii plant in a Hawaii landscape,
dying from attack by banana moths, Opogona sacchari. The
heart of this plant is completely rotten.
Hawaii and is known to occur on Oahu and Hawaii.
The larvae of O. sacchari are generally considered
to be scavengers that feed on decaying and dead plant
material. However, they also can colonize living tissues.
In Hawaii they are perhaps best known as pests of sug
arcane, where they damage the “eyes” (buds) of the
plants, but they also attack many other ornamental and
Published by the College of Tropical Agriculture and Human Resources (CTAHR) and issued in furtherance of Cooperative Extension work, Acts of May 8 and June 30, 1914, in cooperation
with the U.S. Department of Agriculture. Andrew G. Hashimoto, Director/Dean, Cooperative Extension Service/CTAHR, University of Hawai‘i at Mänoa, Honolulu, Hawai‘i 96822.
An equal opportunity/affirmative action institution providing programs and services to the people of Hawai‘i without regard to race, sex, age, religion, color, national origin, ancestry, disability,
marital status, arrest and court record, sexual orientation, or status as a covered veteran. CTAHR publications can be found on the Web site <http://www.ctahr.hawaii.edu/freepubs>.
UH–CTAHR
Banana Moth on Prichardia and Other Palms
food crop plants.
The adult moths are 10–15 mm (3⁄8–5⁄8 inch) long
and have grayish-brown wings with two small but promi
nent black spots on each wing. When the moths are at
rest, their two antennae are folded over the wings or lie
next to the abdomen. Adult females lay tiny eggs into
naturally existing or wound-created crevices on plants;
the eggs hatch in about a week.
Upon hatching, the young caterpillars (larvae) bore
into the plant, eventually producing characteristic frass
deposits. Fully developed caterpillars removed from their
tunnels measure 20–30 mm (3⁄4–11⁄8 inches) long and are
somewhat transparent; it is possible to see some of their
internal organs with the naked eye. A distinguishing
characteristic of the caterpillar is the presence of brown
patches on its top and dark brown “breathing pores”
along its sides.
The larvae pupate within the plant. Following the
emergence of the adult moths from the pupae, empty pu
pal cases may be observed protruding from the stems or
other tissues of the infested plants. The life cycle is com
pleted in about 40–45 days under summer conditions in
Hawaii, a bit longer in cool weather. Thus, a consider
able number of generations per year can be produced.
Damage to Pritchardia palms
The severed petioles of pruned Pritchardia plants at
tract the wound-seeking banana moths that are search
ing for a suitable place to lay their eggs. These plants
also have a large number of natural openings, protected
crevices, and naturally decaying tissues all over the stem.
Adult female moths lay eggs in these locations, and moth
populations begin to increase on the infested plant.
The banana moth’s feeding causes stress for the
Pritchardia plants. Other stressors may be present, such
as drought, weed-whacker damage, fertilizer burn, water
logging, and herbicide or pesticide injury. These factors
contribute to a decline in plant health. This process can
take many months to develop into a significant problem.
Eventually, where plants are thus weakened or where
moth populations are particularly large, young heart
leaves come under attack by the voracious moths. These
succulent, nutrient-rich tissues are particularly suscep
tible to the moths’ feeding, and severe damage can occur
rapidly. Because palms are monocots, with their grow
ing point at the center of the base of the stem, when the
heart-leaves are destroyed, plant death is sure to follow.
2
IP-24 — Sept. 2005
Adult banana moth, Opogona sacchari, and pupal case (magni
fied about 6 X; actual size about 1⁄2 inch long). Photo by Arnold Hara.
Necrotic heart leaves of a dying Pritchardia hillebrandii plant
infested with banana moths.
Necrotic heart leaves (above) and an adjacent leaf (below),
easily pulled out from the center of a dying Pritchardia
hillebrandii plant infested with banana moths. Opportunistic
fungi infest the tissues, accelerating their decomposition.
(Text continued on p. 4.)
UH–CTAHR
IP-24 — Sept. 2005
Significant heart rot of a young Pritchardia hillebrandii plant
infested with banana moth larvae.
A declining Pritchardia hillebrandii plant infested with banana
moths. Rotten tissues are easily detached from the base of
the plant, revealing necrosis of living stem tissues beneath.
Several mature leaves show of signs of stress. From a distance,
the plant may appear to relatively healthy at this stage of
decline. However, upon closer inspection, it becomes evident
that the heart leaves are also rotting.
Banana moth larvae found feeding within an affected heart of
a declining Pritchardia hillebrandii plant.
Opportunistic fungi and caterpillar frass pellets on the surface
of a rotting Pritchardia hillebrandii leaf petiole.
3
UH–CTAHR
Management of the pest
The following management practices may minimize the
detrimental effects of banana moth attacks on
Pritchardia.
Minimize plant stress
Plants under stress are very susceptible to attack by the
banana moth. The most dangerous and common stress
factor in this regard is drought. A second important stress
factor is poor plant nutrition. Keep plants well irrigated
and properly fertilized. Do not let potted palms become
too dry. Use soil testing results to help guide fertilizer
practices. Avoid using herbicides near Pritchardia plants
in landscapes if possible. Although herbicides are safe
to use around most palms, in some cases or for certain
species problems may arise when plants are contacted
by herbicide sprays. Avoid over-pruning of leaves, and
treat pruned surfaces with an approved insecticide such
as one derived from Bacillus thuringiensis (“Bt”). Both
B. t. kurstaki and B. t. aizawai have broad registrations
among ornamental plants.
Intercrop
Host-finding by the moth may be more difficult in di
verse plantings. Therefore, avoid monocropping palms
or placing susceptible plants in exposed positions in
barren landscapes. However, the banana moth is so
polyphagous (has such a wide host range) that intercrop
ping might not work where other hosts for the moth are
present.
4
IP-24 — Sept. 2005
Use approved insecticides
Check with your nearest Cooperative Extension Service
office for the latest information on registered insecticides
that may be sprayed on Pritchardia plants to protect them
from banana moth attack. The most effective products
are probably pyrethroids, useful as contact insecticides
after infestations develop, and Bt products, which have
some residual and preventive effects as moths feed on
tissues that received Bt spray applications.
Learn to recognize the banana moth and symptoms
of its damage. Scout Pritchardia plants regularly for
moths and moth damage. Treat plants as described.
Alternate hosts
Watch for the buildup of moth populations on alternate
hosts in the vicinity of Pritchardia palms. Banana moths
have few qualms about what they eat, it seems. They
occur in grasses, banana, and coffee and can be found
wherever there is decaying vegetation.
Damage to other palms
Banana moth damage is not confined to Pritchardia
palms. Other palm species may also be subject to attack
and show symptoms similar to those described here,
according to the UH-CTAHR Agricultural Diagnostic
Service Center. Mortality due to banana moth attack of
the following palm species has been observed in Ha
waii in recent years: floribunda palm; foxtail palm
(Wodyetia bifurcata); Manila palm (Veitchia merrillii),
and coconut palm (Cocos nucifera). The point of moth
entry is usually either into the young heart leaves or some
place along the stem in natural openings or wounds.

IP-1 taro root aphid - Advanced Crop Science 55

Transcription

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