Title: Florida Entomologist
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Permanent Link: http://ufdc.ufl.edu/UF00098813/00077
 Material Information
Title: Florida Entomologist
Physical Description: Serial
Creator: Florida Entomological Society
Publisher: Florida Entomological Society
Place of Publication: Winter Haven, Fla.
Publication Date: 1988
Copyright Date: 1917
Subject: Florida Entomological Society
Entomology -- Periodicals
Insects -- Florida
Insects -- Florida -- Periodicals
Insects -- Periodicals
General Note: Eigenfactor: Florida Entomologist: http://www.bioone.org/doi/full/10.1653/024.092.0401
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Bibliographic ID: UF00098813
Volume ID: VID00077
Source Institution: University of Florida
Holding Location: University of Florida
Rights Management: Open Access

Full Text

(ISSN 0015-4040)


(An International Journal for the Americas)

Volume 71, No. 2 June 1988


MERSIE, W., AND M. SINGH-Absorption, translocation, and metabolism of "C-
thuringiensin (p-exotoin) in snapbeans ................ ......................... 105
raphy of Anastrepha ludens A. obliqua, and A. serpentina (Diptera te-
phritidae) in Mexico ...................................................................111
EGER, J. E., JR.-A new species of Acrosternum Feiber, Subgenus Chinavia
Orian, from Ecuador (Heteroptera: Pentatomidae: Pentatomini) ......... 120
SNIDER, R. J.-DenisieUa lithophila, a new species from a granite outcrop in
Georgia (Collembola: Sminthurididae) .............................................. 124
JIR6N, L. F., J. S. SOTO-MANITIU, AND A. L. NORRBOM-A preliminary list
of the fruit flies of the genus Anastrepha (Diptera: Tephritidae) in Costa
R ica ............................................................................................. 130
HALL, D. G.-Insects and mites associated with sugarcane in Florida ............ 138
SCARBROUGH, A. G.-A new species of Efferia Coquillett (Diptera: Asilidae),
staminea species group, from Grand Cayman Island, West Indies .......... 150
fruit fly Ceratitis capitata: behavior in nature in relation to different
Jackson traps ............................................................................... 154
ants of the Florida Keys ................................................................. 163
Bibliography of the neotropical cornstalk borer, Diatraea lineloata
(Lepidoptera: Pyralidae) ................................................................. 176
CHAN, K. L., AND J. R. LINLEY-Description ofAtrichopogon wirthi new species
(Diptera: Ceratopogonidae) from leaves of the water lettuce (Pistia
stratiotes) in Florida ...................................................................... 186
PRICE, R. D., AND K. C. EMERSON-Menacanthus dennisi (Mallophaga:
Menoponidae), a new species from the grey currawong (Passeriformes:
Cracticidae) in South Australia ..................................................... 202

Scientific Notes
E. HAGENBUCH, AND D. E. MILNE-Horn Fly (Diptera: Mus-
cidae) control using pyrethriod tags and tail tapes ..................... 205
HUBBARD, M. D., AND E. DOMINGUEZ-SynOnymy of the neotropical
mayfly genera Asthenopus and Asthenopodes (Ephemeroptera:
Polymitarcyidae: Asthenopodinae) .......................................... 207
HAAG, K. H., AND D. H. HABECK-A survey of waterhyacinth weevil
populations (Neochetina spp.) in northern Florida .................. 210
MITCHELL, E. R., R. W. HINES, AND W. W. COPELAND-Heliothis sub-
flexa (Lepidoptera: Noctuidae): Establishment and maintenance of
a laboratory colony ................................................................ 212
Continued on Back Cover

Published by The Florida Entomological Society


President ................................................................................ J. L. Taylor
President-E lect ................................................................... R. S. Patterson
Vice-President ............................ .............. .... J. E. Eger
Secretary ............................. .................... .. E. R. Mitchell
Treasurer ................. .. .... ...................... A. C. Knapp

D. J. Schuster
C. O. Calkins
Other Members of the Executive Committee ..............S. Osborne
J. H. Epler III
G. S. Wheeler
P. G. Koehler
J. R. McLaughlin


Editor ...................... .................................. ............ J. R. McLaughlin

Associate Editors
Arshad Ali Carl S. Barfield Ronald H. Cherry
John B. Heppner Michael D. Hubbard Lance S. Osborne
Omelio Sosa, Jr. Howard V. Weems, Jr. William W. Wirth

Business M manager ........... ................................ .. ................... A. C. Knapp

FLORIDA ENTOMOLOGIST is issued quarterly-March, June, September, and De-
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Membership in the Florida Entomological Society, including subscription to Florida
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Manuscripts and other editorial matter should be sent to the Editor, JOHN R.
MCLAUGHLIN, 462L NW 40th Street, Gainesville, FL 32606.

This issue mailed June 20, 1988

Mersie & Singh: Thuringiensin Metabolism


University of Florida, IFAS
Citrus Research and Education Center
700 Experiment Station Road
Lake Alfred, FL 33850 USA


The absorption, translocation, and metabolism of 'C-thuringiensin (p-exotoxin), an
insecticide derived from Bacillus thuringiensis, were investigated in snapbean
(Phaseolus vulgaris L.). The insecticide was applied to snapbean leaves or roots under
controlled environmental conditions. Thuringiensin was absorbed more by roots than
shoots. Snapbeans absorbed 12 and 17% of the root-applied thuringiensin after 3 and 7
days of application. Less than 3% of the foliar applied thuringiensin was detected in the
whole plant and amounts absorbed at 3 and 7 days were similar. More than 90% of the
root absorbed radioactivity remained in the root at both times of harvest. About 77 and
65% of the absorbed 14C-thuringiensin remained in treated leaf after 3 and 7 days of
foliar application, respectively. Time did not affect the distribution pattern of root or
foliar applied thuringiensin in different parts of snapbeans. In this study, thuringiensin
was not readily absorbed by roots or leaves of snapbean and had limited mobility. The
insecticide was not also readily metabolized by snapbean leaves after 3 and 7 days of
exposures. The practical implications of these results are discussed.


Se investig6 en el frijol, (Phaseolus vulgaris), la absorci6n, translocaci6n, y el
metabolismo de '1C-thuringiensin (p-exotoxin), un insecticide derivado de Bacillus
thuringiensis. El insecticide se aplic6 a las hojas y raices del frijol bajo condiciones
ambientales controladas. Thuringiensin fue absorvido mas por las races que por el
retofio. El frijol absorvi6 el 12 y 17% del thuringiensin aplicado a las raices despu6s de
3 y 7 dias de la aplicaci6n. Menos de un 3% del thuringiensis aplicado al follaje se detect
en toda la plant, y las cantidades absorvidas despues de 3 y 7 dias fueron similares.
Mas del 90% de la radioactividad absorvida por las raices se mantuvo en la raiz durante
las dos cosechas. Cerca del 77 y 65% del 14C-thuringiensin se mantuvo en las hojas
tratadas despues de 3 y 7 dias respectivamente de la applicaci6n foliar. El tiempo no
afect6 el patron de distribuci6n de thuringiensin aplicado a las raices y follaje del frijol.
En este studio, thuringiensin no fue absorvido por las raices y hojas del frijol y tuvo
una mobilidad limitada. El insecticide no fue metabolizado muy rapidamente por las
hojas del frijol despu6s de 3 y 7 dias de ser expuesto. Se discuten las implicaciones
prActicas de estos resultados.

Thuringiensin (p-exotoxin) is an insecticide produced by the entomopathogen bac-
teria, Bacillus thuringiensis. The insecticide is produced commercially by fermentation
and can be formulated as stabilized emulsion, wettable powder, or dust (Burges 1982).
Thuringiensin is a nucleotidic ATP analog (Benz 1966, Bond et al. 1969) which inhibits
the production of DNA-dependent RNA polymerase and consequently the production
of ribosomal RNA (Lecadet & De Barjac 1981). The insecticide has shown potential for
the control of insects on field crops, trees, ornamentals, vegetables, and stored grain
and grain products (Miller et al. 1983). It is most effective against immature Lepidopt-


Florida Entomologist 71(2)

June, 1988

era, Diptera, Coleoptera, Hymenoptera, Isoptera, and Orthoptera (Burgerjon & Mar-
touret 1971).
Although toxicity of thuringiensin has been extensively documented (Burgerjon &
Martouret 1971, Herbert & Harper 1985, Ignoffo & Gregory 1972, Sebesta et al. 1981),
there are few reports on its fate in plants. Wolfenbarger et al. (1972) reported that
thuringiensin was not readily absorbed by cotton (Gossypium hirsutum L.) leaves and
persisted on leaves for 6 to 12 days.
Thuringiensin is primarily a stomach poison and must be ingested to be effective
against plant eating insects. For maximum effectiveness, translocation to the plant
parts eaten by insects is essential. Information on the extent of thuringiensin uptake
by plants and its movement out of the site of application will help to determine how
effective it will be under different application methods. In this study, the uptake, trans-
location, and metabolism of thuringiensin in a potential target crop, snapbean were
Plant Culture

Snapbean variety 'Greencrop' (Asgrow Florida Co.) was grown from seed in a green-
house (24/20C day/night) in planter flats Styrofoamm flats 66 x 33 x 12.7 cm) containing
sand. Plants were watered and fertilized with half-strength Hoagland & Arnon (1950)
nutrient solution as needed.

4C-Thuringiensin Application

At the 3-leaf stage of snapbean, uniform plants were selected and transferred to
1-quart darkened jars with 900 ml half-strength Hoagland and Arnon solution with pH
7 to 7.5. The jars were then transferred to laboratory with 300/200C (day/night) temper-
ature and photosynthetic photon flux density (PPFD) of 200 RE.M.-2 S-1 for 14 h. After
48 h in the above nutrient solution, 100 jpl of 4C-thuringiensin (80,000 dpm with specific
activity of 0.72 gCi/mg, 92% radiochemical purity) was added to each jar. The thurin-
giensin used in this work was supplied by Abbott Laboratories (North Chicago, IL) as
ABG 6162, Lot XDZ41. Hoagland and Arnon solution was added to the jars every 2
days to maintain the original volume of the liquid and the roots were aerated by bubbling
air through the solution. For foliar application, 100 pl of "C-thuringiensin (80,000 dpm)
in 0.1% (v/v) X-77 (surfactant) was applied as droplets on the first true leaf.

Uptake and Translocation

Absorption and translocation of "C-thuringiensin in snapbean was determined quan-
titatively by combustion of plant parts. For root treatments, plants were harvested 3
and 7 days after treatment. The roots were rinsed twice in 25 ml of distilled water and
blotted dry. A 2 ml of aliquot of the root wash solution was added to 15 ml of scintillation
solution and assayed for radioactivity with a Beckman LS 5800 liquid scintillation
counter. The 14C remaining in the Hoagland and Arnon solution was also measured by
assaying 2 ml from each jar. The plants were divided into shoot apex, leaves, stem, and
root. The plant parts were dried at 500C for A8R k, weikged, and ground in a mi l with
40 mm mesh. Each 50 mg sample of the ground tissue was taken and combusted in a
sample oxidizer (OX-300 P. J. Harvey Instr. Corp., Hillsdale, NJ). The CO2 was trapped
in 15 ml of CO2 absorber and scintillation fluid and the radioactivity was quantified as
described aeQys dpm of R C-thuringiensin per 50 mg of dry plant tissue.
calculated as dpm of 14C-thuringiensin per 50 mg of dry plant tissue.

Mersie & Singh: Thuringiensin Metabolism 107

For foliar applied treatments, snapbean plants were harvested at 3 and 7 days after
treatment. The treated leaves of each plant were washed in 25 ml of distilled water.
The 14C in the leaf wash was assayed by a liquid scintillation counter from a 0.5 ml of
aliquot. The plants were separated into leaves above treated leaf, shoot apex, stem
above treated leaf, stem below treated leaf, leaves below treated leaf, and roots. The
14C in each plant part was quantified as described above.

Metabolism of Leaf Applied '1C-Thuringiensin by Snapbean Shoot

Ground shoot tissue saved from the absorption and translocation study was used for
metabolism. The shoot tissue was placed in 100 ml of distilled water and homogenized
with polytron homogenizer at high speed for a total of 5 min. The pellet was washed
with four 20 ml portions of distilled water and the wash solution was poured through
the Buchner funnel. The resulting filtrate was transferred to a 250 ml beaker and the pH
was adjusted to 9.5 with 1N NaOH. The filtrate was concentrated to approximately 3
ml by rotary evaporation under vacuum using a water bath at 400C. A 10 p1 sample of
the concentrated extract was spotted on 5 x 20 cm, 250 P, thin-layer chromatography
plates (microcrystalline cellulose). All spotted plates were developed in the solvent
system containing n-butanol:acetone:acetic acid: 5% NH4OH: distilled water (30:15:10:
10:35) for 15 cm. On separate TLC plates, standard 4C-thuringilensiln was spotted and
developed alongside plates with extract concentrates. The distribution of radioactivity
on plates was assayed by scintillation spectrometry after scraping fifteen 1 cm2 bands
from the origin to the solvent front into scintillation vials containing 10 ml of scintillation
cocktail. Quench corrections were made by an external standard method and counts
were converted to dpm/cm2/band. Two plates were used for each treatment and the
average of the 2 is presented.

Statistical Analysis

Experiments were repeated with at least 3 replications each time. An analysis of
variance was performed for the combined experiments. Where a significant F value was
found, treatment means were separated by the LSD test at the 0.05 probability level.


Thuringiensin Uptake

For the root application, the 14C recovered in the nutrient solution, root washing,
and whole plant was 95% of the applied amount for both times of exposure (Table 1).
Snapbean roots absorbed more of 4C-thuringiensin at 7 days compared to 3 days after
treatment. Root systems, mainly fibrous roots, were larger (370 mg/plant) at 7 days
than at 3 days (210 mg/plant) after treatment and this might have increased the absorp-
tion of the insecticide. About 74 and 61% of the applied '4C was found in the nutrient
solution 3 and 7 days after application, respectively. The root wash solution contained
about 9 and 17% of the applied radioactivity at 3 and 7 days after treatment, respec-
tively. On the whole, less than 20% of the 14C-thuringiensin was absorbed even 7 days
after treatment.
Similarly, less than 3% of the foliar applied insecticide was detected in the whole
plant following sourceleaf washing (Table 2) and snapbean had absorbed similar amounts
of 14C-thuringiensin 3 and 7 days after treatment. More than 90% of the applied radioac-
tivity was found in the leaf wash at both times of harvest which indicated that a large
proportion of the compound remained on the treated leaf surface without penetrating

Florida Entomologist 71(2)

June, 1988


Harvesting '4C in Hoagland's '4C in root
time solution washings 1"C absorbed Recovered
(days) % appliedb % appliedb % appliedb % appliedb

3 74.2 a 9.3 a 12.0 b 95.4 a
7 60.9 b 17.3 a 17.0 a 95.3 a

aData are from 2 experiments (6 replications).
bMeans in a column followed by the same letter are not significantly different as determined by the LSD test at
the 0.05 level of probability.

the leaf tissue. Unlike the root treatment, there was no significant difference in the
amount of leaf absorbed thuringiensin between 3 and 7 days of exposure.
Although snapbean absorbed more root applied than leaf applied thuringiensin, it is
apparent that thuringiensin is not readily absorbed by roots and leaves. Because this
insecticide is primarily targeted for leaf eating insects, foliar application would be of
major interest. Work with "~C-labeled compounds indicates that there are 2 routes by
which molecules may traverse the distance from the leaf cuticle surface into the living
inner cells: a lipoid route and an aqueous pathway (Crafts 1961). Compounds that enter
via the aqueous pathway penetrate slowly, and their penetration is greatly affected by
environmental factors, especially relative humidity. Thuringiensin is a nucleotide de-
rivative, with high water solubility and molecular weight of 700. From its physio-chem-
ical properties, thuringiensin would be expected to readily enter the leaf through the
aqueous phase.
Thuringiensin is not a contact poison and it should be ingested to be effective against
leaf eating insects. For this, it has to be absorbed by the leaf or remain stable on the
leaf surface. However, as our results showed, it is not readily absorbed, and the data
for the leaf wash indicated that it could easily be removed from the surface by water,
i.e., rain. In addition, the low absorption despite the use of the surfactant X-77 indicates
that wetting agents may not increase its penetration into the leaf tissue.

Thuringiensin Translocation

Quantitative measurements (Table 3) showed that more than 90% of the absorbed
radioactivity remained in the root. Higher concentrations of the insecticide were de-
tected in roots at 3 days (2,668 dpm/50 mg) than at 7 days (1,205 dpm/50 mg). This could
be attributed to the relatively faster growth of root than the shoot observed at 7 days
after treatment. It appeared that the higher root biomass after 7 days of growth had


Harvesting '4C in leaf washb '4C-absorbedb Recoveredb
time (days) % applied % applied % applied

3 95.5 1.4 96.9
7 96.0 2.2 98.4

aData are from 2 experiments (6 replications).
bMeans within a column were not significantly different as determined by the LSD test at the
0.05 level of probability.

Mersie & Singh: Thuringiensin Metabolism


3 days 7 days
% of absorbed % of absorbed
Plant part radioactivity radioactivity

Shoot apex 1.9b 3.1 b
Shoot 0.6b 0.9b
Stem 1.2 b 3.0 b
Root 96.2 a 92.8 a

aData are from 2 experiments (6 replications).
bMeans in a column followed by the same letter are not significantly different as determined by the LSD test at
the 0.05 level of probability.

reduced the concentration of the 4C when expressed per 50 mg dry tissue weight. This
dilution effect was evident from results given in Table 2.
For foliar applied thuringiensin, about 77 and 66% of the absorbed radioactivity
remained in the treated leaf at 3 and 7 days, respectively (Table 4). Data indicated that
there was more translocation out of treated leaf at 7 than 3 days of harvest. As indicated
by percent data at 3 days of harvest, similar amounts of '4C were detected in the
untreated plant parts but at 7 days more was translocated to leaves above treated leaf
than other above ground parts.
Maximum effective control of insects can be gained if a compound is translocated to
unsprayed plant parts. A lack of systemic action by thuringiensin means that the insec-
ticide should be applied in sufficient water to achieve adequate coverage or multiple
application may be required.

Metabolism of Leaf Applied 14C-Thuringiensin by Snapbean Shoot

Thin layer chromatography was found to be a sensitive detection method for thurin-
giensin with detection limits as low as 1 jg. The migration of snapbean shoot extract

GIENSIN (80,000 dpm/PLANT)a.

3 days 7 days
% of absorbed % of absorbed
Plant part radioactivityb radioactivityb

Treated leaf 77.4 a 65.5 a
Leaves above treated leaf 3.3 c 16.2 b
Shoot apex 0.5 d 1.7d
Stem above treated leaf 0.2 d 1.7 d
Stem/leaves below treated leaf 4.3 c 4.9 cd
Roots 14.4 b 10.0 b

aData are from 2 experiments (6 replications).
bMeans within a column with similar letters are not significantly different as determined by the LSD test at the
0.05 level of probability.

Florida Entomologist 71(2)


'4C-Distribution (% recovery)

Rf values 3 days 7 days standard

0.2 28 31 6
0.5 49 47 89
0.8 23 22 5

'Values are means of 3 chromatograms.
bStandard = authentic "C-thuringiensin.

on TLC plates were similar at 3 and 7 days exposures (Table 5). There was no evidence
for the metabolism of thuringiensin for either treatment time. Of the total '4C recovered
from the chromatograms of snapbean shoot extracts, 49 and 47% for 3 and 7 days,
respectively, were chromatographed with 4C-thuringiensin with Rf of 0.5. The 14C
retained at Rf 0.2 was 28 and 31 for 3 and 7 days exposure respectively. It is possible
that the 4C-labeled compound which remained at 0.2 may have been intact parent
insecticide which was bound to minute shoot residues which did not migrate from the
origin. "IC-recovered at Rf 0.8 were 23 and 22% for 3 and 7 days harvest, respectively.
There was no significant difference in the amount of radioactivity observed at 0.2 and
0.8 Rf values. Similarly, about 5% of the authentic thuringiensin was observed at 0.8
Rf and was similar to what was observed at the 0.2 Rf.
Thuringiensin is deactivated by dephosphorylation. In mammalian systems, the com-
bination of acidic pH and phosphatase are effective in breaking this compound; ho\, "ver,
in plants the compound is not readily absorbed and metabolized. Wolfenbarger et al.
(1972) reported that thuringiensin persisted in cotton leaves 6 to 12 days without degra-


Florida Agricultural Experiment Station Journal Series no. 8966.


BENZ, G. 1966. On the chemical nature of heat stable toxin of Bacillus thuringiensis
Berliner in Locusta migratoris. J. Invertebr. Pathol. 6: 381-383.
BOND, R. P. M., C. B. C. BOYCE, AND J. FRENCH. 1969. A purification and some
properties of an insecticide exotoxin from Bacillus thuringiensis Berliner.
Biochem. J. 114: 477-488.
BURGERJON, A. AND D. MARTOURET. 1971. Determination and significance of the
host spectrum of Bacillus thuringiensis. Pages 305-325. In H. D. Burges and N.
H. Hussey, Eds. Microbial control of insects and mites. Academic Press, New
BURGES, H. D. 1982. Control of insects by bacteria. Parasitology 84: 79-117.
CRAFTS, A. S. 1961. The chemistry and mode of action of herbicides. Interscience;
New York. pp. 38-39.
HERBERT, D. A. AND J. D. HARPER. 1985. Bioassay of a p-exotoxin of Bacillus
thuringiensis against Heliothis zea larvae. J. Invertebr. Pathol. 46: 247-250.
HOAGLAND, D. R. AND D. I. ARNON. 1950. The water culture method for growing
plants without soil. California Agric. Exp. Sta. Cir. 547. 32 pp.
IGNOFFO, C. M. AND B. GREGORY. 1972. Effects of Bacillus thuringiensis p-exotoxin
on larval maturation, adult longevity, fecundity and egg viability in several
species of Lepidotera. Environ. Entomol. 1: 269-272.

June, 1988


Celedonio-Hurtado et al.: Demography of Anastrepha

LECADET, M. M. AND H. DE BARJAC. 1981. Bacillus thuringiensis beta-exotoxin.
Pages 293-321. In E. W. Davidson, Ed. Pathogenesis of Invertebrate Microbial
Diseases Allanheld Osmum.
MILLER, L. K., A. J. LINGG, AND L. A. BULLA, JR. 1983. Bacterial, viral and fungal
insecticides. Science 219: 715-721.
SEBESTA, K., J. FARKAS, K. HORSKA, AND J. VANKOVA. 1981. Thuringiensin, the
beta-exotoxin of Bacillus thuringiensis. Pages 249-277. In H. D. Burges, Ed.
Microbial control of pests and plant diseases. Academic Press, New York.
1972. Properties of the p-Exotoxin of Bacillus thuringiensis IMC 10,001, against
the tobacco budworm. J. Econ. Entomol. 65: 1245-1248.


Program Mosca del Mediterraneo, DGSPAF, SARH
Apartado Postal 368
30700 Tapachula, Chiapas, MEXICO

Department of Entomology
University of California
Davis, CA 95616


Demographic parameters for Anastrepha ludens (Loew), A. obliqua (Macquart),
and A. serpentina (Wiedeman). reared on two artificial diets and several natural hosts
are reported. These include preadult survival and development rates, adult survival
and fecundity and population parameters such as the intrinsic rate of increase, mean
generation time and stable age distribution.
All three species displayed similar life history patterns. Egg development required
approximately three days, larval development 8-13 days, and pupation 13-17 days. The
number of eggs per female (gross fecundity rate) ranged from 84 to 102. A. serpentina,
compared with the other two species, displayed much higher survival in the first two
weeks of adult life. This led to a higher net reproductive rate despite a lower gross
fecundity relative to the other two species. The three species described here have
different demographic characteristics compared to those of the ecologically similar te-
phritids Ceratitis capitata and Dacus spp.


Se reportan los parametros demogrAficos de Anastrepha ludens (Loew), A. obliqua
(Macquart), y A. serpentina (Wiedemann) criadas en dos dietas artificiales y various
hospederos naturales. Los parametros reportados incluyen, la sobrevivencia y tasas de
desarollo de los estados inmaduros, la sobrevivencia y fecundidad de los adults, y
parametros de poblaci6n tales como la tasa intrinseca de incremento, el tiempo medio
de generaci6n, y la distribuci6n stable de edades.

Florida Entomologist 71(2)

Las tres species presentaron patrons similares en sus ciclos bi6logos. El desarollo
del huevo requiri6 aproximadamente tres dias, el desarollo de la larva de 8 a 13 dias, y
el de la pupa 13 a 17 dias. El nfmero de huevecillos por hembra (tasa bruta de fecun-
didad) vari6 de 84 a 102. A. serpentina, comparada con las otras dos species, present
una mayor sobrevivencia en las primeras dos semanas de vida en estado adulto relative
a las otras dos species. Esto lleva a una mayor tasa neta de reproducci6n a pesar de
una menor fecundidad. Las tres species descritas aqui tienen caracteristicas demog-
raficas diferentes comparandolas con aquellas de los tefritidos ecologicamente similares,
tales como Ceratitis capitata y Dacus spp.

The genus Anastrepha Schiner (Diptera: Tephritidae) is widely distributed in trop-
ical and sub-tropical regions of Mexico, Central America, and South America (Stone
1942). The group includes many important pests of fruit (Baker et al. 1944). Fruit
cultivation in Mexico occupies over 1.2 million hectares and in 1983 had a value of 800
million US dollars (SARH 1983). The 1985 ban by the United States on the use of
ethylene dibromide for the treatment of fruit exports has emphasized the need for
pre-harvest control methods for fruit flies.
Recent work on the demography of the Mediterranean fruit fly, Ceratitis capitata
(Wiedemann), suggests that demographic parameters are important for the develop-
ment of effective control programs, the establishment of efficient mass rearing facilities
for the sterile insect technique (SIT), and the interpretation of trap data (Carey 1982,
Carey & Vargas 1985). Considerable work has been reported on the demography of the
Mediterranean fruit fly (Carey 1984, Vargas et al. 1984), and Dacus spp. (Vargas &
Nishida 1985, Carey et al. 1985, Foote & Carey 1987). Thus, the comparison of life
history strategies in fruit-infesting tephritids may yield insight into the factors which
permit certain species to expand their geographical and host ranges to include many
commercially important fruits (Fitt 1986, Krainacker 1986).
The purpose of this project was to document the preadult and adult life history
parameters for three species of Anastrepha and to analyze these data using demographic
techniques. Despite the economic importance of the genus little or no systematic study
of these traits has been made (Baker et al. 1944, Christenson & Foote 1960, Bateman
1972). The three species studied were the Mexican fruit fly, A. ludens (Loew), the West
Indian fruit fly, A. obliqua (Macquart), and the zapote fruit fly, A. serpentina


Colony Sources and Rearing Conditions

Colonies of 70-150 adults were established from infested fruit collected in Chiapas,
Mexico. Aluja and collaborators (1984, 1987a, 1987b) have reported information on the
population dynamics, hosts and infestation patterns in the region. Flies were held in 60
x 30 x 30 cm cages at 28 3 oC, 75% R.H., and 12:12 LD. Adults received water and
a 3:1 mixture of sugar and yeast hydrolysate as food. All data were collected from Fl

Preadult Parameters

All three Anastrepha species received artificial fruit hosts composed of green paraf-
fin-coated nylon mesh in the form of half spheres. Eggs collected from these receptacles
were placed on moistened black cloth in petri dishes. Percent hatch and duration of the

June, 1988

Celedonio-Hurtado et al.: Demography of Anastrepha

egg stage were determined at 12-h intervals. Ten to twenty first instar larvae were
placed on about 15 g of artificial diet (Table 1) or in the pulp of one of several fruit hosts.
Fruits used included mango (Mangifera indica L.), jobo or mombin (Spondias mombin
L.), canistal or yellow zapote (Pouteria campechiana Baehni), mamee-apple or mamey
(Calocaropm sapota Merr.) star-apple or caimito (Chrysophyllum cainito L.), and
sapodilla or chicozapote (Manilkara achras Mill.). Fresh food was added as needed.
Late third instar larvae were allowed to pupate in sawdust, soil, or sugar cane husks.
Larval development time, pupal duration, and survival were recorded for each species.
Sample sizes are given in Table 2.

Adult Parameters

Fecundity and survival were recorded on separate cohorts of the three species. The
cohorts of A. ludens and A. obliqua consisted of 20 pairs and those of A. serpentina 28
pairs. Fecundity was determined by placing newly closed females with a male, trans-
ferring the females to small cages after copulation, providing food (3:1 mixture of sugar
and yeast hydrolysate), water, and an artificial fruit host for oviposition. Eggs were
counted daily.

Demographic Analysis

Fecundity, mortality, and developmental data for each species were used for stable
population analysis. When larval development rate was measured on more than one
host, the average development time over all hosts for the analysis was used. Egg hatch
was assumed to be 85% for this analysis. The reasons for this assumption are discussed
below. Carey (1982) defines the demographic parameters used in this study.


Preadult Parameters

Egg development times in days (- SD) were: 3.27 ( 0.76) for A. ludens, 3.14 (
0.72) for A. obliqua and 3.41 ( 0.96) for A. serpentina. Percent hatch ranged from
75% for A. ludens to less than 20% for A. serpentina. The low hatch success may have
been due to the susceptibility of eggs to mechanical damage during transfer from the



Wheat bran 12.4% 17.7%
Torula yeast 6.7 13.7
Soy 16.0 -
Sugar cane husks 8.2
Sugar 7.3 10.9
Sodium benzoate 0.6 0.5
Nipagin 0.2 0.5
Hydrochloric acid -0.5
Water 56.8 47.8
100.0% 100.0%

114 Florida Entomologist 71(2) June, 1988

artificial fruit hosts to petri dishes and to dehydration when transfer to the petri dishes
was delayed. Since we felt that this did not reflect biological differences in the life
history strategies of the flies, all subsequent demographic analysis assume a uniform
hatch rate of 85%. This is consistent with egg hatch results obtained with other species
of tephritids (e.g. Carey 1984). More detailed studies of percent egg hatch are required
to determine the relationship between egg hatch and population dynamics.
The duration and success of larval development and pupation for the three species
of Anastrepha reared on artificial diets and natural hosts is given in Table 2. Note that
the rates for all species are quite similar. Larval development required from 8 to 13
days and pupation from 13 to 17 days. The results for preadult survival and development
rates found in this study are similar to those reported by Baker et al. (1944), who
reported a combined egg and larval development time of 14 days at 26C for A. ludens.

Adult Parameters

Survival of the flies is given in Fig. 1. All three species show constant daily mortality
through the life of a cohort. A. serpentina had much higher survival than the other two
species during the first two weeks of adult life. This is reflected in A. serpentina's
higher net reproductive rate and greater expectation of life. In A. ludens and A. obliqua
a few individuals survived much longer than the remainder of the cohort. Male survival
was very similar to female survival in all three species.
Daily reproduction of the flies is shown in Fig. 2. The age at first reproduction was
approximately 10 days. Peak reproductive output occurred shortly after this. Reproduc-
tive parameters and the adult expectation of life are given in Table 3. These parameters
are based on the analysis of adult survival and fecundity only, without considering the
immature stages. Adult expectation of life (mean age of death) ranged from 17.3 for A.
ludens to 30.4 days for A. serpentina. Gross fecundity rate ranged from 83.9 eggs/fe-


Species/Host N X % X % X %

A. ludens
Metapa 12360 10.30 (3.70) 2.5 14.43 (1.61) 40.7 24.73 1.0
Xalapa 565 9.43(3.57) 43.9 16.60 (3.60) 75.0 26.05 32.9
Mango 2394 11.14 (3.66) 13.2 14.11 (0.96) 62.0 25.51 3.1
A. oblioua
Metapa 1025 13.06(5.79) 31.6 15.27(3.08) 97.2 28.33 30.7
Xalapa 201 8.63(1.44) 40.8 14.75 (1.02) 87.5 23.38 35.7
Mango 136 9.03(2.21) 56.8 12.94 (1.98) 52.2 21.97 29.6
Jobo 622 9.53(1.17) 4.5 14.00(0.05) 100.0 23.53 4.4
A. serpentina
Metapa 470 10.76(2.28) 40.0 13.68 (1.38) 53.9 24.64 21.5
Xalapa 967 13.48(5.73) 31.6 15.20 (2.97) 97.2 28.68 30.6
Canistal 103 1.56(3.07) 55.1 15.29(2.83) 95.9 26.85 52.8
Mamey 47 9.37(1.24) 51.2 15.86 (1.55) 85.1 25.23 43.9
Caimito 108 10.29(2.16) 40.8 13.33(1.42) 75.8 23.62 30.9
Sapodilla 130 8.77(2.61) 34.7 14.70(1.51) 55.9 23.50 19.4

Celedonio-Hurtado et al.: Demography of Anastrepha 115


A. ludens

.5 female

% male


A. obliqua



A. serpentina

.5 -



0 30 60 90 120

AGE (days)

Fig. 1. Survival schedules for 3 species of Anastrepha. The data are from wild adult
flies collected as larvae in fruit.

male for A. serpentina to 102 eggs/female for A. ludens. Net fecundity rate (mortality-
weighted fecundity) ranged from 46.8 to 76.3 eggs/female.
The population parameters considering the immature stages and percent in the
stable age distribution are shown in Table 4. The net reproductive rate (Ro) was 9.1
for A. ludens and A. obliqua and 16.5 for A. serpentina. The intrinsic rate of increase

Florida Entomologist 71(2)


A. ludens



SA. obliqua
E o10


10 A. serpentina


0 10 20 30 40 50

AGE (days)
Fig. 2. Gross fecundity schedule for 3 species of Anastrepha. The data are from wild
adult flies collected as larvae in fruit. Arrows refer to the day on which the last female
in the cohort died.

June, 1988

Celedonio-Hurtado et al.: Demography of Anastrepha 117


Meane Expec-
Eggs Age tationd
Grossa Netb per Repro- of
Species Rate Rate Day duction Life

A. ludens 101.8 46.8 4.24 19.4 17.3
A. obliqua 99.5 51.8 3.69 22.1 21.7
A. serpentina 83.9 76.3 1.68 16.7 30.4

gross fecundity rate is expressed as the number of eggs per female that lives to the last possible day of life.
bnet fecundity rate is expressed as the number of eggs per female, considering adult survival.
'mean age of reproduction is the age, in days, in which an average female laid half of the total number of eggs.
expectation of life is the average age of death expressed in days.

(r) ranged from 0.045 to 0.061. Doubling time in the stable population required from 11
to 34 days. The stable age distribution shows that 88 to 90% of the population will be
in the preadult stages.


The differences in larval developmental time and survival within a given species,
comparing the artificial diets and natural hosts, suggests that host-specific larval fitness
will serve as the overriding factor governing the growth potential of a population at
large (Carey 1984, Krainacker 1986). The low survival of A. ludens in Metapa diet and
mango, and A. obliqua in jobo, could be attributed to a number of reasons (eg. nutri-
tional, microbial contamination, temperature, maturity of fruit), but no specific causes
can be determined in this study.
Our results suggest that A. serpentina has a somewhat different life history strategy
compared with A. ludens and A. obliqua. It had a lower gross rate of egg production
and a higher life expectancy. The increased life expectancy--due to a higher survival
during the first two weeks of adult life--offset the lower egg production and led to a
higher net reproductive rate and a shorter mean age of reproduction.


Parameter" SADb (%)
Species R, r DT T E L P A0

A. ludens 9.1 0.047 14.7 46.9 24 42 24 10
A. obliqua 9.1 0.045 15.3 48.5 26 41 21 12
A. serpentina 16.5 0.061 11.4 46.2 25 45 20 10

"Ro = Net reproductive rate; r intrinsic rate of increase; DT = doubling time (days); T = mean generation
time (days).
bSAD = Stable Age Distribution.
'E = Eggs; L = Larvae; P = Pupae, A Adults.

118 Florida Entomologist 71(2) June, 1988

These three Anastrepha species all have longer developmental times and lower
fecundity than the Mediterranean fruit fly, the oriental fruit fly, and the melon fly
studied in Hawaii. (Carey 1984, Vargas et al. 1984, Vargas & Nishida 1985, Foote &
Carey 1987). For example, Carey (1984) reports a net reproductive rate of over 1000
eggs/female for C. capitata and larval to adult development time of about 20 days.
Vargas et al. (1984) obtained a net reproductive rate of over 250 and 400 eggs/female
and developmental time from larva to adult of 17 and 18 days for D. cucurbitae and D.
dorsalis respectively. Baker et al. (1944) report that A. ludens may lay as many as 400
eggs. All of these results were obtained from laboratory colonies. The lower fecundities
recorded in this study of Fl generation wild flies may be a result of the artificial egging
devices, to which the wild flies are not adapted, and the stress of laboratory environ-
ment (Leppla et al 1983).
Foote & Carey (1987) report a ten-fold increase in the fecundity of D. dorsalis lab
strains over wild strains. Perez (1987) found a net reproductive rate of 4.13 for wild A.
ludens compared with 10.24 for laboratory flies. On the other hand, developmental
rates did not differ between wild and laboratory strains.
It is not obvious why the egg, larval and pupal times are longer for these Anastrepha
species than for C. capitata and D. dorsalis. All these species exploit similar hosts. C.
capitata is native from a Mediterranean climate and is much smaller (Christenson &
Foote 1960). Tropical species are often larger than their temperate relatives (Young
1985) and larger species frequently take longer to develop (Bonner 1965). However,
Dacus species are similar in size and are thought to have originated in a tropical or
sub-tropical habitat (Christenson & Foote 1960, Fitt 1986, Fletcher 1987).
All three species described here are pests of economic importance that have success-
fully expanded their geographical and host ranges in conjunction with the spread of
commercial fruit cultivation (Aluja & Liedo 1986, Bateman 1972, Christenson & Foote
1960). These species are each reported on about forty or more different hosts in the
neotropics (Malavasi et al. 1980, Norrbom 1985, Wasbauer 1972). However, the actual
natural host range of these species is more restricted and varies from region to region
(Baker et al. 1944). For example, in Chiapas, Mexico, A. ludens is commonly collected
from fruits of the Rutaceae family, although it also infests mango (Anacardaceae) and
Mastichodendron capri var tempisque (A. C. D.) Cronq. (Sapotaceae). A. obliqua is
commonly collected from fruits of three species of the Anacardaceae family and three
species of the Myrtaceae family. A. serpentina is normally restricted to five species of
the Sapotaceae family (Aluja et al 1987b).
Zolfer (1983) characterizes the tephritid genera Anastrepha, Ceratitis, and Dacus
as consisting of highly fecund host generalists. However this study and recent work on
C. capitata and Dacus spp. demonstrate that there is a great deal of life history vari-
ation among these genera. The reasons for this variation have yet to be determined.


This work was supported by grants from the California Department of Food and
Agriculture, and ongoing support of the Programa Mosca del Mediterraneo, DGSPAF,
SARH, Mexico.


results of general interest for control of Anastrepha spp. (Diptera: Tephritidae).
In: Proc. CEC/IOBC Fruit Fly 'ad-hoc' meeting. XVII Int. Congress Entomol.,
ed. R. Cavalloro. Hamburg, Germany. A. A. Balkema, Rotterdam. pp. 209-215.

Celedonio-Hurtado et al.: Demography of Anastrepha

AND P. LIEDO. 1987a. A survey of the economically important fruit flies (Dipt-
era: Tephritidae) present in Chiapas and few other fruit growing regions in
Mexico. Florida Entomol. 70: 320-328.
AND J. HENDRICHS. 1987b. Natural host plant survey of the economically
important fruit flies (Diptera: Tephritidae) of Chiapas, Mexico. Florida Entomol.
70: 329-338.
ALUJA, M. AND P. LIEDO. 1986. Future perspectives on integrated management of
fruit flies in Mexico. In: Pest control: operations and systems analysis in fruit fly
management. Proc. NATO advanced workshop. M. Mangel, J. R. Carey, and R.
Plant. eds. Springer Verlag. New York. pp 12-48.
BAKER, A. C., W. E. STONE, C. C. PLUMMER, AND M. MCPHAIL. 1944. A review
of studies of the Mexican fruit fly and related mexican species U.S.D.A. Misc.
Publ. No. 531. Washington D. C.
BATEMAN, M. A. 1972. Ecology of fruit flies. Ann. Rev. Entomol. 17: 493-518.
BONNER, J. T. 1965. Size and Cycle. Princeton Univ. Press. Princeton, N.J.
CAREY, J. R. 1982. Demography and population dynamics of the Mediterranean fruit
fly. Ecol. Model. 16: 125-150.
CAREY, J. R. 1984. Host-specific demographic studies of the Mediterranean fruit fly
Ceratitis capitata. Ecol. Entomol. 9: 261-270.
CAREY, J. R. AND R. I. VARGAS. 1985. Demographic analysis of insect mass rearing:
A case study of three tephritids. J. Econ. Entomol. 78: 523-527.
CAREY, J. R., E. J. HARRIS, AND D. 0. MCINNIS. 1985. Demography of a native
strain of the melon fly, Dacus cucurbitae, from Hawaii. Entomol. Exp. Appl.
38: 195-199.
CHRISTENSON, L. D. AND R. H. FOOTE. 1960. Biology of fruit flies. Ann. Rev.
Entomol. 5: 171-192.
FITT, G. P. 1986. The roles of adult and larval specializations in limiting the occurrence
of five species of Dacus (Diptera: Tephritidae). Oecologia (Berl.) 69: 101-109.
FLETCHER, B. S. 1987. The biology of Dacine fruit flies. Ann. Rev. Entomol. 32:
FOOTE, D. S. AND J. R. CAREY. 1987. Comparative demography of a laboratory and
a wild strain of the oriental fruit fly Dacus dorsalis. Entomol. Exp. Appl. 44:
KRAINACKER, D. A. 1986. Demography of the Mediterranean Fruit Fly: Larval Host
Effects. M.S. Thesis, University of California, Davis. 360 pp.
MALAVASI, A., J. S. MORGANTE, AND R. A. ZUCCHI. 1980. Biologia de "moscas-das-
frutas" (Diptera: Tephritidae). I. Lista de Hospedeiros e ocorrencia. Rev. Brasil.
Biol. 40(1): 9-16.
NORRBOM, A. L. 1985. Phylogenetic analysis and taxonomy of the Cryptostrepha,
daciformis, robusta, and shawsi species groups of Anastrephar Schiner (Diptera:
Tephritidae). Ph.D. Dissertation, The Pennsylvania State Univ., Univ. Park.
314 pp.
PEREZ, A. 1987. Tasas de supervivencia y reproduccion de Anastrepha ludens (Loew)
en diferentes hospedantes. Tesis M. C. Centro de Entomologia y Acarologia,
Colegio de Postgraduados, Chapingo, Mexico. 89 pp.
SARH. 1983. Anuario estadistico de la production agricola. Direccion General de
Economia Agricola. Mexico D. F., Mexico.
STONE, A. 1942. The fruit flies of the genus Anastrepha. U.S.D.A. Misc. Publ. No.
439. Washington, D.C.
VARGAS, R. I. AND T. NISHIDA. 1985. Life history and demographic parameters of
Dacus latifrons (Diptera: Tephritidae). J. Econ. Entomol. 78: 1242-1244.
VARGAS, R. I., D. MIYASHITA, AND T. NISHIDA. 1984. Life history and demographic
parameters of three laboratory-reared tephritids (Diptera: Tephritidae). Ann.
Entomol. Soc. Am. 77: 651-656.
WASBAUER, M. S. 1972. An annotated host catalog of the fruit flies of America north
of Mexico (Diptera: Tephritidae). Occ. Pap. No. 19. California Dept. of Agr.
Sacramento, CA, U.S.A.

120 Florida Entomologist 71(2) June, 1988

YOUNG, A. M. 1982. Population Biology of Tropical Insects. Plenum Press, New York.
ZWOLFER, H. 1983. Life systems and strategies of resource exploitation in tephritids.
In: Proc. CEC/IOBC Int. Symp. on Fruit Flies of Economic Importance. Athens,
Greece, 1982. R. Cavalloro ed. A. A. Balkema, Rotterdam. pp. 16-30.


Project Leader, Agricultural Products Department
Dow Chemical U.S.A., 5100 West Kennedy Blvd., Suite 450
Tampa, FL 33609 USA
Research Associate, Florida State Collection of Arthropods
Florida Department of Agriculture & Consumer Services
Gainesville, FL 32602 USA


Acrosternum (Chinavia) gregi n. sp. from Ecuador is described and figured. Key
couplets to supplement Rolston's 1983 key are provided to allow recognition of this new


Se describe y figure del Ecuador a Acrosternum (Chinavia) gregi n. sp. Coplas de
claves para suplementar las claves de 1983 de Rolston se proven para permitir reco-
nocer a esta nueva especie.

Rolston (1983) revised the species of Acrosternum (Chinavia) occurring in the West-
ern Hemisphere. Rider and Rolston (1986) added three species from Mexico to this
genus and provided a key to species occurring in Mexico. Rider (1987) described an
additional species from Cuba and keyed the species occurring in the West Indies. In
1986, I collected a series of an undescribed species in Ecuador. This paper describes
and keys the new species. All measurements are given in millimeters.

Acrosternum (Chinavia) gregi n. sp.

Dorsum medium green, densely punctate, with narrow yellow to orange border
laterally on head, pronotum, coria basally and abdomen; venter light green. Body length
Head 2.1-2.6 long, 2.8-3.3 wide across eyes. Antennae green, segments 3-5 with
apical 1/3-1/2 dark brown to black, lengths of segments 1-5: 0.5-0.6, 1.1-1.4, 1.4-1.7,

Eger: New Ecuadorian Pentatomid 121

1.9-2.2, 2.0-2.2. Rostrum green, apex and median line dark brown to black, reaching
posterior margin of metacoxae, lengths of segments 2-4: 1.6-1.9, 1.0-1.3, 1.0-1.2.
Pronotum 7.0-8.6 wide, 4.8-5.9 long at meson, humeri rounded to broadly angulate,
not produced. Cicatrices immaculate. Legs green. Each ostiolar ruga extending 0.7-0.8
distance from inner margin of ostiole to lateral margin of thorax.
Scutellum 4.8-5.9 long at meson, 4.2-5.4 wide at base, unicolorous. Costal angle of
each corium rounded apically, extending at most onto anterior 1/3 of last connexival
segment. Connexiva with small black spot present at posterolateral angles of segments,
extending onto laterotergite. Abdominal tubercle compressed, produced anteriorly to
about middle of metacoxae. Spiracles pale brown to brown, not located in or near pale
macula or callus.
Basal plates with middle half of posterior margin concave, lateral half of posterior
margin variable, from smoothly convex to strongly produced into posterolateral projec-
tion; projection or convexity not or slightly reflexed toward ninth paratergite (Figs. 8,
9). Mesial margins of basal plates slightly convex to straight. Spermathecal bulb and
pump as in Fig. 12.

d, r


5 I

Figs. 1-7. 1-3. A. gregi. 1. Pygophore, caudal view; posterior wall (pw). 2. Genital
cup, dorsal view; dorsal rim of posterior wall (dr). 3. Pygophore, ventral view. 4-5. A.
marginatum. 4. Pygophore, caudal view. 5. Pygophore, ventral view. 6. A. hilare.
Pygophore, ventral view. 7. A. gregi. Paramere. Dimensional lines equal 1.0 mm.

Florida Entomologist 71(2)

June, 1988

Posterior margin of genital cup with broad V-shaped emargination in ventral view,
distinctly emarginate laterally (Fig. 3); posterior wall with sinuous V-shaped emargina-
tion in caudal view, lateral emargination relatively deep (Fig. 1). Dorsal rim of posterior
wall of genital cup diagonal, mesial margins somewhat denticulate, concave, concavity
bordered anteriorly and posteriorly by distinct tooth (Fig. 2). Paramere as in Fig. 7.
Distribution: Known only from Ecuador.
Holotype: Male, labelled: (a) ECUADOR: Pichincha Prov., Tinalandia; 12 km. E. Sto.
Domingo de los Colorados. ca. 2500 ft., 11-17-V-1986. J. E. Eger, coll. (b) Acrosternum
n. sp., Det. L. H. Rolston '87. Deposited in the Florida State Collection of Arthropods,
Gainesville, Florida.
Paratypes: 1 male, 8 females: 1 male, 7 females labelled same as holotype except lacking
determination label; 1 female labelled: ECUADOR: Pichincha Prov., Tinalandia, 2800'
el., 12 km. E. of Santo Domingo. June 28-30, 1980. Coll. Dan Bogar. Paratypes depo-
sited as follows: Florida State Collection of Arthropods (2 females); L. H. Rolston

* 1



Figs. 8-13. 8-9. A. gregi. Variation in genital plates, ventral view; basal plates (bp),
9th paratergite (pt9). 10. A. marginatum. Genital plates, ventral view. 11. A. hilare.
Genital plates, ventral view. 12-13. A. gregi. 12. Spermathecal bulb and pump. 13.
Anomalous spermathecal bulb. Dimensional lines equal 1.0 mm.

Eger: New Ecuadorian Pentatomid 123

collection, Louisiana State University, Baton Rouge, Louisiana (1 female); American
Museum of Natural History, New York City, New York (1 female); National Museum
of Natural History, Washington, D. C. (1 male, 1 female); British Museum of Natural
History, London, England (1 female); Texas A&M University Collection, College Sta-
tion, Texas (1 female); author's collection (1 female).
Comments: The structure of the dorsal rim of the posterior wall of the genital cup in
A. gregi is similar to that of A. triangulum Rider and Rolston which occurs in Southern
Mexico, but A. gregi can be readily separated from A. triangulum by the lack of yellow
maculae on the base of the scutellum and by the lack of crimson coloration ventrally on
the rostrum. In Ecuador, this species is most likely to be confused with Acrosternum
marginatum (Palisot de Beauvois). The following key couplets may be inserted into the
key provided by Rolston (1983) to allow recognition of A. gregi among the species of
Acrosternum in South America:

42(40). Costal angle of each corium subacute, reaching beyond middle of
last connexival segment ............................. fuscopunctatum (Breddin)
Costal angle of each corium rounded, not surpassing anterior 1/3 of
last connexival segment ......................................... ........... .. 43
43(42). Females: posterior margin of basal plates evenly convex (Fig. 11);
males: emargination of posterior margin of phygophore shallow, with
small median notch from ventral view (Fig. 6) ................... hilare (Say)
Females: posterior margin of basal plates concave mesially, convex
laterally or with posterolateral projection at base of 9th paratergites
(Figs. 8, 9, 10); males: posterior margin of pygophore with broad
V-shaped emargination from ventral view (Figs. 3, 5) ................... 43.1
43.1(43). Females: posterolateral projection of basal plates present or absent
(Figs. 8, 9), but if present, projection only slightly reflexed toward
9th paratergites; males: pygophore with deep lateral emargination in
ventral and caudal views (Figs. 1, 3) ................................ gregi n. sp.
Females: posterolateral projection of basal plates present (Fig. 10)
and strongly reflexed toward 9th paratergites; males: pygophore en-
tire laterally in ventral view (Figs. 4, 5) ..................................
... ................................................ marginatum (Palisot de Beauvois)

The spermathecal bulb of one female examined had a third projection (Fig. 13). This
apparently anomalous condition was also observed by Rider and Rolston (1986) in A.
dubium Rider and Rolston. This species is dedicated to the memory of my younger
brother, Greg.


I am grateful to Dr. L. H. Rolston, Louisiana State University, Baton Rouge,
Louisiana, who determined that this was an undescribed species and provided comments
on the manuscript.


RIDER, D. A. 1987. A new species of Acrosternum Fieber, subgenus Chinavia Orian,
from Cuba (Hemiptera: Pentatomidae). J. New York Entomol. Soc. 95: 298-301.
RIDER, D. A. AND L. H. ROLSTON. 1986. Three new species of Acrosternum Fieber,
subgenus Chinavia Orian, from Mexico (Hemiptera: Pentatomidae). J. New York
Entomol. Soc. 94: 416-423.

124 Florida Entomologist 71(2) June, 1988

ROLSTON L. H. 1983. A revision of the genus Acrosternum Fieber, subgenus Chinavia
Orian, in the Western Hemisphere (Hemiptera: Pentatomidae). J. New York
Entomol. Soc. 91: 97-176.


Department of Zoology
Michigan State University
East Lansing, Michigan 48824 USA


A new species, Denisiella lithophila Snider, is described from Georgia. It appears
to be more closely related to Denisiella serroseta (Bmrner), a South African species,
than Denisiella sexpinnata (Denis 1931), a species from Costa Rica and possibly North
America. Separation of D. lithophila from D. serroseta may be accomplished by pres-
ence of a color pattern, configuration of female antenna and unguicular filament length.
Separation from sexpinnata is by lack of serrate setae on the anal papilla. The type
locality is Decatur County, Georgia, water surface on granite outcrop.


Se describe de Georgia la nueva especie Denisiella lithophila Snider. Esta especie
parece estar relacionada mAs cercamente a Denisiella serroseta (B6rner), que es una
especie de Africa del Sur, que a Denisiella sexpinnata (Denis 1931), que es una especie
de Costa Rica y posiblemente de NorteamBrica. Se puede separar a D. lithophila de D.
serroseta por el patron de color, configuraci6n de la antena de la hembra, y el largo del
filamento unguicular. Se separa de sexpinnata por la falta de setas serradas en la papila
anal. La localidad tipo estA en el granite fuera de la superficie del agua en el Condado
de Decatur, Georgia.

This paper is another in a series describing sminthurid Collembola from the south-
eastern United States. While working at the Department of Entomology, University
of Georgia at Athens, I conducted a number of collecting trips during 1980-1981 in
Georgia. Many new records and species have been found. Here my purpose is to describe
a new species from collections made on Mount Arabia, a granite outcrop. The classifica-
tion used for higher categories follows Betsch (1980).

Denisiella lithophila, NEW SPECIES

COLOR & PATTERN: Dark blue-black with cream markings laid down in irregular
mosaics. Female: Head dark blue becoming lighter dorsally, with cream mosaics on
vertex and surrounding eyepatches; antenna bluish-purple with cream spots basally on
each of first three segments. Abdomen dark blue-black becoming lighter dorsally, with

Snider: New Collembola from Georgia

Denisiella lithophila n. sp.
Fig. 1. Habitus, dorsal aspect, female; 2. Habitus, lateral aspect, female; 3. Habitus,
dorsal aspect, male; 4. Habitus, lateral aspect, male.

irregular blue mosaics forming a mid-dorsal line; anal papilla dorsally with two cream
maculae. Legs dark blue with cream markings basally on each segment. Furcula light
blue; parafurcular lobes white (Figs 1 & 2). Male: Cream with blue maculae. Head light
blue with dorsal cream mosaics, a black spot between antennal bases; antenna violet-
purple, becoming darker distally. Abdomen cream with blue pigment ventrally and
dorsally, creating a light irregular, lateral band; dorsally a dark longitudinal line with
irregular lines radiating off at right angles. Anal papilla light blue with two dorsal
maculae. Legs blue, becoming darker distally. Furcula light blue; parafurcular lobes
light blue, cream-white ventrally (Figs. 3 & 4).
HEAD: Eyes 6 + 6; ocelli subequal (Fig. 5). Mandible with large molar plate, elongate
apex (Fig. 6). Maxilla with 4 galeal teeth (Fig. 7). Outer maxillary lobe with 4 distal


Florida Entomologist 71(2)


58 9

4 15 16
\' -"- 1--5

Denisiella lithophila n. sp.
Fig. 5. ocelli, left side of head; 6. mandible; 7. maxilla; 8. outer maxillary lobe; 9.
labrum; 10. antenna, female; 11. antenna, male; 12. foreleg, coxa; 13. foreleg, trochanter;
14. foreleg, femur; 15. foreleg, tibiotarsus, anterior surface; 16. foreleg, tibiotarsus,
posterior surface; 17 foreclaw.

setae (Fig. 8). Labrum with stout setae (Fig. 9). Mean antennal ratio of female 1:1:1:2,
male 3:3:1.5:2; female ANT IV with subapical papilla and numerous setae; ANT III with
sensilla in separate pockets; ANT II with 8-9 dorsal setae; ANT I with 4 setae (Fig.
10); male ANT IV with subapical papilla, shape subovate; ANT III with large apical
projection and various clasping setae (C1 truncate); ANT II with b6 greatly enlarged,
b, reduced; ANT I simple (Fig. 11). FORELEG: Coxa with simple seta (Fig. 12);

June, 1988

Snider: New Collembola from Georgia

31 32 20

Denisiella lithophila n. sp.
Fig. 18. mesoleg, coxa; 19. mesoleg, trochanter; 20. mesoleg, femur; 2. mesoleg,
tibiotarsus, anterior surface; 22. mesoleg, tibiotarsus, anterior surface; 23. mesoclaw;
24. metaleg, coxa; 25. metaleg, trochanter; 26. metaleg, femur; 27. metaleg, tibiotarsus,
anterior surface; 28. metaleg, tibiotarsus, posterior surface; 29. metaclaw; 30. manub-
rium, dorsum; 30. dens, dorsal surface; 32. dens, ventral surface; 33. mucro, female; 34.
mucro, male; 35. collophore; 36. retinaculum.

128 Florida Entomologist 71(2) June, 1988

trochanter with 2 anterior and 1 posterior setae (Fig. 13); femur with 5 anterior and 4
posterior setae (Fig. 14); tibiotarsus with 5E setae on anterior surface (Fig. 15), poste-
rior surface with 4L setae (Fig. 16); pretarsus with 1+1 setulae, unguis with single
inner and outer teeth, curving, lanceolate; unguiculus with subapical filament reaching
length of unguis, apex weakly swollen (Fig. 17). MESOLEG: Coxa with single anterior
seta and short spine (Fig. 18); trochanter with 2 anterior and 1 posterior setae (Fig.
19); femur with 5 heavy anterior and 6 posterior setae (Fig. 20); tibiotarsus with 4E
setae on anterior surface (Fig. 21), posterior surface with 5L setae (Fig. 22); pretarsus
with 1+ 1 setulae; unguis with single inner and outer teeth, curving, lanceolate; un-
guiculus with subapical filament reaching length of unguis, apex weakly swollen (Fig.
23). METALEG: Coxa with 3 anterior setae (Fig. 24); trochanter with 2 anterior setae
and 1 posterior setulae (Fig. 25); femur with 6 anterior and 2 posterior setae (Fig. 26);
tibiotarsus with 5E setae on anterior surface (Fig. 27), posterior surface with 5L setae,
3 of which are stout and coursely serrate (Fig. 28); pretarsus with 1+1 setulae; unguis
with single inner and outer teeth, curving, lanceolate; unguiculus with subapical fila-
ment reaching length of unguis, apex slightly swollen (Fig. 29). FURCULA: Dorsum of
manubrium with 7+7 setae (Fig. 30); dorsal surface of dens with numerous, long setae,
female with 6 long L setae (Fig. 31), male has bases of L setae greatly expanded; ventral
surface of dens typical of other members of the genus (Fig. 32); mucro of female with
outer lamella smooth, inner surface finely toothed, outer seta present (Fig. 33), male
similar, except both lamellar surfaces smooth (Fig. 34). GREAT ABDOMEN: Collophore
with 1 + 1 subapical setae (Fig. 35); corpus of retinaculum with 2 distal setae, rami with
4 teeth (Fig. 36). Body of female with short curving setae (Fig. 37), male with longer,
stiff setae (Fig. 38). LESSER ABDOMEN: Female anal papilla with gladiform dorsal
setae (Fig. 39), male similar, but reduced in size (Fig. 40). Length: Female 0.75mm,
male 0.50mm.


This new species keys out in Christiansen & Bellinger (1981) to Sminthurides (De-
nisiella) sp., based on the absence of the tibiotarsal organ and presence of modified
setae on the metatibiotarsus, as well as a mucronal seta. The only species so far recorded
from North America is Denisiella sexpinnatus Denis (1931), originally described from
Costa Rica. Christiansen & Bellinger point out that their records from the United

37 38 39

Denisiella lithophila n. sp.
Fig. 37. great abdomen, bothriotrix A,B,C, female; 38. great abdomen, bothriotrix
A,B,C, male; 39. anal papilla, female; 40. anal papilla, male.

Snider: New Collembola from Georgia

States may represent a still different species. Based on descriptions in the literature,
Denisiella lithophila n. sp. may easily be separated from D. sexpinnata on the basis of
color and pattern and the following morphological characteristics:

D. sexpinnatus D. lithophila
dens with 4VeA, 5VeC setae 4VeA, 6VeC setae
metatibiotarsus with 5, strong, toothed spines with 3 strong, toothed spines
anal papilla with 3 + 3 strongly toothed spines spines smooth

In Folsom & Mills (1938), D. lithophila keys out closest to Denisiella serroseta (Bbrner),
(1908). It differs from that South African species by having a cream and dark blue color
pattern in the female; lacking a swollen base on ANT II; and having an unguicular
subapical filament that reaches the apex of the unguis. These two species are remarka-
bly similar in that they lack crenulate or serrate setae on the anal papilla; share the
same mucronal configuration in both male and female; and in the male, lack an inner
tooth on the metaunguis. D. lithophila does have a weak tooth on the fore- and meso-
claws of the male. In their description of Denisiella, Folsom & Mills (1938) mention the
presence of 4 swellings at the bases of the male fore tibiotarsi; D. lithophila does not
exhibit this morphological characteristic.
TYPES: Holotype 9 and allotype S in 95% ethanol, 100 + 9 9 and 25 d~ paratypes in
95% ethanol, and 21 dissection slides representing 10 specimens deposited in the En-
tomology Museum, Michigan State University, East Lansing, Michigan. Collection data:
Georgia, Decatur County, Mount Arabia, water filled depression, granite outcrop, "May
23, 1982", R. J. Snider, collector.


Special thanks are extended to Peter Carrington, graphic artist, Department of
Entomology, Michigan State University, for his renderings of the male and female
habitus sketches; to Dr. D. A. Crossley, Jr. for his help in locating the unique collection
site, and the use of his laboratory facilities; and to Dr. Kenneth A. Christiansen, Grinnell
College and Dr. Peter F. Bellinger, California State University, Northridge for their
review of the manuscript.


BETSCH, J.-M. 1980. Elements pour une monographie des Collemboles SymphyplBones
(Hexapodes, Apterygotes). Mem. Mus. Nat. Hist. Nat., Serie A, 116: 1-227.
BORNER, C. 1908. Collembolen aus Sudafrica nebst einer Studie uber die I. Maxille
der Collembolen. Schultz, Forschungsreise. Denkschr. Med.-Naturw. Ges. Jena.
13: 53-69.
CHRISTIANSEN, K. A. AND P. F. BELLINGER. 1981. The Collembola of North
America north of the Rio Grande, Part IV. Families Neelidae and Sminthuridae.
Grinnell College, Grinnell, Iowa: 1043-1322.
DENIS, J. R. 1931. Contribute alla conoscenza del "Microgenton" di Costa Rica. II.
Collemboles de Costa Rica avec une contribution au species de l'ordre. Bol. Lab.
Zool. Gen. Agr. R. Inst. Super. Agr. Portici, 25: 69-171.
FOLSOM, J. W. AND H. B. MILLS. 1938. Contribution to the knowledge of the genus
Sminthurides Borner. Bull. Mus. Comp. Zool. Harvard, 82: 231-274.

130 Florida Entomologist 71(2) June, 1988


Museo de Insectos, Facultad de Agronomfa
Universidad de Costa Rica, San Jose, Costa Rica

USDA/ARS/Systematic Entomology Laboratory
c/o National Museum of Natural History, NHB 168
Washington, D.C. 20560 USA


Distribution and host plant data for 28 species of Anastrepha known to occur in
Costa Rica are given. These data include the results of an extensive trapping survey
and fruit sampling conducted in 1985-86, a study of museum specimens, and a summary
of information from previous publications. Ten species are reported from Costa Rica for
the first time: A. antunesi Lima, A. bahiensis Lima, A. barnesi Aldrich, A. concava
Greene, A. crebra Stone, A. hamata (Loew), A. robusta Greene, A. tumida Stone, and
two undetermined, probably undescribed species. Variation in host use and adult mor-
phology between Costa Rican and Mexican-Guatemalan populations of A. ludens (Loew)
is discussed.


Se ofrece informaci6n sobre las distribuciones y las plants hospederas en Costa Rica
de 28 species conocidas de moscas de las frutas del g6nero Anastrepha. Se incluyen
los resultados de un extenso muestreo de moscas de este genero por medio de trampas
y por cultivo de frutas infestadas durante 1985-86. A estos resultados se agrega la
information acumulada en publicaciones previas; asi como la de especimenes de museos
recolectados en distintas 4pocas. En este trabajo se informan diez nuevos registros de
species para Costa Rica: A. antunesi Lima, A. bahiensis Lima, A. barnesi Aldrich,
A. concava Greene, A. crebra Stone, A. hamata (Loew), A. robusta Greene, A. tumida
Stone, y dos species no identificadas que probablemente no son descritas. Se discute
variaci6n en hospederas y morfologia entire poblaciones de Costa Rica y de M6xico y
Guatemala de A. ludens (Loew).

Anastrepha Schiner is the most economically important genus of fruit flies in Costa
Rica (Jir6n and Zeled6n 1979, Jir6n and Hedstrdm 1988). In 1985 a project on the
ecology of these flies was begun at the Universidad de Costa Rica, an important part
of which is to determine such basic data as which species of Anastrepha occur in Costa
Rica, what are their distributions within the country, and what host plants do they
infest. In this paper we report the results to date.
Nineteen species of Anastrepha were previously recorded from Costa Rica. The
study of the genus in this country began when Picado (1920) published a paper on A.
striata Schiner associated with guava (P. guajava L.). Salas (1957), in his study on the
Mediterranean fruit fly, Ceratitis capitata (Wiedemann), also determined the host pref-
erences of several Anastrepha species. Jir6n and Zeled6n (1979) determined that A.
striata, A. obliqua (Macquart), and A. serpentina (Wiedemann) are the three most

_ ____.


Jir6n et al.: Costa Rica Anastrepha

economically important fruit fly species in Costa Rica, causing the greatest damage to
locally consumed fruits. Records for additional Anastrepha species have been reported
in more comprehensive taxonomic studies (Stone 1942, Foote 1967) and in various recent
papers (Saunders 1978, Hedstrim et al. 1985, Soto-Manitiu and Jir6n 1988, Gonzalez et
al. 1988).
Here, we summarize this information, report ten new species records for Costa
Rica, and include additional biological data for the previously known species.


Since 1985 L. F. Jir6n, J. Soto-Manitiu, and co-workers at the Facultad de Ag-
ronomia, Universidad de Costa Rica have conducted an extensive survey for Anas-
trepha in Costa Rica. We have reared 4,126 specimens from 440 samples of 18 fruit
species from 198 localities throughout the country. We also collected 11,139 specimens
in 95 McPhail traps, baited with torula yeast, placed in seven localities: Finca La Selva,
Sarapiqui, Prov. Heredia; Centro Ecol6gico La Pacifica, Cafias, Prov. Guanacaste;
Ciudad Col6n, Prov. San Jos6; Orotina, Prov. Alajuela; Lepanto, Prov. Puntarenas; and
Buenos Aires, Prov. Puntarenas.
Unless otherwise noted, specimens collected in our survey are deposited in the
Museo de Insectos, Universidad de Costa Rica. Additional specimens were examined
from the following collections: Academy of Natural Sciences, Philadelphia (ANSP); Na-
tional Museum of Natural History, Smithsonian Institution, Washington, D.C. (USNM);
California Academy of Sciences, San Francisco (CAS); Los Angeles County Museum
(LACM); Museum of Comparative Zoology, Cambridge, Massachussetts (MCZ); Texas
A & M University, (TAMU); University of California, Berkeley (UCB); University of
Wisconsin, Madison (UWM); and Utah State University, Logan (USU).
Norrbom (1985) listed most of the reported host plants for Anastrepha species. Host
data in this paper are drawn from that list unless otherwise noted. The botanical
nomenclature follows Terrell et al. (1986), that of the flies, Stone (1942) and Steyskal


1. A. antunesi Lima
Distribution. Previously known from Panama, Peru, Venezuela, Trinidad and Brazil
(Stone 1942, Caraballo 1981). 1 9 (LACM) was collected at Boca del Rio Barranca, Prov.
Puntarenas, 19-VI-1963 by C. L. Hogue.
Host plants. Genipa americana L. (Rubiaceae), Spondias mombin L., S. purpurea
L. (Anacardiaceae), Dovyalis hebecarpa (Gardner) Warb. (Flacourtiaceae), and Manil-
kara zapota (L.) P. Royen (Sapotaceae). All the reported hosts are present in Costa
Rica, and we have sampled Spondias and Manilkara extensively, but have not yet
found this species in infested fruit.

2. A. bahiensis Lima
Distribution. Previously known from Panama and Brazil (Foote 1967). We tenta-
tively use this name for 1 9 trapped on 25-II-1986 at Buenos Aires. It appears to be
conspecific with specimens from Panama determined by Stone (1942) as A. bahiensis,
although whether or not these are conspecific with populations in Brazil is unresolved.
At least the specimens reared from Helicostylis in Brazil aculeuss tip figured by Stone
1942) may be a different species.
Host plants. Juglans neotropica Diels, J. regia L. (Juglandaceae), Helicostylis poep-
pigiana Tree. (Moraceae), and Eugenia variabilis Baillon (Myrtaceae).

Florida Entomologist 71(2)

June, 1988

3. A. balloui Stone
Distribution. Costa Rica, Panama and Venezuela (Foote 1967, Soto-Manitiu and
Jir6n 1988). We trapped this species only during the dry season (Feb.-May) at Lepanto
and Orotina.
Host plants. Terminalia catappa L. (Combretaceae) and Sterculia apetala (Jacq.)
Karsten (Sterculiaceae). Soto-Manitiu and Jir6n (1988) trapped 21 adults in commercial
mango orchards, but did not recover this species from mango fruits.

4. A. barnesi Aldrich
Distribution. Previously known from Guatemala, Panama, Guyana, and Brazil
(Foote 1967). 1 d (ANSP) was collected by H. R. and E. H. Roberts south of Rinc6n,
Osa Peninsula, Prov. Puntarenas, 7-20-II-1967.
Host plants. Pouteria torta Radlk. (Sapotaceae). Pouteria spp. are widespread in
Costa Rica.

5. A. canalis Stone
Distribution. Costa Rica, Panama, and Venezuela (Soto-Manitiu and Jir6n 1988,
Stone 1942). We have seen 1 9 (USU) collected 6 km south of San Vito, Prov. Pun-
tarenas, 2-V-1967, by D. F. Veirs, 1 9 (UCB) collected 9 mi. south of Turrialba, Prov.
Cartago, 8-VIII-1965, by A. Raske and C. Slobodchikoff, and 2 9 we trapped at Orotina.
Host plants. Turpinia occidentalis (Swartz) Don. (Staphylaceae). The females from
Orotina were trapped in a mango orchard, but we have not recovered A. canalis from
mango fruit.

6. A. chiclayae Greene
Distribution. U.S.A. (Texas), Mexico, Costa Rica, Panama, Venezuela, Peru, and
Argentina (Stone 1942, Caraballo 1981, Gonzalez et al. 1988). We have seen specimens
from Estaci6n Experimental Fabio Baudrit, Prov. Alajuela, Guicimo, Prov. Lim6n,
Finca La Selva, and Buenos Aires. All these localities are in the tropical wet forest zone.
Host plants. Anacardium occidentale L., Mangifera indica L., Spondias purpurea
L. (Anacardiaceae), Psidium guajava L. (Myrtaceae), Passiflora sp., P. quadran-
gularis L. (Passifloraceae), Prunus persica (L.) Batsch, Cydonia oblonga Miller
Rosaoeae), and Citrus spp. Rutaceae). Except for Passiflora, we question whether the
above species are natural hosts for A. chiclayae, at least in Costa Rica. We did not
recover it from mango, mombin, guava, or peach fruit despite extensive sampling.

7. A. concava Greene
Distribution. Previously known from Panama, Ecuador, and Brazil (Foote 1967). 1
Y (ANSP) was collected at Rinc6n, Osa Peninsula, 7-20-II-1967 by H. R. and E. H.
Host plants. Nothing is known about the biology of this species.

8. A. crebra Stone
Distribution. Previously known from Panama and Honduras (Stone 1942). 1 9 (CAS)
was collected at Turrialba, Prov. Cartago, 14-VII-1965 by H. G. Real.
Host plants. Quararibea asterolepis Pittier (Bombacaceae).

9. A. distinct Greene
Distribution. U.S.A. (Texas) (not currently established), Mexico, Guatemala, Costa
Rica, Panama, Colombia, Ecuador, Peru, Guyana, Venezuela, Trinidad, and Brazil
(Stone 1942, Caraballo 1981). We collected it from many localities in the tropical wet
forest zone (Jir6n and Hedstrom 1988).
Host plants. Thirteen species of Inga (Fabaceae) and 11 other species in eight plant
families. Mango is a reported host in Peru, but we have not found this fruit infested by

Jir6n et al.: Costa Rica Anastrepha

A. distinct in Costa Rica. We reared it from fruit from wild and cultivated trees of
Inga edulis Mart. (Jir6n and Hedstr6m 1988).

10. A. fraterculus (Wiedemann)
Distribution. Throughout continental America from U.S.A. (Texas) (not currently
established) and Mexico to Argentina and Chile, Trinidad, Tobago (Stone 1942, Foote
1967). We trapped or collected it at Santiago de San Ram6n, Prov. Alajuela, Cafias,
Finca La Selva, and Buenos Aires.
Host plants. 68 species in 23 plant families. We reared it only from guava (Psidium
guajava L.), and it does not appear to be a species of great economic significance in
Costa Rica (Jir6n and Zeled6n 1979, Jir6n and Hedstrbm 1988).

11. A. hamata (Loew)
What we are calling A. hamata may actually be a complex of sibling species.
Distribution. Previously reported from Panama and Brazil (Foote 1967). 1 9 (ANSP)
was collected by H. R. and E. H. Roberts 3-10 mi. south of Rinc6n, Osa Peninsula,
Prov. Puntarenas, 7-20-II-1967, and we collected 1 9 at Estaci6n Experimental Fabio
Baudrit, Prov. Alajuela on 24-IX-1987.
Host plants. No hosts have been recorded, but A. sagittata (Stone), which is closely
related or perhaps even conspecific with A. hamata, has been reared from Pouteria
campechiana (Kunth) Baehni (Sapotaceae).

12. A. irretita Stone
Distribution. Costa Rica and Panama (Foote 1967, Soto-Manitiu and Jir6n 1988). We
trapped 1 9 at Centro Ecol6gico La Pacifica.
Host plants. No hosts known.

13. A. leptozona Hendel
Distribution. Mexico, Guatemala, Costa Rica, Panama, Venezuela, Bolivia, Guyana,
and Brazil (Foote 1967, Aluja and Martinez 1985, Soto-Manitiu and Jir6n 1988). We
collected 1 specimen at Golfito, Prov. Puntarenas.
Host plants. Chrysophyllum cainito L., Pouteria caimito (R. & P.) Raldk., P.
campechiana (Kunth) Baehni, Micropholis mexicana Gilly, Lucuma (s. lat.) sp.
(Sapotaceae) and Cratequs sp. (Rosacea) (Aluja et al. 1987, Norrbom 1985).

14. A. limae Stone
Distribution. U.S.A. (Texas), Costa Rica, Panama and Venezuela (Foote 1967,
Caraballo 1981, Hedstr6m et al. 1985). We collected 10 adults from Estaci6n Experimen-
tal Fabio Baudrit, Prov. Alajuela, Guicimo, Prov. Lim6n, and Finca La Selva.
Host plants. Passiflora quadrangularis L. We collected 3 adults on fruit of a Passif-
lora species (Hedstrom et al. 1985) and reared 5 specimens from fruit of a Passiflora

15. A. ludens (Loew)
Distribution. U.S.A. (Texas), Mexico, Guatemala, El Salvador, Honduras,
Nicaragua, and Costa Rica (Stone 1942; unpublished data). We examined 1 S and 2 9
(USNM) reared in San Jos6 by Ballou in 1936 (Stone 1942) and 1 S and 3 9 (TAMU,
USNM) reared by Fischel & Gilstrap at Santa Rosa, Prov. Heredia, 2-VI-1980.
Host plants. A. ludens has been reared from at least 49 plant species belonging to
18 families, although many are doubtfully natural hosts in the field. Mango and Citrus
are heavily infested in Mexico (Aluja and Martinez 1985) and Guatemala (Eskafi and
Cunnigham 1987), but we have not recovered A. ludens from samples of fruit from these
plants or during our extensive trapping (more than 15,000 specimens total). All of the
specimens from Costa Rica that we have examined were reared from fruit of Casimiroa

134 Florida Entomologist 71(2) June, 1988

edulis Llave & Lex. (Rutaceae). This plant is not native to Costa Rica, but C. tetrameria
Millsp., which is also a reported host of A. ludens, occurs from southern Mexico to
Costa Rica (Standley 1937).
The reason that mango, citrus, and other commercial hosts do not appear to be
attacked in Costa Rica should be further investigated. Climatic factors may be involved,
or perhaps competition from other species, especially A. obliqua on mango, but the
difference in host usage could also be due to genetic differences between populations of
A. ludens. The Costa Rican females examined differ slightly from Mexican and
Guatemalan females in having the aculeus (ovipositor) tip very weakly or non-serrate
(only one specimen from Santa Rosa had even a few weak serrations), but this may be
clinal variation, because several females from Nicaragua that we examined also have
the aculeus tip weakly serrate. Additional material from Central America is needed to
further study this variation, but the existence of Central American A. ludens popula-
tions that appear to vary morphologically and in host usage from Mexican populations
casts doubt on the hypothesis of Baker et al. (1944), that A. ludens occurred only in
northeastern Mexico prior to the 1800's. The difference in the aculeus also has important
implications for species concepts within Anastrepha and for the identification of these
flies, both of which are heavily based on aculeus morphology. The Costa Rican females
will not run to A. ludens in the keys of Stone (1942) or Steyskal (1977).

16. A. manihoti Lima
Distribution. Costa Rica, Panama, Venezuela, Brazil (Foote 1967, Saunders 1978).
Saunders (1978) reported it from Turrialba, Prov. Cartago, and we collected it at Buenos
Aires and Finca La Selva. All three localities are in the tropical wet forest zone.
Host plants. Manihot esculenta Crantz (= aipi Pohl, dulcis (Gmel.) Pax.) (Euphor-
biaceae). The larvae have been reported to infest both the buds and stalks and the seed
capsules (Lima 1934, Pofia and Bellotti 1977, Caraballo 1981, Saunders 1978, Jir6n and
Hedstrbm 1988). In Costa Rica, A. manihoti has been reared only from the buds and

17. A. montei Lima
Distribution. Costa Rica, Panama, Venezuela, Brazil, Paraguay, and Argentina
(Foote 1967, Aluja et al. 1987). Stone (1942) reported six specimens (USNM) from
Higuito, San Mateo, Prov. Alajuela collected by P. Schild. We have confirmed their
Host plants. Manihot esculenta Crantz (= aipi Pohl, dulcis (Gmel.) Pax.) (Euphor-
biaceae). The larvae infest the seed capsule (Stone 1942).

18. A. obliqua (Macquart)
Distribution. Greater and Lesser Antilles, U.S.A. (Texas, Florida) (not currently
established), Mexico, Central America (including Costa Rica), Panama, Venezuela,
Ecuador, and Brazil (Foote 1967). We have found it at numerous localities below 1300 m.
Host plants. At least 64 species in 24 families, although many are not confirmed
natural hosts in the field. Many of the preferred hosts belong to the family Anacar-
diaceae (Whervin 1974), and in Costa Rica A. obliqua is the major pest on mango and
commercial Spondias species (Jir6n and Hedstrbm 1988).

19. A. panamensis Greene
Distribution. Costa Rica and Panama (Foote 1967, Soto-Manitiu and Jir6n 1988). We
trapped 1 2 at Orotina.
Host plants. Chrysophyllum cainito L. and C. panamense Pittier (Sapotaceae).
Although we have reared other Anastrepha species from 38 samples of infested C.
cainito fruit from various localities, we have not yet recovered A. panamensis.

Jir6n et al.: Costa Rica Anastrepha

20. A. pickeli Lima
Distribution. Costa Rica, Panama, Peru, Venezuela, Brazil, and Argentina (Foote
1967, Hedstrim et al. 1985). We collected this species at Estaci6n Experimental Fabio
Baudrit, Prov. Alajuela and Buenos Aires.
Host plants. Manihot esculenta Crantz (= aipi Pohl, dulcis (Gmel.) Pax.) (Euphor-
biaceae), Quararibea magnifica Pittier, and Q. turbinata Poir (Bombacaceae). Jir6n
and Hedstrim (1988) reared it from seed capsules of yuca (M. esculenta).

21. A. robusta Greene
Distribution. Previously known from Sourthern Mexico, Guatemala, and Panama.
Records from Venezuela and Brazil are erroneous (Norrbom 1985). One S (USNM) was
collected by R. Holzenthal at Rio Los Ahogados, Quebrada Grande, Prov. Guanacaste,
Host plants. No hosts known.

22. A. schausi Aldrich
Distribution. Known only from the type locality, Juan Vifias, Prov. Cartago, Costa
Host plants. Nothing is known about the biology or host plants (Norrbom and Kim

23. A. serpentina (Wiedemann)
Distribution. U.S.A. (Texas), Mexico, Central America, Ecuador, Peru, Venezuela,
Guyana, Trinidad, Argentina, Brazil (Foote 1967).
Host plants. About 40 species in 13 families. Species of Sapotaceae appear to be the
primary hosts in Costa Rica, and A. serpentina is the major fruit fly pest on three
commercial crops, Manilkara zapota (L.) P. Royen, Chrysophyllum cainito L., and
Pouteria caimito (R. & P.) Radlk. Jir6n and Hedstrdm (1988) also reared it from Dios-
pyros digyna Jacq. (Ebenaceae).

24. A. spatulata Stone
Distribution. U.S.A. (Texas), Mexico, Costa Rica, Panama (Foote 1967). We trap-
ped 2 9 in a mango orchard at Centro Ecol6gico La Pacifica, 19-II-1986.
Host plants. No hosts known.

25. A. striata Schiner
Distribution. U.S.A. (Texas) (not currently established), Mexico, Guatemala, Hon-
duras, Costa Rica, Panama, Colombia, Ecuador, Peru, Bolivia, Venezuela, Surinam,
Trinidad, Brazil (FernAndez-Y6pez 1953, Foote 1967, Eskafi and Cunningham 1987).
Host plants. 23 species in 10 families, most of which have been found naturally
infested in the field (Berg 1959, Norrbom 1985, Jir6n and Hedstr6m 1988). Species of
Myrtaceae, and especially Psidium, are the primary hosts. In Costa Rica A. striata is
the major pest on common guava (P. guajava L.); over 97% of the tephritids Jir6n and
Hedstrom (1987) reared from this fruit were A. striata. Mexz6n and Jir6n (in prep.)
found that on the Caribbean slope of Costa Rica, populations are maintained throughout
the year by shifting among the following hosts: P. guajava, P. friedrichsthalianum
(Berg) Ndz., P. savanarum Donn. Sm., Persea americana Mill., and Spondias mombin

26. A. tumida Stone
Distribution. Previously known only from Panama (Foote 1967). We examined 4
specimens from La Lola, Prov. Lim6n: 1 6 and 1 9 (USNM) collected 1-VI-1961 by J.
HernAndez, 1 9 (UWM) collected 25-IV-1957 by M. J. Stelzer, and 1 & (UWM) collected
31-VII-1957 by R. D. Shenefelt.
Host plants. No hosts known.

Florida Entomologist 71(2)

27. Anastrepha undetermined sp. A
At Finca La Selva, between 29-IV and 23-V-1986, we trapped 42 adults of an unde-
termined Anastrepha species. It is probably undescribed, but further taxonomic study
is needed.

28. Anastrepha undetermined sp. B
We examined 1 9 and 2 & (MCZ), collected by P. and D. Allen at Palmar, Prov.
Puntarenas, V-1949, of a second, probably undescribed species also represented by a
large series (USNM) from Arraijan, Panama.


As a result of our survey and study of museum specimens, the number of species of
Anastrepha known from Costa Rica has been increased from 11 to 28 (including the two
unnamed species). Considering the number of species recorded for Panama (more than
60), it is likely that the acutal number is much higher, and we expect it to increase as
our project continues. Certainly, species such as A. acris Stone, A. alveata Stone, and
A. cordata Aldrich, known from Panama and from Mexico or other parts of Central
America (Stone 1942; Aluja et al. 1987), can be expected to occur in Costa Rica as well.
Our extensive sampling of infested fruits from about 200 localities during 1985-1986
confirms the findings of Jir6n and Zeled6n (1979), that three species, A. obliqua, A.
striata, and A. serpentina, are the major tephritid pests of the most commonly con-
sumed fruits in Costa Rica (Jir6n and Hedstrom 1988). A. fraterculus and A. ludens are
not of economic significance here, and our examination of specimens of the latter species
revealed minor differences between females from Costa Rica and from Mexico and
Guatemala. The significance of these differences needs to be further investigated, espe-
cially considering the apparent difference in host preferences between populations from
the two areas.


We thank the following colleagues of L. F. Jir6n and J. Soto-Manitiu at the Univer-
sidad de Costa Rica for their assistance in the field survey: R. G. Mexz6n, H. J. Lezama,
I. M. Gonzalez, I. Chac6n, and L. E. Cordero. We also thank L. D. G6mez (Jardin
BotAnico Las Cruces), H. P. Sauter (Univ. of California, Riverside), M. Aluja S. (Univ.
of Massachussetts), M. A. Condon (Smithsonian Institution), and R. D. Gordon, F. C.
Thompson, and R. V. Peterson (Systematic Entomology Laboratory) for their reviews
of earlier drafts of the manuscript. We are grateful to the curators of the collections
listed in the methods section for the loan of valuable study material. The field studies
conducted in this project were partially supported by the Consejo Nacional de Inves-
tigaciones Cientificas y Tecnol6gicas de Costa Rica (CONICIT) and the Vice-rectoria
de Investigaci6n, Universidad de Costa Rica. Work by L. F. Jir6n at the National
Museum of Natural History, Washington, D.C. was supported by a short term visitors
grant from the Smithsonian Institution.


AND P. LIEDO. 1987. A survey of the economically important fruit flies (Dipt-
era: Tephritidae) present in Chiapas and a few other fruit growing regions in
Mexico. Florida Entomol. 70: 320-329.
J. HENDRICHS. 1987. Natural host plant survey of the economically important

June, 1988

Jir6n et al.: Costa Rica Anastrepha 137

fruit flies (Diptera: Tephritidae) of Chiapus, Mexico. Florida Entomol. 70: 329-
ALUJA, M. AND I. MARTINEZ. 1985. Manejo integrado de las moscas de las frutas
(Diptera: Tephritidae). Program Mosca del Mediterraneo, SARH, Mexico. 241
BAKER, A. C., W. E. STONE, C. C. PLUMMER, AND M. MCPHAIL. 1944. A review
of studies on the Mexican fruitfly and related Mexican species. United States
Dept. Agric. Misc. Publ. No. 531, 155 pp.
BERG, G. H. 1959. Mosca de las frutas. I parte. Manual Entomol6gico para Inspectores
de Cuarentena Vegetal, OIRSA, San Salvador. 66 p.
CARABALLO, J. 1981. Las moscas de frutas del genero Anastrepha Schiner, 1868
(Diptera: Tephritidae) de Venezuela. Ph.D. dissertation, Universidad Central de
Venezuela, Maracay. 210 pp.
ESKAFI, F. M. AND R. T. CUNNIGHAM. 1987. Host plants of fruit flies (Diptera:
Tephritidae) of economic importance in Guatemala. Florida Entomol. 70: 116-
FERNANDEZ-YEPEZ, F. 1953. Contribuci6n al studio de las moscas de las frutas del
g6nero Anastrepha Schiner (Diptera; Trypetidae) de Venezuela. Caracas, II Con-
greso de Ciencias Naturales y afines 7: 5-41.
FOOTE, R. H. 1967. Family Tephritidae (Trypetidae, Trupanidae). IN: P. E. Vanzolini
and N. Papavero, eds., A catalogue of the Diptera of the Americas South of the
United States, Fasc. 57. Dept. Zool., Secr. Agric., Sao Paulo. 91 pp.
GONZALEZ, I. M., H. J. LEZAMA, AND L. F. JIR6N. 1988. Anastrepha fruit flies
(Diptera; Tephritidae) in Costa Rica: Three new records. Rev. Biol. Trop. 36:
(in press).
HEDSTROM, I., J. SOTO-MANITIU, AND L. F. JIR6N. 1985. Two species of fruit flies,
Anastrepha (Diptera; Tephritidae), new to Costa Rica. Rev. Biol. Trop. 33:
JIR6N, L. F. AND I. HEDSTROM. 1988. Occurrence of fruit flies of the genera Anas-
trepha and Ceratitis (Diptera; Tephritidae), and their host plant availability in
Costa Rica. Florida Entomol. 71: 62-73.
JIR6N, L. F. and R. ZELEDON. 1979. El g6nero Anastrepha (Diptera, Tephritidae)
en las principles frutas de Costa Rica y su relaci6n con pseudomiasis humana.
Rev. Biol. Trop. 27: 155-161.
NORRBOM, A. L. 1985. Phylogenetic analysis and taxonomy of the cryptostrepha,
daciformis, robusta and schausi species groups of Anastrepha Schiner (Diptera;
Tephritidae). Ph.D. dissertation, The Pennsylvania State University. 355 pp.
NORRBOM, A. L. AND K. C. KIM. 1988. Revision of the schausi group of Anastrepha
Schiner, with a discussion of the terminology of the female terminalia in the
Tephritoidea. Ann. Entomol. Soc. America 81: 164-173.
PICADO, C. 1920. Historia del gusano de la guayaba. Publ. coleg. Seforitas (Serie A.
N 2). San Jose, Costa Rica. 28 pp.
SALAS, L. A. 1957. Informe sobre el studio de la mosca del Mediterraneo en Costa
Rica. Publ. Univ. Costa Rica, ser. Agron. no. 1. 153 pp.
SAUNDERS, J. L. 1978. Cassava production and vegetative growth related to control
duration of shoot flies and fruit flies. Cassava Protec. Workshop, CIAT, Cali,
Colombia, p. 215-219.
SOTO-MANITIU, J. AND L. F. JIRON. 1988. Anastrepha fruit flies (Diptera; Tep-
hritidae) in Costa Rica: Four new records. Rev. Biol. Trop. 36: (in press).
STANDLEY, P. G. 1937. Flora of Costa Rica, Part II. Publ. 392. Field Museum of
Natural History, Chicago.
STEYSKAL, G. C. 1977. Pictoral key to species of the genus Anastrepha (Diptera:
Tephritidae). Entomological Society of Washington, Washington, D.C., 35 pp.
STONE, A. 1942. Fruit flies of the genus Anastrepha. United States Dept. Agric.,
Misc. Publ. No. 439. 112 pp.
TERRELL, E. E., S. R. HILL, J. H. WIERSEMA, AND W. E. RICE. 1986. A checklist
of names for 3,000 vascular plants of economic importance. United States Dept.
Agric., Agric. Handbook No. 505. 244 pp.
WHERVIN, L. W. VAN. 1974. Some fruit flies (Tephritidae) in Jamaica. Pans 20:

138 Florida Entomologist 71(2) June, 1988


Research Department
United States Sugar Corporation
Clewiston, Florida 33440


A list of insects and mites associated with sugarcane in Florida is presented.
Phytophagous species are listed along with their parasitoids and predators. A literature
review of sugarcane entomology in Florida is also given.


Se present una lista de insects y Acaros asociados con la cafia de azdcar en la
Florida. Se incluyen species que se alimentan de plants junto con sus parAsitos y de-
predadores. Tambi6n se da un resume de la literature de la entomologiA de la cafia de
azdcar en la Florida.

Sugarcane has been grown commercially in southern Florida since the 1920s and has
become one of the most important crops in the State. Some 400,000 acres are currently
grown each year in a general area extending around the lower half of Lake Okeechobee
in Glades, Hendry, Palm Beach, and Martin counties. Cane is planted during the fall
and winter months and reaches maturity 12 to 15 months later. Usually, one plant-cane
crop and two to four ratoon crops are harvested. Fields average around 40 acres in size,
but some are as large as 80 acres.
Sugarcane in Florida is attacked by a full complement of arthropod pests including
root, stalk, and foliage feeders. Different pest problems develop during different sea-
sons each year, but pest problems can occur all year long due to the subtropical climate
of the sugarcane region. Sugarcane pests in Florida were reviewed by Ingram et al.
(1938) and by Gifford (1964). During the last 23 years, however, a number of new pests
such as Perkinsiella saccharicida Kirkaldy and Melanaphis sacchari (Zehntner) have
appeared in Florida cane. In addition to new pests, new information on other pests and
their natural enemies has become available over the last 23 years.
During 1981-1987, I compiled records of insects and mites associated with sugarcane
in Florida along with their predators and parasitoids. The list is based on a literature
review and on specimens I collected in sugarcane. Frequent trips were made each year
to different areas across the sugarcane growing area of Florida to collect specimens.
Specimens were also collected during other sugarcane research projects. Many
phytophagous species were collected and held in a laboratory to determine if they had
been attacked by parasitoids.
Orders are presented alphabetically as are families within orders. Pest species are
listed along with their parasitoids and, in some cases, specific predators. The more
important pest species are denoted by an asterisk (*). General predators and some
noteworthy non-pest species are also listed. Throughout the list, insect and mite species
that were common in cane are denoted by superscript "a". All species I personally
observed are designated by superscript "b". Where possible and appropriate, a general
comment on each listed species is presented along with references pertaining to the

Hall: Florida Sugarcane Pests

species in Florida sugarcane. Finally, the taxonomist who identified specimens for me
is named (det. = determined by). Affiliations for most of the taxonomists are ab-
breviated: "BMNH" is the British Museum of Natural History, London; "BRI" is the
Biosystematics Research Institute, Agriculture Canada, Ottawa, Ontario; "FSCA" is
the Florida Center for Arthropod Systematics, Florida State Collection of Arthropods,
Division of Plant Industry, Gainesville, Florida; "SI" is the Smithsonian Institute,
Washington, D.C.; and "USDA" is the Systematic Laboratory, Insect Identification and
Beneficial Insect Introduction Institute, United States Department of Agriculture,
Beltsville, Maryland.


Abacarus officinari K.-Collected by Ru Nguyen near Belle Glade during 1983.

*Oligonychus stickneyi (McGregor)ab-A common pest of leaves. Damage by this
mite is sometimes extensive. Miticides occasionally used for control. (Hall
1986a, Strayer 1975) (det. H. A. Denmark, FSCA)
Phytoseiidae-Fundiseius cesi (Muma)b, Neoseiulus umbraticus
(Chant)b (det. H. A. Denmark, FSCA)
Paratetranychus simplex (Banks)-(Box 1953, Ingram et al. 1951)

Steneotarsonemus bancrofti (Michael)-First found on sugarcane stalks in Florida
at Canal Point in 1922 by E. W. Brands. (Ingram et al. 1938, Ingram et al.

Pronematulus sp.b-The feeding habits of this mite in cane were not known. (det.
H. A. Denmark, FSCA)


Lobopoda spb-The larvae resemble wireworms and were occasionally encountered
in the soil. Apparently saprophytic. (det. R. E. Woodruff, FSCA)

Calosoma scrutator (Fab.)b-A general predator.
Scarites subterraneous Fab.ab-A general predator common in cane. (det. R. E.
Woodruff, FSCA)

Prionus sp-(Box 1953)

Diabrotica spp-(Gifford 1964)

General predators of aphids and other small arthropods:
Coleomegilla maculata fuscilabris (Mulsant)b-(det. R. D. Gordon)
Cycloneda sanguinea (Lin.)ab--(Gifford 1964) (det. R. D. Gordon)

Florida Entomologist 71(2)

June, 1988

Diomus melsheimeri Weise-(Gifford 1964) (There were no FSCA records for this
species in Florida)
Diomus terminatus Sayab-(Gifford 1964, Hall 1987a)(det. R. D. Gordon, USDA)
Hippodamia convergens Guerinab-(det. R. D. Gordon)
Ola v-nigrum Mulsantb-(det. R. D. Gordon)
Stethorus utilis (Horn)b-Observed feeding on eggs of Oligonychus stickneyi.
(det. R. D. Gordon)

Pachneus litus (Germar)b-Large infestation levels of adults observed in several
fields during 1987. (det. F. N. Young, FSCA)
Sphenophorus coesifrons (Gyllenhal)b-Low population levels sometimes found in
cane, large levels uncommon. (Hall and Remik 1982) (det. R. E. Woodruff,
Sphenophorus venatus vestitus (Chittenden)b-Low population levels sometimes
found in cane, large levels uncommon. (Hall and Remik 1982) (det. R. E.
Woodruff, FSCA)

Aeolus dorsalis Say-(Box 1953, Wilson 1940)
A. perversus (Brown)b--Uncommon in cane. (det. E. C. Becker, BRI)
Conoderus amplicollis (Gyllenhal)ab-Common wireworm in cane. (Box 1953, In-
gram et al. 1938, Ingram et al. 1951, Wilson 1940, Wilson 1946) (det. E. C.
Becker, BRI)
C. falli (Lane)ab--Common wireworm in cane. (Gifford 1964, Strayer 1975) (det.
E. C. Becker, BRI)
C. rudis Brownab--Common wireworm in cane. (det. E. C. Becker, BRI)
C. scissus (Schaeffer)b-Adults common in light-traps operated in cane fields. (det.
E. C. Becker, BRI)
Parasitoid reared from unidentified wireworms of the genus Conoderus:
Anomalon ejuncidum (Say)b (Ichneumonidae) (det. V. K. Gupta, FSCA)
Dolopius sp-(Wilson 1940)
Glyphonyx bimarginatus Schaefferab-Small wireworm often present. (det. E. C.
Becker, BRI)
Ischiodontus sp.b--Localized populations of this wireworm are sometimes en-
countered. (det. T. J. Spilman, USDA)
*Melanotus communis (Gyllenhal)ab-An important soil pest. Insecticides routinely
applied at planting time for control. (Box 1953, Bregger et al. 1959, Cherry
and Hall 1986, Gifford 1964, Hall 1982, Hall 1985b, Hall and Cherry 1985,
Ingram et al. 1938, Ingram et al. 1951, Samol and Johnson 1973, Strayer
1975, Wilson 1940, Wilson 1946) (det. E. C. Becker, BRI)
Bethylidae-Pristocera armifera (Say)b (det. A. S. Menke,
Neotrichophorus carolinensis (Schaeffer)--A large wireworm occasionally seen in
cane. (det. E. C. Becker, BRI)
Orthostethus infuscatus (Germar)b--Adults sometimes collected at light-traps op-
erated in cane fields. (Box 1953, Ingram et al. 1938, Ingram et al. 1951)
(det. E. C. Becker, BRI)

Carpophilus humeralis (Fab.)b--Commonly associated with decaying seedpieces
in the soil. (det. W. A. Connell, USDA)

Hall: Florida Sugarcane Pests

Anomola marginata (Fab.)ab-Common. (Gordon and Anderson 1981, Hall 1987b,
Prewitt and Summers 1981)
Cyclocephala parallel Caseyab-Common. (Boucias et al. 1986, Bregger et al.
1959, Cherry 1984a, Cherry 1985, Gifford 1964, Gordon and Anderson
1981, Hall 1987b, Ingram et al. 1938, Ingram et al. 1951, Prewitt and Sum-
mers 1981, Strayer 1975, Watve and Shuler 1985, Watve et al. 1981)
Dyscinetus morator (Fab.)b-Adults commonly collected at light traps in cane
fields, but larvae rarely found in cane. (Gordon and Anderson 1981)
Euphoria sepulchralis (Fab.)b-Common in some areas. (Gordon and Anderson
1981, Strayer 1975)
*Ligyrus subtropicus (Blatchley)ab--This large grub can cause extensive damage.
Infestations common but usually localized and occur primarily in organic
soils. (Boucias et al. 1986, Cherry 1983a, Cherry 1983b, Cherry 1984a,
Cherry 1984b, Cherry 1985, Gordon and Anderson 1981, Hall 1987b, Mil-
ler and Bell 1985, Prewitt and Summers 1981, Sosa 1984a, Sosa 1984b,
Sosa and Beavers 1985, Strayer 1975, Summers 1974, Summers 1978a,
Summers et al. 1981, Watve and Shuler 1985, Watve et al. 1981)
Phyllophaga latifrons (LeConte)ab--Common in some areas. (Box 1953, Gordon
and Anderson 1981, Hall 1987b, Ingram et al. 1938, Ingram et al. 1951,
Prewitt and Summers 1981)
Pupae of the following parasitoid species were often collected in the soil around
cane infested by grubs, notably C. parallel:
Tiphiidae-Tiphia floridana floridana Robertsonb, Tiphia sppb
(det. R. W. Carlson, USDA)
Scoliidae-Scolia bicincta Fab.b (det. R. W. Carlson, USDA)

General predators in sugarcane. (Adams et al. 1981):
Anotylas insignitus (Gravenhorst)
A. nanus (Erichson)
Acrotona hebiticornis Notman
Atheta macrops Notman
A. spp
Belonuchus pallidus Casey
Diochus schaumii Kraatz
Meronera venustula (Erichson)
Philonthus hepaticus Erichson
Sunius debilicornis (Wollaston)
Thoracophorus sp
Tinotus sp


Salina beta Christiansen & Bellingerab-A yellowish, fast-moving species often
present on the underside of sugarcane leaves. (det. M. M. Bush-Davis, SI)

Lepidocyrtus cyaneus Tullberg-(Box 1953, Ingram et al. 1951)
Onychiurus armatus (Tullberg)-(Ingram et al. 1951)
Pseudosinella violent (Folsom)-(Ingram et al. 1951)

Florida Entomologist 71(2)


Labidura riparia (Pallas)ab-A general predator, notably of Diatraea saccharalis.
(Ingram et al. 1951)


Robberfly larvae sometimes appear to be important predators of scarab grubs.
Diogmites neoternatus Bromley-(det. A. G. Scarbrough, FSCA) (R. H. Cherry,
pers. comm.)
prob. Diogmites esuriens Bromleyb-(det. S. W. Bullington, Virginia Polytech.
Instit. and St. Univ.)
Triorla interrupta (Macquart)b-(det. S. W. Bullington, Virginia Polytech. Instit.
and St. Univ.)

Euxesta stigmatias Loew-Adults often in sugarcane fields. (det. H. V. Weems,
FSCA) (0. Sosa, pers. comm.)

Allograpta exotica (Wiedemann)b-A general predator of aphids and mites. (Hall
1987a) (det. F. C. Thompson, USDA)


Cyrtomenus ciliatus (Palisot de Beauvois)b-Low population levels found in a few
fields. (det. J. E. Eger, Dow Chemical USA)
Pangaeus bilineatus (Say)b-Low population levels found in a few fields. (det. J. E.
Eger, Dow Chemical USA)

Andrallus spinidens (Fab.)b-Occasional predator, notably of Mocis latipes. (det.
J. E. Eger, Dow Chemical USA)
Podisus maculiventris (Say)b--Occasional predator. (det. J. E. Eger, Dow Chem-
ical USA)


Hysteroneura setariae (Thomas)-(Gifford 1964, Ingram et al. 1938) (There were
no FSCA records for this aphid in Florida sugarcane)
Melanaphis sacchari (Zehntner)ab-Common and sometimes present at large
levels. (Hall 1987a, Mead 1978, Summers 1978b)
Aphidiidae-Lysiphlebus testaceipes (Cresson)b (det. P. M.
Marsh, USDA)
Rhopalosiphum maidis (Fitch)-(Gifford 1964, Ingram et al. 1938) (There were no
FSCA records for this aphid in Florida sugarcane)
*Sipha flava (Forbes)ab-An important pest of cane. Common but frequently local-
ized within a cane field. Insecticides occasionally used for control. (Bregger

June, 1988

Hall: Florida Sugarcane Pests 143

et al. 1959, Gifford 1964, Hoffman 1959, Ingram et al. 1938, Ingram et al.
1951, Strayer 1975)

Prosapia bicincta (Say)b-Low population levels sometimes occur but no economic
damage has been reported. (Gifford 1964, Ingram et al. 1938, Ingram et al.
1951, Strayer 1975)

Draeculacephala portola Ballab--Common but has not been reported to cause eco-
nomic damage in Florida cane. (Gifford 1964, Ingram et al. 1951, Strayer
Mymaridae-Lymaenon koebelei (Perkins)
Trichogrammatidae-Ufens niger (Ashmead)
Homalodisca insolita (Walker)b-(det. J. P. Kramer, USDA)

Two leafhoppers, Graminella nigrifrons (Forbes) and Balclutha caldwelli
Blocker, were collected in sweepnet samples taken in young cane fields. Draecu-
lacephala product (Walker) and D. inscripta (Van Duzee) were collected at black-
light traps operated in cane fields. (cicadellids det. J. P. Kramer, USDA)

Myndus crudus Van Duzeeb-Sometimes present in low numbers. (det. J. P.
Kramer, USDA)

*Pulvinaria elongata Newsteadab-Damage by this scale can be severe. Localized
infestations sometimes occur. These scales frequently controlled by nat-
ural enemies. (Ingram et al. 1951, Williams et al. 1969)
Aphelinidae-Coccophagus lycimnia (Walker)b, Coccophagus
spb, Encarsia spb
Encyrtidae-Homosemion spb, Metaphycus flavus (Howard)b,
Metaphycus sppb, prob. Trichomasthus spb. Cheiloneurus
pulvinariae Dozierb was a hyperparasite of M. flavus. (para-
sitoids det. M. E. Schauff, USDA)

Perkinsiella saccharicida Kirkaldyab--Common but has not caused serious damage
in Florida cane. (Hall 1985b, Nguyen et al. 1984, Sosa 1982, Sosa 1983b;
Sosa 1985b, Sosa and Cherry 1982)
Miridae-Tytthus parviceps (Reuter)b (det. T. J. Henry, USDA)
Mymaridae-Anagrus sp
Saccharosydne saccharivora Westwoodab-Common but usually not present in
large enough numbers to be damaging. (Bregger et al. 1959, Gifford 1964,
Ingram et al. 1938, Ingram et al. 1951, Strayer 1975)
Miridae-Tytthus parviceps (Reuter)b (det. T. J. Henry, USDA)
Reduviidae-Zelus longipes (Lin.) (Released into cane during
1960 and may still be present; F. D. Bennett, pers. comm.)

Florida Entomologist 71(2)

June, 1988

Mymaridae-Anagrus armatus (Ashmead)
Trichogrammatidae-Paracentrobia (=Abbella) spp
Dryinidae-Psuedogonatopus variistriatus Fen.b
Stylopidae-Stenocranophilus quadratus Pierce

The following delphacids were collected in sweepnet samples taken in
young cane fields: Sogatodes molinus Fennah, Delphacodes propinqua (Fieber),
Delphacodes puella (Van Duzee), Sogatella kolophon (Kirkaldy), Pissonotus piceus
(Van Duzee), and Megamelus gracilis Beamer. (delphacids det. J. P. Kramer,

Aspidiella sacchari (Cockerell)-(Box 1953, Dekle 1976, Williams et al. 1969)
Encyrtidae-Adelencyrtus moderatus (Howard) (det. J. Noyes,
BMNH) (F. D. Bennett, pers. comm.)

Dysmicoccus boninsis (Kuwana)ab-Common but not regarded as an economic pest.
(Box 1953, Gifford 1964, Ingram et al. 1938, Ingram et al. 1951, Strayer
1975, Warner 1941)
Encyrtidae-Metaphycus sp.b (det. M. E. Schauff, USDA)
Dysmicoccus brevipes Cockerell-On sugarcane at the Miami World Collection of
Sugarcane. (det. A. Hamon, FSCA) (F. D. Bennett, pers. comm.)


Over 30 different species of ants have been reported in Florida sugar-
cane fields (Adams et al. 1981a, Adams et al. 1981b, Carroll 1970, Prewitt et al.
1981). The imported red fire ant, Solenopsis invicta Buren, is common. It may be
an important predator of Diatraea saccharalis and other insects.


Dicranoctetes sp.b--Low population levels of this leafminer sometimes occur. (Hall
1983) (det. R. W. Hodges, USDA)
Eulophidae-Chrysocharis imbrasus (Walker)b, Cirrospilus sp.b
(det. M. E. Schauff, USDA)

prob. Lerema accius Abbot and Smithb-Uncommon.

Agrotis ipsilon (Hufnagel)b-(Box 1953, Ingram et al. 1938, Ingram et al. 1951)
(det. R. W. Poole, USDA)
Agrotis malefida Guenee-(Box 1953, Ingram et al. 1938, Ingram et al. 1951)
Agrotis subterranea (Fab.)b--(Box 1953, Ingram et al. 1951) (det. R. W. Poole,


Hall: Florida Sugarcane Pests 145

Anicla infecta (Ochsenheimer)b-(det. R. W. Poole, USDA)
Elaphira chalcedonia (Hubner)-(Box 1953, Ingram et al. 1938, Ingram et al. 1951)
Elaphira nucicolora (GuenBe)-(Box 1953, Ingram et al. 1938, Ingram et al. 1951)
Leucania latiuscula Herrich-Schaffer-(Gifford 1964, Ingram et al. 1938, Ingram
et al. 1951, Strayer 1975, Wylie 1946)
Tachinidae-Archytas piliventris Van der Wulp, Belvosia luteola
Coquillett, Eucelatoria rubentis (Coquillett)
Braconidae-Cotesia floridanus Muesebeck, C. rufocoxalis
Eulophidae-Euplectrus plathypenae Howard
Ichneumonidae-Ichneumon sp near laetus Brulle, Netelia
emorsa Townsend, Ophion ancyloneura Cameron, undet. sp
of Ichneumonini
Leucania scirpicola (Guenee)b--Low levels are common in some areas. (det. R. W.
Poole, USDA)
Braconidae-Cotesia rufocoxalis (Riley)b (det. P. M. Marsh,
Meropleon cosmion Dyar-(Ingram et al. 1951)
Mocis latipes Gueneab-1Infestations frequently occur, especially on some vari-
eties. Can cause extensive defoliation. (Gifford 1964, Hall 1985c, Strayer
1975) (det. D. M. Weisman, USDA)
Sarcophagidae-Sarcodexia sternodontis (Townsend)b (det.
N. E. Woodley, USDA)
Tachinidae-Chetogena sp.b (det. N. E. Woodley, USDA)
Ichneumonidae-Enicospilus sp.b (det. L. A. Stange, FSCA),
Gambrus ultimus (Cresson)b (det. V. K. Gupta, FSCA)
Braconidae-Rogas sp.b (det. S. R. Shaw, USDA)
Chalcididae-Spilochalcis sp.b (det. E. E. Grissell, USDA)
Prodenia eridania (Cramer)-(Gifford 1964)
Spodoptera frugiperda J. E Smithab-Large infestations sometimes occur. (Gif-
ford 1964, Ingram et al. 1938, Ingram et al. 1951, Strayer 1975) (det. R. W.
Poole, USDA)
Tachinidae-Lespesia archippivora (Riley)b (det. N. E. Woodley,
Braconidae-Meteorus autographae Muesebeckb (det. P. M.
Marsh, USDA)
Parasitoids reported to be active against some cutworms and armyworms
(Box 1953, Ingram et al. 1938):
Braconidae-Agathis texana (Cresson)
Ichneumonidae-Enicospilus purgatus (Say), Paniscus ocellata
Tachinidae-Eucelatoria comosa (Van der Wulp)

Diatraea crambidoides Grote-(Box 1953)
Diatraea evanescens Dyarb-Low infestation levels occur. (det. D. C. Ferguson,
*Diatraea saccharalis (Fab.)ab-An important, widespread pest. Insecticides often
used for control. (Adams et a. 1981b, Alvarez and Kidder 1981, Box 1953,

Florida Entomologist 71(2)

June, 1988

Bregger et al. 1959, Carroll 1970, Charpentier et al. 1965, Gifford 1964,
Gifford 1965, Gifford and Mann 1967, Hoffman 1959, Hall 1981, Hall 1986b,
Hall 1986c, Ingram et al. 1938, Ingram et al. 1951, Long and Hensley 1972,
Mathes et al. 1953, Prewitt et al. 1982, Reagan 1984, Reagan et al. 1973,
Rice 1981, Scaramuzza 1942, Sosa 1981, Sosa 1983a, Sosa 1985a, Strayer
1975, Summers 1976a, Summers 1976b, Summers et al. 1976, Summers
et al. 1977, Taylor 1944, Ulloa et al. 1982, Williams et al. 1969, Wilson
1941, Wilson 1942)
Braconidae-Agathis stigmatera (Cresson)b, Cotesia flavipes
(Cameron)b (det. P. M. Marsh, USDA)
Trichogrammatidaeb-Trichogramma fasciatum (Perkins), Tri-
chogramma minutum Riley
Elasmopalpus lignosellus (Zeller)ab--Large infestations sometimes occur. (Breg-
ger et al. 1959, Gifford 1964, Ingram et al. 1938, Ingram et al. 1951,
Mathes et al. 1953, Strayer 1975)
Tachinidae-Chetogena floridensis (Townsend)b (det. N. E.
Woodley, USDA)
Braconidae-Orgilus elasmopalpi Muesebeckb (det. P. M.
Marsh, USDA)
Herpetograma bipunctalis (Fab.)b-The southern beet webworm is uncommon in
cane. (det. J. B. Heppner, FSCA)
Marasmia trapezalis (Guenee)ab-Large infestations sometimes occur. (Strayer
1975) (det. D. C. Ferguson, USDA)
Braconidae-Agathis discolor (Cresson)b, Agathis texana (Cres-
son)b, Chelonus (Microchelonus) spb, Dolichogenidea spb,
Rogas laphygmae Viereckb (det. P. M. Marsh, USDA)
Tachinidae-Chetogena floridensis (Townsend)b (det. N. E.
Woodley, USDA)


Chrysoperla externa (Hagan)b-A general predator of aphids and mites. (det. L. A.
Stange, FSCA)

Micromus subanticus (Walker)b-A general predator of aphids and mites. (det.
L. A. Stange, FSCA)


Schistocera obscura (Fab.)bm-Localized infestations sometimes occur.

Gryllus assimilis (Fab.)b
Gryllus firmus Scudderb

Mole crickets sometimes kill young cane shoots.


Hall: Florida Sugarcane Pests 147

Scapteriscus acletus Rehn and Hebardb-(det. D. A. Nickle, USDA)
Scapteriscus vicinus Scudderb--(det. D. A. Nickle, USDA)


Ectopsocopsis cryptomeridae (Enderlein)b-Sometimes present during late spring.
Appeared to feed on the sugarcane rust fungus. (det. E. L. Mocford, Ill.
State Univ.)


I am grateful to the taxonomists who identified specimens for me. Rick A.
Armstrong provided invaluable assistance with collecting, mounting and submitting
specimens for identification. Diana Ford typed the manuscript. For their special assist-
ance during this project, I would like to thank F. D. Bennett, R. H. Cherry, O. Sosa,
E. C. Becker, S. W. Bullington, J. E. Eger, H. A. Denmark, M. S. Irey, and R. E.


ADAMS, C. T., C. S. LOFGREN, AND R. BEUTELL. 1981a. Interrelationship of the
red imported fire ant and staphylinids collected in pitfall traps in Florida sugar-
cane fields. Semi Annual Report of Research conducted on Imported Fire Ants,
Sci. Educ. Adm., Agr. Res., Gainesville, FL and Gulfport, MS. Rept. 80(1):
1981b. Interrelationship of ants and the sugarcane borer in Florida sugarcane
fields. Envir. Entomol. 10(3): 415-418.
ALVAREZ, J. AND G. KIDDER. 1981. Economic thresholds for sugarcane borer popu-
lations in Florida. Proc. Amer. Soc. Sugar Cane Techn. 9: 104.
BOUCIAS, D. G., R. H. CHERRY, AND D. L. ANDERSON. 1986. Incidence of Bacillus
popilliae in Ligyrus subtropicus and Cyclocephala parallel (Coleoptera:
Scarabaeidae) in Florida sugarcane fields. Envir. Entomol. 15: 703-705.
Box, H. E. 1953. List of sugar-cane insects-a synonymic catalogue of the sugar-cane
insects and mites of the world, and of their insect parasites and predators, ar-
ranged systematically. London: Commonwealth Instit. Entomol., 41, Queen's
Gate, London, S.W.7. 101 pp.
TEL. 1959. A brief review of sugarcane research in Florida, 1939-1959. Proc.
Soil and Crop Sci. Soc. Fla. 19: 287-294.
CARROLL, J. F. 1970. Role of ants as predators of the sugarcane borer, Diatraea
saccharalis. Unpub. MS Thesis, University of Fla. 70 pp.
CHARPENTIER, L. J., J. R. GIFFORD, AND R. MATCHES. 1965. Present status of
biological control of the sugarcane borer in continental United States. Proc. In-
ternat'l. Soc. Sugar Cane Techn. 12: 1287-1294.
CHERRY, R. H. 1983a. Flooding to control the grub Ligyrus subtropicus in Florida
sugarcane. Univ. Fla., Belle Glade Agr. Res. Ed. Center Res. Rept. EV-1983-7.
7 pp.
1983b. Contact toxicities of ten insecticides to the sugarcane grub, Ligyrus
subtropicus (Coleoptera: Scarabaeidae). Fla. Entomol. 66(4): 503-506.
1984a. Spatial distribution of white grubs (Coleoptera: Scarabaeidae) in
Florida sugarcane. J. Econ. Entomol. 77: 1341-1343.
1984b. Flooding to control the grub Ligyrus subtropicus (Coleoptera:
Scarabaeidae) in Florida sugarcane. J. Econ. Entomol. 77: 254-257.

148 Florida Entomologist 71(2) June, 1988

1985. Seasonal phenology of white grubs (Coleoptera: Scarabaeidae) in Florida
sugarcane fields. J. Econ. Entomol. 78(4): 787-789.
CHERRY, R. H. AND D. G. HALL. 1986. Flight activity of Melanotus communis
(Coleoptera: Elateridae) in Florida sugar cane fields. J. Econ. Entomol. 79:
DEKLE, G. W. 1976. Florida armored scale insects. Arthropods of Florida. Fla. Dept.
Agr. Vol. 3, 345 pp.
GIFFORD, J. R. 1964. A brief review of sugarcane insect research in Florida, 1960-
1964. Proc. Soil and Crop Sci. Soc. Fla. 24: 449-453.
GIFFORD, J. R. 1965. Goniozus indicus as a parasite of the sugarcane borer. J. Econ.
Entomol. 58: 799-800.
GIFFORD, J. R. AND G. A. MANN. 1967. Biology, rearing, and a trial release of
Apanteles flavipes in the Florida Everglades to control the sugarcane borer. J.
Econ. Entomol. 60: 44-47.
GORDON, R. D. AND D. M. ANDERSON. 1981. The species of Scarabaeidae (Coleopt-
era) associated with sugarcane in south Florida. Fla. Entomol. 64(1): 119-138.
HOFFMAN, C. H. 1959. Entomology Research Division programs relating to Florida
insect problems. Fla. Entomol. 42: 1-10.
HALL, D. G. 1981. A biological-chemical IPM program for the sugarcane borer. Proc.
2nd Inter-Amer. Sugar Cane Sem.-Insect and Rodent Pests: 89-95.
S1982. A parasite, Pristocera armifera (Say), of the wireworm Melanotus com-
munis (Gyll.) in south Florida. Fla. Entomol. 65: 574.
1983. A leaf miner, Dicranoctetes sp (Lepidoptera: Elachistidae), infesting
sugarcane in south Florida. Fla. Entomol. 66: 521.
- 1985a. Damage by the corn wireworm, Melanotus communis (Gyll.), to plant
cane during germination and early growth. J. Amer. Soc. Sugar Cane Techn. 4:
S1985b. Sugarcane delphacid control, small plot insecticide test, 1984. Insect.
and Acar. Tests. 10: 248-249.
.1985c. Parasitoids of grasslooper prepupae and pupae in south Florida sugar-
cane. Fla. Entomol. 68: 486-487.
.1986a. Oligonychus stickneyi (McGregor): a mite pest of sugarcane in Florida.
J. Amer. Soc. Sugar Cane Techn. 6: 134-135.
S1986b. Seasonal activity of parasitoids against sugarcane borer larvae in
Florida. J. Amer. Soc. Sugar Cane Techn. 6: 19-23.
.1986c. Sampling for the sugarcane borer (Lepidoptera: Pyralidae) in sugar-
cane. J. Econ. Entomol. 79: 813-816.
S1987a. The sugarcane aphid, Melanaphis sacchari, in Florida sugarcane. J.
Amer. Soc. Sugar Cane Techn. 7: 26-29.
.1987b. Seasonal flight activity of sugarcane grubs in Florida. J. Amer. Soc.
Sugar Cane Techn. 7: 39-42.
HALL, D. G. AND M. D. REMIK. 1982. Billbugs in Florida sugarcane. Sugar J. 45(5):
HALL, D. G. AND R. H. CHERRY. 1985. Contact toxicities of eight insecticides to the
wireworm Melanotus communis (Coleoptera: Elateridae). Fla. Entomol. 68:
INGRAM, J. W., H. A. JAYNES, AND R. N. LOBDELL. 1938. Sugarcane pests in
Florida. Proc. Internat'l. Soc. Sugar Cane Techn. 6: 89-98.
TIER. 1951. Pests of sugarcane and their control. U.S.D.A. Cir. 878. 38 pp.
LONG, W. H. AND S. D. HENSLEY. 1972. Insect pests of sugarcane. Ann. Rev.
Entomol. 17: 149-176.
DUGAS. 1953. Current status of sugarcane-insect investigations in the United
States. Proc. Internat'l. Soc. Sugar Cane Techn. 8: 560-567.
MEAD, F. W. 1978. Sugarcane aphid (Melanaphis sacchari (Zehntner))-Florida-
new continental United States record. Coop. Plant Pest Rept. 3(34): 475.
MILLER, J. D. AND M. G. BELL. 1985. Life cycle of the white grub and its effect on
sugarcane. J. Amer. Soc. Sugar Cane Techn. 4: 38-42.

Hall: Florida Sugarcane Pests 149

NGUYEN, RU, 0. SOSA, JR., AND F. W. MEAD. 1984. Sugarcane delphacid, Perkin-
siella saccharicida Kirkaldy 1903. Fla. Dept. Agric. and Consum. Serv., Div.
Plant Ind., Entomol. Circ. 265. 2 pp.
fields for effective sugarcane borer control. J. Amer. Soc. Sugar Cane Techn. 1:
PREWITT, J. C. AND T. E. SUMMERS. 1981. White grubs of sugarcane in south
Florida. Proc. 2nd Inter-Amer. Sugar Cane Sem.-Insect and Rodent Pests:
1981. Known distribution of the imported fire ant Solenopsis invicta Buren in
Florida sugarcane fields: benefit or problem for the future. Proc. Amer. Soc.
Sugar Cane Techn. 8: 160.
REAGAN, T. E. 1984. Insecticide resistance studies with Florida and Louisiana sugar-
cane borer populations. J. Amer. Soc. Sugar Cane Techn. 3: 90.
REAGAN, T. E., S. D. HENSLEY, AND J. B. GRAVES. 1973. Status of insecticide
resistance in sugarcane borer populations in Louisiana. J. Econ. Entomol. 66:
RICE, E. R. 1981. Biological-chemical control of the sugarcane borer in Florida. Sugar
J. 43(9): 17-19.
SAMOL, H. H. AND S. R. JOHNSON. 1973. Effect of some soil pesticides on sugarcane
yields in Florida. Proc. Amer. Soc. Sugar Cane Techn. 2: 37-40.
SCARAMUZZA, L. C. 1942. Results attained in the biological control of Diatraea sac-
charalis (F.) in Florida. J. Econ. Entomol. 35: 642-645.
SOSA, 0., JR. 1981. Sugarcane borer, Diatraea saccharalis, in Florida: a review.
Proc. 2nd Inter-Amer. Sugar Cane Sem.-Insect and Rodent Pests: 145-152.
1982. Discovery of a new insect pest of sugarcane in Florida, Perkinsiella
saccharicida Kirkaldy, a North American record. Proc. 3rd Inter-Amer. Sugar
Cane Sem.-Varieties and Breeding: 223-226.
1983a. Sugar cane borer survey of the 1980-1981 sugarcane variety tests in
Florida. J. Amer. Soc. Sugar Cane Techn. 2: 86-87.
1983b. Sugarcane delphacid discovered in Florida. Sugar J. 45: 16.
1984a. Effect of white grub (Coleoptera: Scarabaeidae) infestations on sugar-
cane yields. J. Econ. Entomol. 77: 183-185.
- 1984b. Losses caused by the white grub Ligyrus subtropicus in sugarcane. J.
Amer. Soc. Sugar Cane Techn. 3: 91-92.
1985a. Evaluation of sugar cane clones for resistance to the sugar cane borer
in Florida. J. Amer. Soc. Sugar Cane Techn. 4: 118.
1985b. The sugarcane delphacid, Perkinsiella saccharicida (Homoptera: De-
lphacidae), a sugarcane pest new to North America detected in Florida. Fla.
Entomol. 68(2): 357-360.
SOSA, 0., JR. AND R. CHERRY. 1982. Status of Perkinsiella saccharicida Kirkaldy,
the sugarcane planthopper in Florida, and some background information on this
pest. Belle Glade AREC Res. Rept. EV-1982-5. 8 pp.
SOSA, 0., JR. AND J. B. BEAVERS. 1985. Entomogenous nematodes as biological
control organisms for Ligyrus subtropicus (Coleoptera: Scarabaeidae) in sugar-
cane. Envir. Entomol. 14(1): 80-82.
STRAYER, J. 1975. Sugarcane insect control. Fla. Coop. Ext. Serv., Plant Protect.
Pointers, Ext.Entomol. Rept. No. 40. 6 pp.
SUMMERS, T. E. 1974. Florida sugarcane attacked by white grubs in 1972. Proc.
Amer. Soc. Sugar Cane Techn. 3: 124.
1976a. Some calculated costs and returns from monitoring Florida sugarcane
for the sugarcane borer, Diatraea saccharalis (F.). Proc. Amer. Soc. Sugar Cane
Techn. 5: 130.
1976b. Parasitism and control of the sugarcane borer by field released
Lixophaga diatraeae (T.). Proc. Amer. Soc. Sugar Cane Techn. 5: 131.
1978a. Flooding for the control of the white grub Bothynus subtropicus in
Florida. Proc. Amer. Soc. Sugar Cane Techn. 7: 128.

150 Florida Entomologist 71(2) June, 1988

-- 1978b. Sugarcane aphid (Melanaphis sacchari). Coop. Plant Pest Rept. 3(35):
SUMMERS, T. E., E. G. KING, D. F. MARTIN, AND R. D. JACKSON. 1976. Biological
control of Diatraea saccharalis (Lep.: Pyralidae) in Florida by periodic releases
of Lixophaga diatraeae (Dipt.: Tachinidae). Entomophaga. 21: 359-366.
SUMMERS, T. E., J. R. ORSENIGO, AND G. KIDDER. 1977. Field monitoring to deter-
mine threshold of economic loss due to the sugarcane borer, Diatraea sac-
charalis. Proc. Amer. Soc. Sugar Cane Techn. 6: 148-149.
Field evaluation of insecticides for control of the white grub Bothynus sub-
tropicus in Florida sugar cane. Proc. Amer. Soc. Sugar Cane Techn. 8: 162.
TAYLOR, D. J. 1944. Life history studies of the sugarcane moth borer, Diatraea
saccharalis Linn. Fla. Entomol. 27: 10-13.
ULLOA, M., M. G. BELL, AND J. D. MILLER. 1982. Losses caused by Diatraea
saccharalis in Florida. J. Amer. Soc. Sugar Cane Techn. 1: 7-10.
WARNER, J. D. 1941. Sugar cane mealy bug control on seed cane, with special refer-
ence to cold water treatment, at the North Florida experiment station. Fla.
Entomol. 24(3): 6-7.
WATVE, C. M. AND K. D. SHULER. 1985. A summary of research activities on white
grubs injurious to Florida sugarcane. J. Amer. Soc. Sugar Cane Techn. 4: 73-79.
WATVE, C. M., J. D. MILLER, M. G. BELL, AND K. D. SHULER. 1981. A summary
of research activities on white grubs injurious to Florida sugarcane. Proc. 2nd
Inter-Amer. Sugar Cane Sem.-Insect and Rodent Pests: 51-59.
1969. Pests of Sugar Cane. Elsevier Pub. Co. 568 pp.
WILSON, J. W. 1940. Preliminary report on wireworm investigations in the
Everglades. Fla. Entomol. 23: 1-6.
1941. Biological control of Diatraea saccharalis in the Florida Everglades
during 1940 and 1941. Fla. Entomol. 24: 52-57.
-- 1942. Correlation of sugar yields with the percent of joints bored by Diatraea
saccharalis (F.). Fla. Entomol. 25(2): 19-24.
--. 1946. Present status of the wireworm problem in south Florida. Fla. State
Hort. Soc. 59: 103-106.
WYLIE, W. D. 1946. Entomological control investigations. United States Sugar Cor-
poration, Research Department Doc. 123 (unpublished).


Department of Biological Sciences
Towson State University
Baltimore, Maryland 21204


Efferia caymanensis is described as a new species from Grand Cayman Island, West
Indies. This species is the first Efferia reported from the Cayman Islands and the first
member of the staminea group reported from the West Indies. Illustrations of the
terminalia are included.

Scarbrough: New West Indian Asilid


Se describe a Efferia caymanensis como una nueva especie de la Isla de Gran
Cayman, en las Indias Occidentales. Esta especie es el primer Efferia que se report
de la Isla Cayman, y el primer miembro del grupo de staminea reportado de las Islas
Occidentales. Se incluyen ilustraciones de la terminalia.

While conducting a survey of insects on Grand Cayman Island, West Indies, Dr.
Eugene Gerberg of Baltimore, Maryland collected 4 specimens of an asilid that proved
to be undescribed. The species belongs to Efferia Coquillett, a large genus restricted
to the New World (Hull, 1962). At least 105 species occur in the neartic (Wilcox 1966)
and 135 in the neotropics (Martin and Papavero 1970). All of the 16 West Indian species
belong to the aestuans group, and most are reported from the larger islands, the excep-
tions being E. cazieri (Curran) and E. vauriei (Curran) from the Bimini Islands in the
Bahamas (Curran 1953) and E. tortola (Curran) from Tortola Island in the British
Virgin Islands (Curran 1928). The two species from the Bahamas (Curran 1953) and E.
gossei Farr (Farr 1965) from Jamaica are the most recently described species from the
West Indies. None have been reported from the Cayman Islands.
This new species traces to the Efferia staminea species group in available keys (Hine
1919, Wilcox 1966) and is the first member of this group to be reported from the West
Indies. This group is characterized by having the furcation of the R4 + 5 vein being
distinctly before the base of the 1st medial cell, mesonotum with short hairs on the
anterior 1/2 and several long bristles on the posterior 1/2, and several strong bristles
on the margin of the scutellum. In addition, it differs from the aestuans group in that
the R5 vein curves forward and meets the costal margin anterior to the wing tip. In
the aestuans group, this vein curves posteriorly and meets the costal margin behind the
wing tip (Hine 1919). The following is presented at this time in order to provide a name
for the species in conjunction with the on-going insect survey of the island.

Efferia caymanensis, n. sp.
Figures 1-2
Male.-Body largely reddish, length 17.8 mm excluding terminalia. Face mostly
black with pale yellowish pollen, gibbosity red with sparse yellowish pollen, mystax
with largely black bristles above but sparsely intermixed and densely bordered by
white hairs and short bristles; lower 1/3 of mystax with mostly whitish bristles and 2-3
coarse black bristles laterally. Gena black with yellow-brown pollen grading to black
below and numerous black bristly hairs. Width of vertex about two-thirds that of frons
just above antenna. Palpus with vestiture mostly black, several white hairs on basal
1/2 ventrally and laterally. Proboscis black with long white hairs below. Antenna mostly
black, narrow apices of scape and pedicel and basal 1/3 of stylus reddish; dorsal vestiture
of scape and pedicel black, that ventrally and laterally mostly or entirely white; length
of stylus subequal to basal 3 antennal segments; flagellum length about 1/2 that of
stylus. Occiput mostly grayish pollinose with mostly white hairs, large black pollinose
spot on each side of occiput with several black hairs and 9-11 coarse, black bristles; 4-5
additional thinner black postocular bristles present laterally beyond black pollinose
Thorax mostly reddish with yellow to pale yellow pollen; scutum with 2 admedial
black longitudinal stripes, narrowly separated their entire lengths, and 2 black lateral
spots, posterior spot subtriangular; scutal vestiture black mostly or entirely, hair on
anterior 1/2 short, equal to subequal scape length; several bristles on posterior 1/3
becoming increasingly longer and thicker posteriorly. Dorsal hairs of scutellum mostly


Florida Entomologist 71(2)

June, 1988


',11 ,T 8

Fig 1. Efferia caymanensis n. sp., male terminalia, lateral view. Abb: Lp= lower
forceps; scale = 1.0 mm. 2. Efferia caymanensis n. sp., female terminalia, lateral view.
Abb: T8 = tergum 8; scale = 1.0 mm.
black with sparse white hairs intermixed on basal 1/3; margin of scutellum with 7-8 black
bristles and bristly hairs, the bristles about twice length of dorsal hairs. Mesoanepister-
num, mesokatepisternum, meron and metaepimeron largely to entirely black; vestiture
of mesoanepisternum and katatergite mostly black; mesokatepisternum with only sparse
black hairs; pleural vestiture otherwise whitish. Halter yellow.
Wing light yellowish with reddish to reddish brown veins. Costal dilation absent.
Vein R5 curving forward and meeting costa well before wing tip; spur of vein R4 almost
twice length of basal vein; furcation of vein R4 + 5 about two-thirds distance between
r-m crossvein and base of first medial cell; r-m crossvein at middle of discal cell.
Coxae black anteriorly, reddish elsewhere; fore coxa with abundant thin whitish
bristles and 3-4 coarse bristles, one of these black; hind coxa with 4-5 black hairs or
weak bristles. Trochanter black ventrally, hind trochanter with 4-5 black bristles. Fem-
ora largely reddish, black ventrally except narrow apices; all femora with abundant long
whitish hairs ventrally and posteriorly, hair densest and longest below fore femur;
anterior and dorsal surfaces of fore and mid femora entirely or mostly with short black
hairs, those of dorsal surface of fore femur mixed with long, thin, wavy black hairs;
hindfemur with mostly white hairs anteriorly and dorsally. Hind femur with 3 bristles
anteriorly, 3 subapically and dorsally, 7 anteroventrally and 5 in a cluster basopos-
teriorly. Tibiae with narrow apices slightly blackish red, hairs long, thin pale yellow;
tarsi, fore and hind tibiae with dense pad of short reddish or slightly orangish hairs
ventrally. Tarsi blackish red with black bristles; fore tarsus with several long, thin
whitish hairs laterally.
Abdomen reddish with slight blackish tint; terga 1-2 and 6-8 mostly black or blackish
pollinose, dark areas on terga 1-6 subtriangular, terga 7-8 with sparse blackish pollen;
call of tergum 1 sparsely and lateral margins of terga 2, 6-8 densely yellow pollinose;
remaining areas of terga white pollinose. Lateral margins of terga 1 and 2, narrow
apical margin of tergum 2 and terga 3-5 entirely with long white hairs, those on terga
3-5 dense and parted medially; tergum 6 with short white hairs laterally and basally;
black pollinose areas of terga 1, 2 and 6-8 mostly or entirely with black hairs, those on
terga 1 and 2 long laterally; terga 7-8 with hairs contrastingly short. Tergum 1 with 3-4
coarse black bristles. Sterna white pollinose with white hairs, those on basal 5 dense
and long, much shorter and sparser on apical 3 with sparse intermixed black hair.
Terminalia (Fig. 1) reddish with black hairs, surstylus long and slender, widest at
or just before middle, with a distinct posteroapical flange. Lower forceps very slender,
with an anterior adbasal thumblike process.


Scarbrough: New West Indian Asilid


Female.-Differs from male as follows: Length 13.8-17.0 mm, excluding terminalia.
Stylus 2.1 to 2.5 times length of flagellum. Facial pollen slightly darker yellow than in
males; occiput with black pollinose areas much smaller than on male. Hair of scutellum
variable, either mostly white or black; pleuron with hair mostly white, sparse black
hairs restricted to mesoanepisternum; katatergite of 1 female without black bristles.
Femora or hind coxa with 0-3 black bristles or hairs. Abdominal terga 1-4 with broad
triangular black pollinose spots dorsally, remaining areas white with mostly or entirely
white hairs; hairs of basal 5 segments much shorter and less abundant than on males.
Terga 5-7 with sparse black pollen, lateral margins with dense yellow pollen, hairs
mostly black. Ovipositor (Fig. 2) mostly black, sometimes partially reddish; length 5.0
mm, width at middle 1/9 length.
Holotype 3, Grand Cayman, BWI, E. Foldmans Bay, 8 May, 1970, E. J. Gerberg.
Allotype Y Grand Cayman Island, BWI, Boatswain Point, Lime Tree Estate, 16 Feb.
1985, E. J. Gerberg.
Paratypes.-9, same data and collector as allotype, 27 Feb, 1987; 2 9, Grande
Cayman Island, BWI, Vicksville, NW Pt. 17 Aug. 1975, E. J. Gerberg. Holotype,
allotype and 2 paratypes deposited in the United States National Museum of Natural
History, Washington, D. C. and 1 paratype in the private collection of G. J. Gerberg.
Etymology.-The name caymanensis refers to the type locality, Grand Cayman


I am very grateful to Dr. Eugene Gerberg for bringing this species to my attention
and the following curators for permitting me to make comparisons of this new species
with existing West Indian types and other specimens: Charles Vogt, Museum of Com-
parative Zoology, Harvard University [cubensis (Bromley), brunnescens (Bromley)],
Randall Schuh, American Museum of Natural History, New York [cazieri (Bromley),
forbesi (Curran), grandis (Hine), pachychaeta (Bromley), tortola (Curran)], T. H. Farr,
Institute of Science, Jamaica caudexx (Walker), fortis (Walker), fulvibarbis (Macquart),
grossi Farr, haloesus (Walker), portoricensis (Hine)], Lloyd Knutson, Biosystematics
and Beneficial Insects Institute, USDA, ARS, Beltsville, % United States National
Museum [the entire Efferia collection]; Eric Fisher, California Department of Agricul-
ture, Sacramento, and Robert Lavigne, University of Wyoming, Laramie, for reviewing
the manuscript. The Faculty Research Committee of Towson State University provided
funds for publication and reprints.


CURRAN, C. H. 1928. Insects of Porto Rico and the Virgin Islands. Diptera or two-
winged flies, 1:23, New York Academy of Sciences, Scientific Survey of Puerto
Rico and the Virgin Islands, Vol. 11. New York.
1953. The Asilidae and Mydaidae of the Bimini Islands, British West Indies
(Diptera). Amer. Mus. Nov. 1644: 4-5.
FARR, T. H. 1965. The robber-flies of Jamaica (Diptera: Asilidae). Bull. Inst. Jamaica,
Sci. Ser. 13(2): 31.
HINE, J. S. 1919. Robberflies of the genus Erax. Ann. Entomol. Soc. Amer. 12:
HULL, F. M. 1962. Robber flies of the World. The genera of the family Asilidae.
Smithson. Inst. Bull. 2: 474-479.
MARTIN, C. H. AND NELSON PAPAVERO. 1970. A catalogue of the Diptera of the
Americas south of the United States. Family Asilidae. Museu de Zoologia, Uni-
versidade de Sio Paulo. 35b: 62-69.
WILCOX, J. 1966. Efferia Coquillett in America north of Mexico (Diptera: Asilidae).
Proceed. Californis Acad. Sci. (ser. 4), 34(2): 85-234.

Florida Entomologist 71(2)


Program Moscamed, D.G.S.P.A.F.-S.A.R.H.
Apartado Postal 368
30700 Tapachula, Chiapas, MEXICO


Wild Mediterranean fruit flies Ceratitis capitata (Wied.) were exposed to 6 different
combinations of Jackson trap designs and colors, in a coffee plantation near Antigua,
Guatemala. Their pattern of arrival, landing, and capture or escape during different
hours of the day and trap environments were observed.
Maximum catch was recorded during rain-free afternoons, in tall and dense coffee
vegetation. The standard Jackson trap caught significantly more flies than the open
bottom Jackson trap, and yellow traps were significantly more effective than white
traps. An unknown number of flies were attracted to the surroundings of traps but did
not approach them during the allocated 30 min. observation periods. Of the flies ap-
proaching a trap, 18.6% were caught initially, and another 7.8% were caught during the
observation period as a result of agonistic interactions between males accumulating
inside the standard Jackson traps. The large presence of males remaining inside the
traps suggests that the daily percentage of capture is much higher than the 26.4%
documented during the short observation periods. Male aggregation and territoriality
inside and around the traps, as well the pattern of male-female attraction, indicate that
the parapheromone trimedlure has an effect similar to the medfly male pheromone.


Moscas del MediterrAneo silvestres (Ceratitis capitata Wied.) fueron expuestas a 6
diferentes combinaciones de disefo y color de trampas Jackson en una plantaci6n de
caf6 en las cercanias de la ciudad de Antigua, Guatemala. Se estudiaron los patrons de
arribo, aterrizaje y capture o escape durante diversas horas del dia y diseho de trampa.
MAxima capture se obtuvo durante tardes sin lluvia bajo condiciones de vegetaci6n
densa de caf6. La trampa Jackson estandar capture mAs moscas que la Jackson sin base
(differencia significativa) y las trampas amarillas capturaron mAs moscas que las blancas
(diferencia significativa). Un nuimero desconocido de moscas fue atraido a la cercania de
las trampas pero no se dirigi6 directamente a las mismas durante los periods de obser-
vaci6n (30 min). De las moscas que se dirigieron a las trampas, 18.6% fueron capturadas
inmediatamente y otro 7.8% fue capturado como resultado de interacciones agresivas
entire machos que se iban agregando dentro de la trampa. El gran nuimero de machos
que permanece dentro de la trampa sugiere que el porcentaje diario de capture es much
mayor al 26.4% determinado durante los cortos perfodos de observaci6n. El compor-
tamiento de agregaci6n y de territorialidad observado dentro y alrededor de las tram-
pas, asi como el patron de atracci6n macho-hembra, indican que la paraferomona trimed-
lure produce un efecto similar a la feromona natural emitida por los machos.

The successful application of the Sterile Insect Technique (SIT) is largely based on
the existence of an effective monitoring system that makes the early detection of the

'Current address and to whom reprint requests should be addressed: Dept. of Entomology, Univ. of Mas-
sachusetts, Amherst MA 01003.

June 1988

Villeda et al.: Medfly Behavior

target pest at low densities possible. Present large scale SIT programs against the
Mediterranan fruit fly generally use the standard sticky white Jackson trap baited with
trimedlure (Beroza et al. 1961). Dresner (1970) and Gutierrez (1972) compared this trap
with other traps, showing that the former is more sensitive at low medfly population
levels and preserves specimens for longer periods without contamination of unmarked
by marked flies. Ortiz & Reyes (1982) reached similar conclusions in studies conducted
in Egypt, as did Ortiz et al. (1983) working in Mexico under conditions of varying
elevations and climate.
Compared with other tropical fruit fly pests, this medfly trapping system has a
relatively small distance of attraction, mainly due to the low attractancy of trimedlure
(Sivinski & Calkins 1986). Consequently there have been various attempts to improve
trap attractancy. Evaluation of new trap features is, however, mostly based on trap
catches and the behavior of flies responding to these stimuli remains largely unknown.
Only Prokopy & Economopoulos (1975) observed olive flies (Dacus oleae) (Gmelin) ap-
proaching McPhail traps, and Davis et al. (1984) carried out observations of Caribbean
fruit flies Anastrepha suspense (Loew) approaching food and color-baited open bottom
Jackson traps.
The objective of this study was to systematically observe medfly behavior in re-
sponse to stimuli such as trap environment, trap design and color during different hours
of the day under natural conditions.


The study was carried out in "El Potrero", a coffee plantation in Antigua, Guatemala,
at an elevation of 1400 m, during April-May 1983, after the coffee harvest. An area of
approximatley 30,000 m2 was selected in which six circles replicationss) were distri-
buted, each formed by six traps (treatments). Distance between traps was 20 m. Three
of these circles were placed in plots of dense and tall (2.0-2.5m) coffee plants, whereas
the three others were located in plots of relatively open and short (1.0-1.5m) coffee. In
both types of plots there was a similar, small number of out of season coffee berries.
Two versions of the Jackson traps were used: the standard, triangular closed-bottom
trap, with a sticky insert; and a modified open trap, without bottom or insert, but sticky
inner sides (see Davis et al. 1984). Two colors were used, the white of the regular trap,
a yellow (Cromatone machine yellow No. 615), and a striped combination of both
(Greany et al. 1982). The six treatments, used in each of six replicates, were compared
in a split-plot design over two different coffee areas.
Traps were exposed for a total of 12 days, and were rotated each 24 hours one
position within their respective circle. Two ml of trimedlure were applied to new
Richmond cotton wicks in both trap types at the beginning of each of the six-day cycles.
The sticky material (Stickem Special), was applied to all the surface of either insert or.
Each day, traps or inserts were changed at 0600 h and from that time to 1800 h, all
traps were checked each two hours. Trapped flies were counted by sex and removed.
Every two hours, temperatures and relative humidities were recorded. The 2-h periods
between checking traps were spent observing flies and recording their behavior from
arrival to capture or escape. Equal observation time was assigned to each treatment
(30 min. at a time per trap type) and was rotated throughout the hours of the day, the 12
observation days, as well as between the 2 observers. The observation of flies remaining
inactive on, in or near a trap was abandoned once the 30 min. observation time had
elapsed. Flies that were observed and caught by the trap were removed from it at the
end of the observation period and placed into labeled vials with 50% ethyl alcohol so
that their gonads could be examined (Guillen 1983).

156 Florida Entomologist 71(2) June, 1988


With Rainy
Rainy Afternoon
Afternoon y S.D. Relative Temperature
Hour (X) (1 Day) (11 Days)1 Humidity C

1800-0600 (0) 0 0.0 99.0 12.0
0600-0800 (2) 43 9.4 6.2 76.9 18.6
0800-1000 (4) 142 134.4 28.7 44.5 25.4
1000-1200 (6) 121 125.8 40.7 39.5 27.5
1200-1400 (8) 72 125.5 64.7 49.5 28.2
1400-1600 (10) 0 210.5 71.6 60.3 22.5
1600-1800 (12) 0 254.4 59.2 98.4 19.9

'Linear regression equation y= 1.96 + 20.75X, significant at the 0.1% level.


I. Trap Catches
A total of 9837 flies were trapped, of which 0.2% were females and 99.8% were
males. Females caught were virgins. Daily catches varied considerably during the 12
days, fluctuating between an average of 10.5 and 43.9 flies per trap per day. In Table
1 the average number of flies trapped per 2-hour period of the day is shown with
average temperatures and relative humidities recorded at the end of each period. The
daily catch rate increased throughout the hours of the day, following a highly significant
linear regression (F = 99.8***, P<0.001). The maximum capture rate occurred in the
afternoon. On day six, however, it rained the whole afternoon and no flies were caught;
the only catch was in the morning.
Of the 9837 flies trapped, 60.5% were caught in the closed Jackson traps, whereas
the other 39.5% were caught in the open bottom traps (Table 2). Yellow traps caught
38.0% of all flies, striped 31.8% and white 30.2%. Closed traps were superior to the
open bottom traps (F = 7.2**, P<0.01), while the open yellow trap caught significantly
more flies than the open striped and the open white traps.
The difference between the two coffee environments was larger than between treat-
ments. Traps in the tall and dense coffee plots caught significantly more medflies than
those in short and open coffee (Table 2).
II. Behavioral Observations
Observations of fly behavior in relation to their pattern of approach to the traps,
showed that 75.4% of all flies observed arriving over a range of approx. 10 to 50 cm,
approached from above or on a horizontal plane, and 24.6% from below (Table 3). Also
43.8% of the flies approached a trap either in a direct or relatively straight flight, while
56.2% approached in a looping, zigzagging or curved flight, often landing first nearby.
Only in arrivals from below were significantly fewer direct than looping flights observed
(Table 3).
The observed initial landing site of medflies on the closed Jackson trap, shown in
Table 4, indicates a significant preference for the surfaces of the two trap sides: 80.8%
of the landings on 66.7% of the total trap surface, compared to only 19.2% of the
landings on both surfaces of the trap bottom, (33.3% of the trap surface), (X2 = 8.95**,

Villeda et al.: Medfly Behavior 157


Closed Closed Closed' Open Open Open' Environ-
Replicate White Yellow Striped White Yellow Striped ments2

Dense- 1 339 470 345 193 287 190
tall 2 443 525 568 322 400 303 352.la
coffee 3 356 407 406 234 327 222
Open- 1 233 279 219 114 198 142
short 2 205 208 214 131 149 148 194.4b
coffee 3 255 264 219 147 223 152
X Traps2 305.2ab 358.8a 328.5ab 190.2c 264.0b 192.8c 273.3

'Alternating two cm wide white and yellow bands.
2Values followed by the same letter are not significantly different at the 1% confidence level (environment) and
5% (trap types) according to Duncan's MRT.

P<0.01). Also, comparing observed initial landing site on sticky against non-sticky
surfaces, there was a significant preference for non-sticky surfaces on the closed Jackson
trap (X2 = 5.82*, P<0.05), and on the open bottom trap (X2 = 3.4**, P<0.01).
Of 1002 flies observed approaching a trap, only 18.6% were caught initially. Of those
flies that approached traps, 4.9% never contacted the trap during the 30 min. observa-
tion period, but only flew over it or circled it one or several times; 65.6% landed on a
non-sticky surface and either remained mostly motionless during the rest of the assigned
observation period (12.8%), or walked upon arrival for some time over non-sticky sur-
faces (52.8%), avoiding the stickum whenever their front legs and protracted mouth
parts contacted it. Flies that first landed on a non-sticky surface never were trapped
by walking onto the stickum. On a few occasions flies were seen sitting for short periods
on a several-days-old wick. Up to nine males were seen at one time mostly motionless
inside the closed Jackson trap. This behavior was common during hot weather, windy
and cloudy conditions and during rains, when flies apparently looked for shelter.
Only 29.5% of flies were observed to land directly on the stickum. Buzzing loudly,
all tried to become free. Those landing near the outer edge of the stickum generally
managed to escape by alternately moving their legs and by fanning their wings until
they reached a place free of stickum; 62.7% never escaped. Inserts nearing saturation,
with ca. 2 flies/cm2 (equivalent to some 25-35 flies/trap/day on a seven day exposure

Origin of Approach

Form of % from % from % from
approach Above Horizontal Plane Below Total

Looping 31.5 7.5 17.2 56.2
Direct 28.6 7.8 7.4** 43.8
TOTAL 60.1 15.3 24.6 100.0

(X2 = 15.8**, P<0.01)

158 Florida Entomologist 71(2) June, 1988


% of fly landings % of fly landings
on trap sides/ on trap bottom/
% of total surface % of total surface
outside inside outside inside

Closed Jackson traps 1.33 1.09 0.69 0.461
Open bottom traps 1.37 0.631 -

'Sticky surfaces

cycle), had a rapidly increasing rate of escapes. In the open bottom trap a large portion
of flies landing on the inner sticky sides managed to get free again, accounting for a
majority of escapes from stickum.
An unknown number of flies were attracted to the surroundings of the traps, but
did not approach them during the 30 min. observation period, remaining on foliage and
branches at different heights within a 1.0 m tridimensional radius. This became apparent
when one brushed against a coffee plant bearing a trap, flushing a considerable number
of flies which flew up and then settled again on nearby foliage.
Daily, beginning in the mornings, an aggregation of males apparently forming a lek
was observed on and around the trap on the underside of the foliage. Most males
interacted territorially, with arriving males challenging resident males. Only a few
times and for short periods were males seen calling on a trap. On 2 occasions females
were observed approaching a trap from the foliage. Males around the trap started
intensive wing fanning but were not able to attract the females, which left the trap
Walking and wing waving males regularly met other walking males or males that
remained motionless and mostly without calling at varying distances from the source of
trimedlure. This often led to territorial encounters similar to those observed in leks
(Zapien et al. 1983), where pheromone-calling males on the underside of a leaf are
confronted by males landing on the same leaf (territory). These encounters are short
and generally take place without physical contact. After facing briefly, one of the males
typically makes a sudden forward movement and the other male reacts immediately by
jumping away. During these encounters inside the closed Jackson trap frequently the
male jumping back fell into the stickum. In 78 of 104 male-male encounters recorded
inside a trap, one of the opponents fell into the stickum, raising the overall percentage
of observed capture during the 30 min. observation period per trap to 26.4% of approach-
ing flies.
Male territoriality, however, also had the opposite effect. Encounters on the bottom
or outer sides of the closed Jackson trap resulted in one of the opponents leaving the
trap. As flies were not marked it was not possible to distinguish returning from newly
attracted males. Most likely however, many of these flies remain in the area of attraction
and return. As interactions continued throughout the day, the cumulative daily catch
might be much higher than indicated by our 30-minute samples.


New analysis of medfly pheromone components (Baker et al. 1985) have confirmed
no structural relation with trimedlure. Our observations, however, support some recent

Villeda et al.: Medfly Behavior


speculation about trimedlure acting similarly to the male-produced medfly sex pheromone,
and therefore as an analogue to the lek-forming pheromone (Chambers 1977, Burk &
Calkins 1983, Prokopy & Roitberg 1984, Sivinski & Calkins 1986). Males are apparently
attracted to the trimedlure baited trap as they are attracted to a large male mating
aggregation or lek. Approaching males, aggregating on the surrounding foliage and the
trap, interact territorially as in leks, where pheromone calling males, apparently per-
ceived as competitors, are challenged by attracted arriving males (Zapien et al. 1983,
Hendrichs & Hendrichs, unpublished data).
Males get caught basically in two ways: either on their first arrival at the trap while
landing on the sticky surface, or once inside the trap and while participating in male-
male encounters. A larger sticky surface under the trimedlure source of the trap should
therefore catch a higher percentage of attracted flies. P. Baker (unpublished data)
comparing the larger sized Delta trap with the Jackson trap, found this to be true. Also,
by placing a trap carefully with no foliage in the immediate vicinity, more flies should
be chanelled onto or into the trap, increasing possibly the direct landings on the sticky
surface or the interactions between males inside (Drummond et al. 1984).
The poorer performance in capturing medflies of the trimedlure-baited open bottom
trap differs from the findings for A. suspense by Davis et al. (1984). Our results indicate
that medflies tend to land on upper surfaces and then tend to walk to bottom surfaces.
This however, could partly be the result of the repellency of sticky surfaces and the 60
angle of trap sides. The open bottom trap still attracted apparently more flies, but ac-
counted also for most escapes from the stickum, as a large portion of those landing on
the sticky inner side managed to get free again, possibly helped by gravity. The closed
Jackson trap was more effective in capturing them and additionally benefitted from the
agonistic encounters between males within the trap. Furthermore, the open bottom
trap is less practical and runs the risk of losing trapped flies due to dripping of the
stickum during hot weather.
The important role of trap placement is confirmed (Farias & Nakagawa 1970). Not
only is the possibility of capturing flies higher when traps are placed on fruiting hosts,
but also when they are located in dense vegetative environments. Determining where
leks form primarily in nature acquires additional importance, so that traps, which appa-
rently provide a stimulus similar to an established lek, are placed in suitable locations
(Hendrichs & Hendrichs, unpublished data).
The color effect, despite being secondary to chemical attractant, trap placement and
trap design, confirmed the trends found by Hentze (1984), supporting the notion that
yellow surfaces constitute bright foliage mimics (Prokopy and Economopoulos 1976,
Nakagawa et al. 1978, Cytrynowicz et al. 1982). According to Katsoyannos (1987),
however, the yellow preference varies with the type of background foliage. Striping,
successfully employed along with a fruit mimicking color for A. suspense (Greany et al.
1982, Davis et al. 1984), was not as attractive to medfies when used with a foliage
mimic. Jackson traps of a yellow in the 500-520nm region could be considered for
medflies (Agee et al. 1982). A considerable increase in the capture of beneficial insects,
as has been reported for the sticky yellow Rebell trap (Delrio 1986), does not seem to
occur with a yellow Jackson trap.
The principal period of pheromone calling observed for C. capitata under similar
conditions (Prokopy & Hendrichs 1979) was from 1030-1400 hrs., when males aggregate
and pheromone "chorus" from the bottom leaf surface. Significant trapping captures,
under the warmer weather of this study, extended from 0800-1800, reaching a maximum
in the afternoons. (Under Egyptian summer conditions the main capture is in the morn-
ings, however also after male lek-establishment reaches its peak; Hendrichs unpublished
data). Apparently traps compete with leks for males during the main calling period.
Also, some of the attracted males might be arrested by other calling males in an ex-

160 Florida Entomologist 71(2) June, 1988

tended lek around the trap before they come to the trap. In the afternoons fewer males
were seen calling, and fewer males might be arrested before arriving at a trap. Possibly
as important is the fact that the peak of capture is apparently delayed some time after
the peak of attraction due to the accumulation of males inside the trap before they get
caught as a result of their agonistic interactions.
The observed attractancy pattern also might be related to the existence of the two
mating modes or strategies that have been observed in medfly males (Prokopy & Hen-
drichs 1979). While aggregating in the midmornings in leks on the foliage in order to
attract females, males actively move around before and after this period, searching for
ovipositing females on the fruit, thereby increasing their possibility of entering the
influence radius of a trap. One could speculate here that trimedlure could act as a "fruit
attractant" such as alpha-copaene (Teranishi et al. 1987) and other fruit volatiles (Light
et al. 1988), males responding to it more during hours when females are likely to be
ovipositing, although polyphagous medflies seem to locate individual fruits at closer
range more through visual than olfactory cues (Feron 1962, Sanders 1968, Prokopy &
Economopoulos 1976, Nakagawa et al. 1978).
The nearly exclusive capture of males confirms a common pattern (Nakagawa et al.
1970), that is to be expected if the parapheromone trimedlure acts analogously to the
medfly sex pheromone (Burk & Calkins 1983). Females generally are attracted only a
few times in their life to leks, whereas males do so daily to increase their mating success
(Zapien et al. 1982, Hendrichs & Hendrichs, unpublished data). Females, mostly those
that are sperm deficient, are therefore only regularly caught when males are scarce or
population density is low (Nakagawa et al. 1981). Fitt (1981), reported similar patterns
of response for female Dacinae, establishing a direct competition for females between
the synthetic releasing trap and the wild male pheromone. Furthermore, females ap-
proaching a lek (or trap) respond at close range more to sound and visual stimuli of
individual males (Nakagawa et al. 1981, Prokopy & Roitberg 1984), being mostly ar-
rested before nearing a trap. The observation of pair formations within a 1 meter radius
around the trap (Hendrichs, unpublished data) supports this.
Recently, Drew (1987) has pointed out that this pattern of attraction, at least in
Dacinae, is the result of the lure acting as a feeding attractant for females and a sex
attractant for males. Unlike Dacus flies, however, that feed vigorously on cuelure or
methyleugenol, C. capitata flies attracted to trimedlure were not observed feeding on
the lure, (although Nadel & Peleg 1965, found that not only virgin but also starved
females respond to trimedlure and that their response is proportional to their degree
of starvation; and McInnis & Warthen 1988, report males feeding on trimedlure).
One general implication for medfly survey programs under similar conditions is that
trap catches during the rainy season, which has rainy afternoons daily, will be based
on the lower morning catches and are therefore not comparable to dry season trap data
(Hendrichs et al. 1982). Fruit sampling is an alternative survey and detection method
without some of the disadvantages of the present trapping system (including the dif-
ferentiation of sterile and wild adults), and that samples a much larger larval population
(Carey 1982). Fruit sampling as a means of detection should be given more importance,
mainly during the rainy seasons and in areas where sterile flies are being released.


We wish to express our special thanks to Fernando Tapia for his field support, and
to Carlos A. Bustamante the owner of the plantation. Rafael Mata assisted with logis-
tical support. Moscamed Program Guatemala provided valuable information on popula-
tion levels prior to this work. Jorge Guillen assisted in the dissection of gonads. We
also greatly acknowledge the critical reviews of an earlier draft by Drs. Peter Baker,
Carrol Calkins, Ring Carde, Daniel Papaj and Ronald Prokopy.

Villeda et al.: Medfly Behavior 161


Spectral sensitivities and visual attractant studies on the Mediterranean fruit fly
C. capitata, olive fly D. oleae, and the European cherry fruit fly R. cerasi.
(Diptera: Tephritidae). Z. ang. Entomol. 93: 403-12.
BAKER, R., R. H. HERBERT, AND G. G. GRANT. 1985. Isolation and identification
of the sex pheromone of the Mediterranean fruit fly Ceratitis capitata (Wied.).
J. Chem. Soc., Chem. Commun. 73: 824-825.
1961. New attractants for the Mediterranan fruit fly. J. Agr. Food Chem. 9:
BURK, T. AND C. 0. CALKINS. 1983. Medfly mating behavior and control strategies.
Florida Entomol. 66: 3-18.
CAREY, J. R. 1982. Demography and population dynamics of the Mediterranean fruit
fly. Ecol. Modelling 16: 125-150.
CHAMBERS, D. L. 1977. Attractants for fruit fly survey and control. pp. 327-344. In:
Chemical control of insect behavior: theory and applications. H. H. Shorey and
J. J. McKelvey, Jr. (eds.) John Wiley & Sons, N.Y.
CYTRYNOWICZ, M., J. S. MORGANTE, AND H. M. L. DE SOUZA. 1982. Visual re-
sponses of South American fruit flies Anastrephafraterculus, and Mediterranean
fruit flies, Ceratitis capitata, to colored rectangles and spheres. Environ. En-
tomol. 11: 1202-1210.
DAVIS, J. C., H. R. AGEE, AND D. C. CHAMBERS. 1984. Trap features that promote
capture of the Caribbean fruit fly Anastrepha suspense. J. Agric. Entomol. 1:
DELRIO, G. 1986. Biotechnical methods for Ceratitis capitata Wied. pp. 11-21. In:
Fruit Flies of Economic Importance 1984. R. Cavalloro (ed.). Balkema, Rotter-
DREW, R. A. I. 1987. Behavioural strategies of fruit flies of the genus Dacus (Diptera:
Tephritidae) significant in mating and host plant relationships. Bull. ent. Res.
77: 73-81.
DRESNER, E. 1970. A sticky trap for Mediterranean fruit fly survey. J. Econ. Ent.
63: 1813-6.
DRUMMOND, F., E. GRODEN, AND R. J. PROKOPY. 1984. Comparative efficacy and
optimal positioning of traps for monitoring apple maggot flies (Diptera: Te-
phritidae). Environ. Entomol. 13: 232-235.
FARIAS, G. J. AND S. NAKAGAWA. 1970. Host vs. nonhost plants as sites for baited
traps for Mediterranean fruit flies. J. Econ. Ent. 63: 662-3.
FERON, M. 1962. L'instinct de reproduction chez la mouche mediterranean des fruits
Ceratitis capitata Wied. (Dipt. Trypetidae). Comportement sexuel. Comporte-
ment de ponte. Rev. Path. veg. Entomol. Agric. Fran. 41: 1-129.
FITT, G. P. 1981. Responses by female Dacinae to "male" lures and their relationship
to patterns of mating behaviour and pheromone response. Ent. & exp. app. 29:
GREANY, P. D., A. K. BURDITT, JR., AND D. L. CHAMBERS. 1982. Effectiveness
of Jackson traps for fruit flies improved by addition of colored patterns. Florida
Entomol. 65: 374.
GUILLEN, J. 1983. Manual for the differentiation of wild (fertile) Mediterranean fruit
flies, from irradiated (sterile) ones. Program Moscamed, D.G.S.V.-S.A.R.H.
Talleres Graficos de la Naci6n, M6xico, D.F. M6xico. 102 pp.
GUTIERREZ, S. 1972. Test of traps for detecting the Mediterranean fruit fly Ceratitis
capitata. Folia Entomologica Mexicana 23-24: 58-59.
HENDRICHS, J., G. ORTIZ, P. LIEDO, AND A. SCHWARZ. 1983. Six years of success-
ful medfly programme in Mexico and Guatemala. pp. 353-365. In: Fruit Flies of
Economic Importance. R. Cavalloro (ed.) Balkema, Rotterdam.
HENTZE, F. 1984. Efecto del color en la atracci6n de mosca del Mediterraneo. pp.
390-6. In: Memorias del II Congreso Nacional de Manejo Integrado de Plagas.

162 Florida Entomologist 71(2) June, 1988

KATSOYANNOS, B. I. Some factors affecting field responses of Mediterranean fruitflies
to colored spheres of different sizes, pp. 469-473. In: Fruit Flies. A. P.
Economopoulos (ed.) Elsevier Science, Amsterdam.
LIGHT, D. M., E. B. JANG, AND R. A. FLATH. 1988. Electroantennogram responses
of the Mediterranean fruit fly, Ceratitis capitata, to the volatile constituents of
nectarines. J. Chem. Ecol. (in press).
NADEL, D. J. AND B. A. PELEG. 1965. The attraction of fed and starved males and
females of the Mediterranean fruit fly, Ceratitis capitata, to trimedlure. Isr. J.
Agr. Res. 15: 33-86.
MCINNIS, D. O. AND J. D. WARTHEN, Jr. 1988. Mediterranean Fruit Fly (Diptera:
Tephritidae): Attraction of males to leaf or stem exudates of Ficus and Litchi
trees. J. Econ. Entomol. (in press).
NAKAGAWA, S., G. J. FARIAS, AND L. F. STEINER. 1970. Response of female
Mediterranean fruit flies to male lures in the relative absence of males. J. Econ.
Ent. 63: 227-9.
T. URAGO, AND E. J. HARRIS. 1978. Visual orientation of Ceratitis capitata
flies to fruit models. Ent. exp. & appl. 24: 193-198.
NAKAGAWA, S., L. F. STEINER, AND G. J. FARIAS. 1981. Response of virgin female
Mediterranean fruit flies to live mature normal males, sterile males and trimed-
lure in plastic traps. J. Econ. Ent. 74: 566-567.
ORTIZ, G. AND J. REYES. 1982. An ecological and logistic study on eradication of the
Mediterranean fruit fly from Egypt. Intl. Atomic Energy Agency, Vienna. 55 pp.
ORTIZ, G., L. GARCIA, AND J. REYES. 1983. Evaluaci6n de tres diferentes trampas
para capturar Ceratitis capitata. XVIII Congr. Nac. de Entomologia. Tapachula,
PROKOPY, R. J. AND A. P. ECONOMOPOULOS. 1975. Attraction of laboratory-cul-
tured and wild Dacus oleae flies to sticky-coated McPhail traps of different colors
and odors. Environ. Entomol. 4: 187-192.
PROKOPY, R. J. AND A. P. ECONOMOPOULOS. 1976. Color responses of Ceratitis
capitata flies. Z. ang. Ent. 80: 437-7.
PROKOPY, R. J. AND J. HENDRICHS. 1979. Mating behavior of Ceratitis capitata on
a field caged host tree. Ann. Entomol. Soc. Am. 72: 642-648.
PROKOPY, R. J. AND B. ROITBERG. 1984. Foraging behavior of true flies. Am. Scien-
tist 72: 41-9.
SANDERS, W. 1968. Die Eiablage der Mittelmeerfruchtfliege Ceratitis capitata Wied.
Ihre Abhiingigkeit von Farbe und Gliederung des Umfeldes. Z. Tierpsycho. 19:
SIVINSKI, J. M. AND C. O. CALKINS. 1986. Pheromones and parapheromones in the
control of Tephritids. Florida Entomol. 69: 157-168.
GHAM, AND S. GOTHILF. 1987. Recent developments in chemical attractants
for tephritid fruit flies. Amer. Chem. Society Symp. No. 330, Chapt. 38: 431-438.
ZAPIEN, G., J. HENDRICHS, P. LIEDO, and A. CISNEROS. 1983. Comparative mating
behavior of wild and mass reared sterile medfly Ceratitis capitata (Wied.) on a
field caged host tree. II. Female mate choice, pp. 397-407. In: Fruit Flies of
Economic Importance. R. Cavalloro (ed.) Balkema, Rotterdam.
ZAPIEN, G., J. HENDRICHS, M. ALUJA, AND R. MATA. 1983. Comparative mating
behavior of wild and mass reared sterile medflies on a field caged host tree. I.
Male territoriality and lekking behavior. XVIII Congr. Nac. de Entomologia,
Tapachula, Mexico.

Deyrup et al.: Florida Ants



Archbold Biological Station
P. O. Box 2057
Lake Placid, FL 33852

Museum of Comparative Zoology
Harvard University
Cambridge, MA 02138

Department of Entomology and Nematology
University of Florida
Gainesville, FL 32611

Department of Biology
Carleton University
Ottawa, K1S 5BC


A new survey of the ants of the Florida Keys increases the known fauna from 30 to
83 species. An annotated list provides data on habitats, collection sites, and location of
vouchers. Solenopsis corticalis Forel, Leptothorax torrei (Aguayo) and Monomorium
ebeninum Forel are new records for the U.S. The fauna includes 27 exotics and 31
species native to the southeastern Coastal Plain; most of the remaining species are
Antillean. There are 2 possibly endemic species. The proportion of known exotics (33%)
in the fauna is the highest for any area in the U.S. There is evidence that populations
of certain exotics are increasing with increasing disturbance of remaining native
habitats. Species diversity in the Keys is probably limited by 1) limited habitat diver-
sity, 2) lack of easy access to the rich ant fauna of the Neotropics, and 3) unsuitability
of the climate and habitats for many species found farther north.


Un muestreo nuevo de las hormigas de los Cayos de la Florida, aument6 el co-
nocimiento de la fauna de 30 a 83 species. Una anotada lista provee datos sobre la
habitaci6n, lugares de colecta, y la localidad de los testigos. Solenopsis corticalis Forel,
Leptothorax torrei (Aguayo), y Monomorium ebeninum Forel, son nuevos records para
los Estados Unidos. La fauna incluye 27 species ex6ticas y 31 nativas a los Llanos
Costales del sudeste; la mayoria del resto de las species son Antillanas. Posiblemente
2 species sean end6micas. La proporci6n de species ex6ticas conocidas (33%) de la
fauna, es la mAs alta que la de cualquier Area de los Estados Unidos. Hay evidencia que
poblaciones de ciertas species ex6ticas estAn aumentado con el aumento de disturbios
de la habitaci6n native que queda. Las diversidad de species en los Cayos estA probab-
lemente limitada por: 1) diversidad limitada de la habitaci6n, 2) falta de acceso fAcil a
la rica fauna de hormigas del Neotr6pico, y 3) lo poco convenient del clima y habitaci6n
para muchas species que se encuentran mAs al norte.

164 Florida Entomologist 71(2) June, 1988

In 1958 E. O. Wilson surveyed the ants of the Florida Keys; the resulting list was
published in 1964, along with notes on the geographic origin and distribution of the
species. In the ensuing 30 years there have been extensive changes in the habitats of
southern Florida, probably affecting the ant fauna. There have also been modifications
in our taxonomic concepts of some species. Additionally, more intensive collecting
methods, as well as possible influxes of additional species, have more than doubled the
number of species known from the Keys. A current survey of the ants of the entire
state now provides a better perspective on the biogeography of the ants of the Keys.
For these reasons, it seems useful to review the ant fauna.
One purpose of this paper, therefore, is to provide an annotated list of species that
can be compared with the earlier study, and that will be a baseline for future documen-
tation of the accelerating disruptions of the ecosystems of tropical Florida. From an
ecological standpoint the ants are an excellent group to monitor because they are so
dominant in terrestrial tropical ecosystems (Brues 1952). One may say that, as go the
ants, so go the arthropods.
A second goal is to examine the biogeography of the ants of the Florida Keys. The
diversity and derivation of the fauna is now open to new interpretations. The distribu-
tion of exotics and the proportion of exotics in the fauna can be reviewed. The presence
or absence of species can often be correlated with habitat and climate, now that we
know more about the ecology of Florida ants. We should even be able to make a few
predictions about the findings of the next survey, which we might consider scheduling
for the year 2018.


Ants were collected on numerous trips to the Florida Keys during 1982-87, and on
trips made by our colleagues listed below. Specimens were collected by simple searching
techniques and by extracting ants from litter samples by standard modified Berlese
funnels. A few species were collected in Malaise traps. Voucher specimens are deposited
in the following collections, abbreviated in the species list: Archbold Biological Station,
Lake Placid, FL (ABS), Museum of Comparative Zoology, Harvard, Cambridge, MA
(MCZ), collections of Norman Carlin (NC), Gary Umphrey (GU), James Trager (JT).
The additional collectors whose initials are in the annotated list are Dr. Edward Wilson
(Harvard Museum of Comparative Zoology, Cambridge, MA), Dr. James Wolfe and
David Smith (Archbold Biological Station), Alan Herndon (Everglades National Park,
Homestead, FL), Marc Minno (University of Florida, Gainesville), Chester Winegarner
(DeFuniak Springs, FL), Dr. Stewart Peck and Dr. Jarmila Kukalova-Peck (Carleton
University, Ottawa, ONT), Dr. Stewart Peck and Dr. Jan Klimaszewski (the latter
collector from Petawawa, ONT). Some records of species of Pseudomyrmex are taken
from Ward, 1985, abbreviated (PW). Specimens were identified by the authors.
Since some of the biogeographical discussion is dependent on the geographic origin
of the ant populations in the Keys, we explain our presumptions on the provenance of
each species in the annotated list. Statements about the presence of a species in other
parts of Florida are based on our own unpublished collecting records unless otherwise
attributed. Our understanding of the taxonomy of the ants of the Keys also depends to
some extent on unpublished work. We briefly mention taxonomic ambiguities, but avoid
taxonomic innovations in this paper.
The small size and great dispersion of the islands of the Florida Keys prevents
preparation of a compact map of our collecting sites. We refer the reader to the Florida
Atlas and Gazeteer, available from the DeLorme Publishing Co., P. 0. Box 298,
Freeport, Maine 04032.

Deyrup et al.: Florida Ants


Amblyopone pallipes (Haldeman). Males of this native species were collected in
Malaise traps on Big Pine (S&JP), Sugarloaf (S&JP), Cudjoe Key (NC). This species is
usually found in upland forested areas. Vouchers: NC, GU.
Aphaenogasterflemingi Smith. This species is widely distributed in upland habitats
throughout the southeastern U.S. There are only a few records from southern Florida,
probably because in south Florida upland habitats are widely scattered and much of
their native vegetation eradicated. This species is native to Florida. The Keys popula-
tion is probably strongly isolated. Habitat in the Keys is dry open pineland, now found
only on Big Pine. Big Pine (GU). Vouchers: GU
Aphaenogaster miamiana Wheeler. This form is a member of a complex of several
named species and subspecies that require further study (Carroll 1975). The form known
as A. miamiana occurs throughout southern and central Florida. This species is native.
The habitat in the Keys is tropical hammocks; nests are usually in rotten wood on the
ground. Elliott (MD), Largo (EW, MD), Upper Matecumbe (MD), Big Pine (MD, GU,
S&JP), Big Torch (S&JP), Middle Torch (S&JP). Vouchers: ABS, GU.
Brachymyrmex, sp. nr. depilis Emery. It is probable that two species of yellow
Brachymyrmex have been combined under the name depilis in Florida; the genus is in
urgent need of revision. The Keys form occurs throughout southern and central Florida,
and is almost certainly native. Habitat in the Keys is rotten wood and deep leaf mold
in shaded situations. Elliott (MM, MD), Adams (MM), Totten (MM), Largo (JT, MD),
Plantation (DS, MD), Grassy (JT, MD) Long (MD), Big Pine (GU, JT, MD), No Name
(JT, MD, S&JP), Middle Torch (S&JP), Big Torch (S&JP). Vouchers: ABS, GU, JT.
Brachymyrmex, sp. nr. obscurior Forel. It is probable that two or more species
have been combined under the name obscurior in Florida. The Keys form occurs
throughout Florida. It has a strong preference for open, grassy, disturbed areas such
as roadsides and sparse lawns, but is also found in naturally disturbed areas, particularly
beaches. We have no good reason to consider this species exotic, though its populations,
if not its geographic range, must have increased with the recent increase in disturbed
habitats. Nests are in soil. Largo (MD), Bahia Honda (JT, MD), Big l,ine (MD), M!iddle
Torch (S&JP), Stock Island (MD), Key West (NC, JW, MD). Vouchers ABS, JT, GU.
Brachymyrmex sp. Three microgyne queens were collected on No Name (S&JP),
4-5 June 1986, by car netting. These queens show characteristics of social parasites (E.
O. Wilson, personal communication), though no social parasites are known in the genus
Brachymyrmex. Large numbers of males and queens of Brachymyrmex sp. nr. depilis
were taken in the same samples. Vouchers: GU.
Camponotus abdominalis floridanus (Buckley). Some myrmecologists consider this
form a distinct species, C. floridanus, others do not consider it merits subspecific status.
This native species occurs throughout Florida. The habitat in the Keys is dead wood
and trash piles found along the beach, in the margins of hammocks, and in rocky pine-
land. Elliott (MD, NC), Largo (EW, NC, MD), Plantation (EW, MD), Upper
Matecumbe (NC, MD), Indian Key Fill (GU), Grassy (JT, MD), Bahia Honda (MD,
S&JP), No Name (JT, MD), Big Pine (NC, EW, JT, GU, MD). Ramrod (NC), Sugarloaf
(NC, GU), Shark (NC), Saddlebunch 1 (NC), Saddlebunch 2 (JW), Saddlebunch 5 (NC),
Big Coppitt (JW), Boca Chica (JW), Stock Island (MD), Key West (NC, MD). Vouchers:
Camponotus decipiens Emery. This native arboreal species has usually gone under
the names C. rasilis Wheeler or C. sayi Emery. We follow Snelling (in press) in the
use of the name C. decipiens. It is widely distributed in Florida, but generally absent
in the extreme south, possibly because its habitat is dominated by C. planatus. It
occurs on Key West, where C. planatus is uncommon. Vouchers: ABS.

Florida Entomologist 71(2)

Camponotus impressus (Roger). This species is native to Florida and occurs
throughout the state. Habitat is hollow twigs and weed stems. This species appears to
be rather scarce in the Keys, and is found primarily in large red mangroves (Cole 1983).
Big Pine (EW). Vouchers: MCZ.
Camponotus inaequalis Roger. We have two collections that appear to represent
this Caribbean species. This species is rather similar to C. tortuganus, and the entire
southern C. maculatus complex could use revision. Accordingly, we do not formally
add this species to the U.S. fauna. One colony was found in a rotten sea grape branch
about 1.5 m above the ground. Bahia Honda (GU), Key West (NC). Vouchers: GU, NC.
Camponotus planatus Roger. In Florida this tropical Caribbean species is restricted
to the extreme southern portion of the state. We consider this is probably a native
species. The habitat is hollow twigs, old termite galleries in dead wood, and occasionally
in grass culms. This is the dominant ant of Florida's tropical hammocks. Elliott (MD),
Largo (EW, MD), Upper Matecumbe (MD), Long (CU), Grassy (JT, MD), Big Pine
(NC, JT, GU, MD), No Name (MD, S&JP), Middle Torch (S&JP), Sugarloaf (S&JP),
Stock Island (MD), Key West (EW). Vouchers: ABS, NC, JT, GU.
Camponotus tortuganus Emery. This native species is found through the southern
third of inland Florida, and further north along the coasts. The natural habitat is dead
wood or trash piles in open areas. Nests are often in wall voids of houses. Elliott (MD),
Plantation (EW, MD), Bahia Honda (GU), Big Pine (EW, GU, MD), No Name (JT, MD,
S&JP), Middle Torch (GU), Sugarloaf (NC), Shark (NC), Saddlebunch 5 (NC), Big
Coppitt (JW), Key West (NC, JW, MD). Vouchers: ABS, JT, NC, GU.
Cardiocondyla emeryi Forel. An exotic species found throughout Florida. Nests are
in soil in open grassy areas. Elliott (MD), Largo (MD), Plantation (EW), Upper
Matecumbe (MD), Bahia Honda (MD), Big Pine (MD, S&JP), Sugarloaf (MD), Key
West (EW, MD), Dry Tortugas (CW). Vouchers: ABS, GU.
Cardiocondyla nuda (Mayr). An exotic species found throughout Florida. Nests are
in soil in open grassy areas. Elliott (MD), Largo (MD), Upper Matecumbe (MD), Vaca
(K&P), Big Pine (MD, K&P), Sugarloaf (MD), Key West (MD). Vouchers: ABS, GU.
Cardiocondyla venustula Wheeler. An exotic species occurring sporadically
throughout Florida. Nests are in soil in open grassy areas. Upper Matecumbe (MD),
Bahia Honda (JT, MD). Vouchers: ABS, JT.
Cardiocondyla wroughtonii (Forel). An exotic species found throughout Florida.
Nests are in hollow twigs and branches. Key West (MD). Vouchers: ABS.
Conomyrma bureni Trager. A native species found throughout Florida. Nests are
in open grassy areas, including beach dunes. Elliott (MD, NC), Largo (MD), Bahia
Honda (MD), Big Pine (EW, JT, NC, GU, MD), Stock Island (MD), Key West (EW,
NC, MD). Vouchers: ABS, JT, NC, GU.
Crematogaster ashmeadi Mayr. This native southeastern species occurs throughout
Florida. Nests are in dead tree limbs, hollow twigs and in weed stems. Elliott (MD),
Largo (EW, MD), Plantation (EW), Lower Matecumbe (MD), Grassy (JT, MD), Fat
Deer (GU), Ohio (GU), Bahia Honda (MD), Big Pine (EW, JT, GU, MD), No Name
(JT), Key West (EW, JW). Vouchers: ABS, JT, GU.
Crematogaster atkinsoni Wheeler. A widespread native southeastern species found
in coastal areas. The carton nests are usually in grass tussocks. Big Pine (NC, MD),
Saddlebunch 2 (JW). Vouchers: ABS, NC.
Crematogaster minutissima Mayr. This native southeastern species occurs through-
out Florida, but appears to be scarcer in the southern part of the state. Nests are in
deep humus. Elliott (MD), Big Pine (K&P). Vouchers: ABS, GU.
Cyphomyrmex minutus Mayr. There are two species of Cyphomyrmex in Florida,
C. minutus in southern and central Florida, and C. rimosus Forel, in northern Florida
and adjoining states. The distributions of the two species overlap in central Florida.

June, 1988

Deyrup et al.: Florida Ants

According to Snelling (personal communication), the species found in south Florida and
the Keys is the West Indian species C. minutus. C. rimosus is probably recently
introduced, as there is no northern Cyphomyrmex mentioned by Creighton (1950). As
far as we know, C. minutus is native in the Keys. Nests are in the soil, usually under
rocks or logs. Elliott (MD, NC), Largo (EW, MD), Plantation (EW, MD), Upper
Matecumbe (MD), Bahia Honda (MD, GU, K&P), Big Pine (JT, NC, K&P, MD), No
Name (JT, MD), Big Torch (S&JP), Summerland (K&P), Sugarloaf (MD, S&JP),
Saddlebunch 2 (JW), Saddlebunch 5 (NC), Boca Chica (JW), Stock Island (MD, K&P),
Key West (MD). Vouchers: ABS, NC, JT, GU.
Discothyrea testacea Roger. This native southeastern species occurs throughout
Florida. Nests are in humus in wooded areas. A single specimen was collected from No
Name (MD). Voucher: ABS.
Eurhopalothrix floridana Brown and Kempf. The distribution of this species ex-
tends from the Keys into northern Florida. All other Eurhopalothrix are neotropical,
and E. floridana itself has been collected in Mexico (W. L. Brown, 1965, personal
communication). This species was not discovered until 1960 (Brown & Kempf), and was
not known to be widely distributed until we began our survey of Florida ants, but it is
sufficiently cryptic to have avoided notice for a long time. This is shown by a damaged
specimen, discovered by Dr. David R. Smith, in the Pergande collection (USNM); the
specimen was collected in Key West in 1887. E. floridana is one of a number of species
that we will treat as native even though its relationships and habits make an exotic
origin plausible. Largo (MD), Upper Matecumbe (MD), Big Pine (K&P), No Name
(MD), Key West (Pergande). Vouchers: ABS, GU, U.S. National Museum.
Forelius pruinosus (Roger). This species has been recently transferred from the
Genus Iridomyrmex (Snellling & Wheeler 1979). The species known as F. pruinosus
shows considerable variation and may be a species complex. F. pruinosus is native in
south Florida. The habitat of this species is beaches and dry rocky or sandy open areas.
Elliott (MD, NC), Largo (MD), Lower Matecumbe (GU), Grassy (MD), Bahia Honda
(MD), Big Pine (NC, JT, GU, MD), No Name (JT), Saddlebunch 2 (JW), Boca Chica
(JW), Key West (NC). Vouchers: ABS, JT, NC, GU.
Hypoponera inexorata (Wheeler). This species has a general austral distribution in
North America, and is probably native. Its habitat in the Keys is tropical hammocks.
Largo (MD), Bahia Honda (MD), Big Pine (GU), Big Torch (P&K), Sugarloaf (MD),
West Summerland (K&P). Vouchers: ABS, GU.
Hypoponera opaciceps (Mayr). This species has an austral general distribution in
North America and occurs through South America (Smith 1979). This species generally
occurs in moist disturbed habitats, including accumulations of beach wrack. Largo (MD),
Long (GU), Big Pine (MD), No Name (MD), West Summerland (GU), Stock Island
(K&P), Dry Tortugas (CW). Vouchers: ABS, GU.
Hypoponera opacior (Forel). A widespread species native in the southern United
States. Nests are in soil and rotten wood in mesic and xeric forests. Elliott (MD), Totten
(MM), Largo (JT, MD), Plantation (MD), Upper Matecumbe (MD), Grassy (JT, MD),
Big Pine (MD, K&P), Sugarloaf (S&JP). Vouchers: ABS, JT, GU.
Hypoponera punctatissima (Roger). An exotic species found throughout Florida.
Its habitat is accumulations of organic matter in moist places. Largo (MD), Big Pine
(MD, K&P), Saddlebunch 2 (JW), Stock Island (MD), Key West (JW). Vouchers: ABS,
Leptothorax allardycei (Mann). This species was known as Macromischa floridana
(Wheeler) until Baroni Urbani (1978) revised the subgenus Macromischa. It is also
known from the Bahamas. We consider this as probably a native species. The habitat
is dead vines in dense tropical hammock. This species may be nocturnal. Elliott (MD),
Largo (MD), Big Pine (S&JP), Big Torch (S&JP), Sugarloaf (S&JP). Vouchers: ABS,

Florida Entomologist 71(2)

June, 1988

Leptothorax torrei (Aguayo). This species is known only from Cuba (Baroni Urbani
1978) and from the Florida Keys (new U.S. record). It is a small inconspicuous ant, not
likely to be collected except by Berlese extraction, and its recent discovery in the Keys
need not indicate this species was recently introduced. Largo (MD), Big Pine (GU, MD,
K&P), No Name (MD), Key West (GU). Vouchers: ABS, GU, MCZ.
Leptothorax (Dichothorax) sp. nr. pergandei. This species appears closely related to
L. pergandei Emery, but differs from south Florida populations in color and in the shape
of the postpetiole. The habitat, a salt marsh, is also unlike that of pergandei, which
inhabits dry, sparse woodlands. We provisionally consider this a native undescribed
species. The habitat of this species, as far as it is known, is a salt marsh with tussocks
of Sporabilis grass. Largo (MD) and Big Pine (GU). Vouchers: ABS, GU.
Monomorium destructor (Jerdon). This exotic species is not common in Florida, and
may be on the decline in the Keys, except for Key West, where it is a dominant urban
ant. The habitat is disturbed areas around buildings. Largo (EW), Plantation (EW),
Key West (NC, GU, MD). Vouchers: ABS, GU.
Monomorium ebeninum Forel. This West Indian species has been collected only
once in the Keys (new U.S. record). We consider this species a poorly established
exotic. The habitat was the edge of a tropical hammock. Upper Matecumbe (MD).
Vouchers: ABS.
Monomorium floricola (Jerdon). An exotic species found throughout southern
Florida. Nests are in dead twigs and vines, usually in disturbed areas. Elliott (MD),
Largo (EW, MD), Plantation (EW), Indian Key Fill (GU), Grassy (MD), Bahia Honda
(MD), Big Pine (MD, GU), No Name (MD), Mliddle Torch (S&JP), Big Torch (GU),
Sugarloaf (MD, GU), Stock Island (MD), Key West (MD), Dry Tortugas (CW). Vou-
chers: ABS, GU.
Monomorium pharaonis (Linnaeus). An exotic species found in and around buildings
in urban areas throughout Florida. Elliott (NC), Plantation (EW), Big Pine (MD), Vou-
chers: ABS.
Myrmecina americana Emery. This native species is found through much of North
America, usually in wooded areas. In the Keys specimens have been collected in ham-
mocks adjacent to the beach. Elliott (MD), Plantation (MD), Big Pine (S&JP), No Name
(S&JP), Middle Torch (S&JP), Big Torch (S&JP), Sugarloaf (S&JP). Vouchers: ABS,
Neivamyrmex opacithorax (Emery). This native species is widely distributed in the
southern U.S. It usually occurs in upland wooded areas. Bahia Honda (Phillip Ward
collector) Big Pine (S&JP), Middle Torch (S&JP), Cudjoe (S&JP), Key West (J. Bunch
collector). Vouchers: GU, JT.
Odontomachus brunneus (Patton). This appears to be a native southeastern species.
Nests are under rocks and in rotten wood in moist wooded areas. Largo (MD), No Name
(MD, GU, S&JP). Vouchers: ABS, GU.
Odontomachus ruginodis Wheeler. This species is found through southern Florida,
but is more common in coastal areas. 0. ruginodis appears to be a West Indian species
which we tentatively consider native in the Keys. Nests occur in soil and under stones,
usually in moderately open areas. Elliott (NC, MD), Largo (EW, JT, MD), Plantation
(MD), Upper Matecumbe (MD), Lower Matecumbe (MD), Grassy (JT, MD), Fat Deer
(GU), Bahia Honda (MD, GU), Big Pine (EW, JT, NC, GU, S&JP, MD), No Name
(S&JP, MD), Middle Torch (S&JP), Big Torch (S&JP), Sugarloaf (MD), Saddlebunch 1
(NC), Saddlebunch 2 (JW), Saddlebunch 5 (ND), Boca Chica (JW), Stock Island (S&JP),
Key West (JW, NC, MD). Vouchers: ABS, JT, NC, GU.
Pachycondyla stigma (Fabricius). This pantropical species seems to be extraordinar-
ily mobile, using either natural or man-assisted methods, or both. Until we have a
clearer idea of how P. stigma moves about, we are treating it as a Florida native. The


Deyrup et al.: Florida Ants

principle habitat appears to be the rotten bark of dead trees, usually in relatively open
sites. A single collection has been made in the Keys, on Elliott Key (MD). Vouchers:
Paratrechina bourbonica (Forel). This exotic species occurs throughout Florida,
usually in disturbed habitats around buildings. Beach and mangrove areas, which are
naturally disturbed habitats, are also highly suitable. Elliott (MD), Largo (MD), Upper
Matecumbe (MD), Grassy (MD), Big Pine (MD), No Name (JT, MD), Saddlebunch 5
(NC), Shark (NC), Boca Clca (JW), Key West (EW, JW, MD), Dry Tortugas (CW).
Vouchers: ABS, NC, JT.
Paratrechina concinna Trager. This native species occurs throughout Florida, usu-
ally in rotten wood in or near swamps or near swamps or marshes. A single collection
was made from a rotten palm stump in a rocky brackish swamp on Key Largo (MD).
Vouchers: ABS.
Paratrechina guatemalensis (Forel). This apparently exotic species seems to be
expanding its range in southern Florida. P. guatemalensis, unlike the other exotic
Paratrechina of the Keys, is at home in densely wooded areas, and is a dominant ant
of some tropical hammocks. Largo (MD), Plantation (MD), Bahia Honda (MD), Big Pine
(JT, MD), No Name (MD), Sugarloaf (MD), Stock Island (MD), Dry Tortugas (NC).
Vouchers: ABS, JT, NC.
Paratrechina longicornis (Latreille). An exotic species found throughout Florida
around buildings, in disturbed areas and on beaches. On Big Pine tl:is species occurs in
open pinelands. Elliott (MD), Largo (EW, JT, MD), Plantation (EW, MD), Upper
Matecumbe (MD), Indian Key Fill (GU), Grassy (MD), Bahia Honda (JT, MD), Big Pine
(NC, JT, GU, MD), No Name (JT, S&JP, MD), Sugarloaf (S&JP), Saddlebunch 2 (JW),
Big Coppitt (JW), Stock Island (MD), Key West (NC, JT, MD), Dry Tortugas (CW,
NC). Vouchers: ABS, JT, NC, GU.
Paratrechina wojciki Trager. This native species inhabits upland forests throughout
Florida. Elliott (MD) and No Name (MD). Vouchers: ABS.
Pheidole dentata Mayr. This native species occurs in many habitats throughout
Florida. Nests are in soil or in rotten wood. Elliott (MD), Largo (MD), Bahia Honda
(MD), Big Pine (MD, GU, JT), No Name (JT, MD), Sugarloaf (S&JP), Saddlebunch 2
(JW), Shark (NC), Stock Island (MD), Dry Tortugas (CW). Vouchers: ABS, JT, NC,
Pheidole dentigula M. R. Snth. This native species is found in mesic sites throughout
Florida. Nests are in moist soil or rotten wood. Largo (MD), Grassy (MD), No Name
(MD). Vouchers: ABS.
Pheidole floridana Emery. There is some confusion about the application of the
names P. floridana, P. flavens Roger, and P. anastasii Emery. A publication on the
Pheidole of Florida (Naves 1985) has done little to correct tls situation. Traditionally
(Creighton 1950, Smith 1979) the name P. floridana has been applied to a widespread
upland species that has a distinctive matte area on the base of the first gastral tergite
and very evenly rugose head. This is the species that we report from the Keys. Its
habitat is dry partially shaded areas; nests are usually in soil. Elliott (MD, NC), Largo
(MD), Plantation (DS), Grassy (MD, JT), Big Pine (EW, GU, MD), No Name (MD),
Sugarloaf (MD, S&JP), Middle Torch (S&JP), Big Torch (S&JP), Saddlebunch 2 (JW),
Boca Chica (JW), Stock Island (K&P). Vouchers: ABS, GU.
Pheidole megacephala (Fabricius). In Florida this exotic species is usually found in
disturbed areas near buildings. Nests are in the soil and under objects on the ground.
Largo (MD), Big Pine (JT), Key West (NC, MD), Dry Tortugas (NC). Vouchers: ABS,
Pheidole moerens Wheeler. This exotic species nests in moist sites in disturbed
areas and in mesic woodlands. Plantation (DS, MD), Bahia Honda (MD), Big Pine (JT,
MD), Stock Island (MD), Key West (MD). Vouchers: ABS, JT.

Florida Entomologist 71(2)

Platythyrea punctata (Smith). This neotropical species is widely distributed in the
southern third of Florida, but is seldom abundant. We consider this a native species.
Nests are usually in dead wood in forested areas. Largo (EW), Vaca (GU), Big Pine
(JT, GU, S&JP), No Name (JT, S&JP), Middle Torch (S&JP), Sugarloaf (S&JP), Stock
Island (MD). Vouchers: ABS, JT, GU.
Pogonomyrmex badius (Latreille). This native harvester ant has been collected only
once on the Keys and may not be established there. P. badius is a common beach species
in both the east and west coast of Florida, and it would not be surprising if there were
a resident coastal population in the Keys. The collection site, Saddlebunch 5 (NC), is a
very long way from the mainland. Voucher: NC.
Pseudomyrmex cubaensis Forel. All the south Florida Pseudomyrmex, with the
exception of P. seminole, are neotropical species and potentially exotics, although P.
mexicanus is the only documented exotic. All species, including P. mexicanus, can be
found in natural habitats. In the absence of any clear evidence, we are considering all
these (except P. mexicanus) as Florida natives. Nests of P. cubaensis are in hollow
twigs and plant stalks. Elliott (MD), Largo (PW), Lower Matecumbe (PW), Big Pine
(JT, GU, S&JP, MD), No Name (PW), Saddlebunch 5 (NC), Key West (NC, PW, MD).
Vouchers: ABS, JT, NC, GU.
Pseudomyrmex ejectus (F. Smith). Nests are in hollow twigs and plant stalks. Elliott
(MD), Largo (MD, PW). Vouchers: ABS.
Pseudomyrmex elongatus (Mayr). Nests are in hollow twigs and plant stalks. Elliott
(MD), Largo (EW, JT, PW, MD), Plantation (EW), Upper Matecumbe (MD), Bahia
Honda (MD), Big Pine (EW, GU, S&JP, JT, PW, MD), No Name (JT, MD), Sugarloaf
(S&JP), Saddlebunch 5 (NC), Boca Chica (JW), Key West (EW, NC, MD), Dry Tortugas
(CW). Vouchers: ABS, JT, NC, GU.
Pseudomyrmex mexicanus (Roger). Nests are in hollow twigs or large plant stalks.
Elliott (MD), Largo (JT, PW, MD), Lower Matecumbe (MD, GU), Big Pine (MD, GU,
JT). Vouchers: ABS, JT, GU.
Pseudomyrmex pallidus (F. Smith). Nests are usually in plant stalks. Elliott (MD),
Largo (JT, PW, MD), Plantation (MD), Lower Matecumbe (PW), Grassy (MD), Bahia
Honda (PW), Big Pine (EW, GU, PW, MD), No Name (PW), Middle Torch (S&JP),
Cudjoe (GU, PW), Saddlebunch 5 (NC), Key West (PW). Vouchers: ABS, NC, JT, GU.
Pseudomyrmex seminole Ward. Nests are in plant stalks. Largo (PW), Bahia Honda
(MD). Vouchers: ABS.
Pseudomyrmex simplex (F. Smith). Nests are usually in hollow twigs. Elliott (MD),
Largo (MD, PW), Plantation (MD), Grassy (MD), Big Pine (MD, PW), No Name (MD,
JT), Big Torch (S&JP), Sugarloar (S&JP). Vouchers: ABS, JT, GU.
Quadristruma emmae (Emery). This exotic species is abundant throughout southern
Florida. Nests are in leaf litter, often in disturbed sites. Elliott (MD), Largo (MD),
Plantation (MD), Lower Matecumbe (MD), Long (S&JP), Bahia Honda (K&P), Big Pine
(JT, GU, K&P, MD), Big Torch (K&P), No Name (MD), Sugarloaf (JW, S&JP), Boca
Chica (JW), Stock Island (K&P). Vouchers: ABS, JT, GU.
Smithistruma dietrichi (M. R. Smith)., This widespread native species is found in
deep leaf litter and rotten wood in wooded areas. Grassy (MD), Bahia Honda (K&P),
Big Pine (K&P), No Name (MD). Vouchers: ABS, GU.
Solenopsis corticalis Forel. This tropical species might be native in extreme south
Florida; there are no published U.S. records of this species, though Florida specimens
have been available for several years. Nests are in hollow twigs and plant stems. Big
Pine (MD), No Name (JT, MD). Vouchers ABS, JT.
Solenopsis geminata (Fabricius). This widespread neotropical species is probably
native to the Keys. S. geminata usually nests in sandy soil in open areas, especially
beaches. Elliott (MM, MD, NC), Largo (JT, MD), Plantation (MD), Upper Matecumbe


June, 1988

Deyrup et al.: Florida Ants

(NC), Bahia Honda (GU, MD), Big Pine (EW, NC, JT, GU, MD), No Name (JT, MD),
Sugarloaf (MD), Shark (NC), Stock Island (MD), Dry Tortugas (CW, NC). Vouchers:
Solenopsis gobularia littoralis Creighton. This native southeastern species lives in
open sandy areas and can also be found in tussocks in salt marshes. Elliott (MD), Largo
(MD), Plantation (EW), Ohio (S&JP), Big Pine (EW, JT, GU, MD), Saddlebunch 2
(JW), Geiger (JW). Vouchers: ABS, JT, GU.
Solenopsis invicta Buren. This exotic species is not nearly as dominant in the Keys
as elsewhere in Florida, possibly because S. geminata is better adapted to coastal areas.
Nests are in soil, usually in moist unshaded sites. Elliott (MD), Saddlebunch 5 (NC),
Key West (NC). Vouchers: ABS, NC.
Solenopsis picta Emery. This native southeastern species nests in twigs, holes in
the bark of live pine trees, and plant stems. Elliott (MD), Plantation (EW), Upper
Matecumbe (EW), Bahia Honda (MD), Key West (MD). Vouchers: ABS.
Solenopsis tennesseensis M. R. Smith. This native southeastern species occurs in
soil or leaf litter in wooded areas. Elliott (MD), Largo (EW, MD), Plantation (MD),
Grassy (JT, MD), Long (MD), Bahia Honda (MD), No Name (JT, MD), Boca Chica
(JW), Stock Island (MD). Vouchers: ABS, JT.
Solenopsis sp. This small yellow species in the subgenus Diphorhoptrum is found
throughout Florida. It has not been formally described. This species is probably native
to the Keys. Nests are in leaf litter, usually in shaded habitats. Elliott (MD), Largo
(MD), Plantation (DS, MD), Lower Matecumbe (MD), Grassy (JT, MD), Bahia Honda
(JT, MD), Big Pine (JT, MD), No Name (MD), Sugarloaf (MD), Boca Chica (JW), Stock
Island (MD), Key West (JW, MD). Vouchers: ABS, JT.
Strumigenys eggersi Emery. This neotropical species is probably exotic in Florida;
it is currently found through the southern two-thirds of the state. Nests are in leaf litter
in shaded or partially shaded areas. Largo (MD), Plantation (DS, MD), Upper
Matecumbe (MD), Big Pine (MD, K&P), Sugarloaf (JW), Stock Island (K&P, MD), Key
West (JW). Vouchers: ABS, GU.
Strumigenys gundlachi (Roger). This neotropical species is probably exotic in
Florida; it is found only in the extreme southern part of the state. Nests are in leaf
litter in shaded sites. Elliott (MD), Largo (EW, MD), Upper Matecumbe (MD), Grassy
(MD), Long (MD), Vaca (K&P), Big Pine (MD, K&P), No Name (JT, MD), Sugarloaf
(MD, K&P). Vouchers: ABS, JT, GU.
Strumigenys louisianae Roger. The range of this species extends from the southern
U.S. to southern South America. It is presumably native in Florida. Nests are in leaf
litter in moist shaded areas. Largo (JT), Plantation (MD), Big l:,ine (K&P), No Name
(MD, GU), Big Torch (S&JP). Vouchers: ABS, JT, GU.
Strumigenys silvestrii Emery. This neotropical species is probably exotic in Florida.
It is known from a few widely scattered localities in the state. Nests are in leaf litter.
Plantation (DS), Big Pine (MD), West Summerland (K&P). Vouchers: ABS, GU.
Tapinoma litorale Wheeler. This species occurs in the West Indies and the southern
third of Florida. We consider this a native species. Nests are in twigs and vines, usually
in exposed or partially exposed sites. Elliott (MD), Largo (EW), Plantation (EW),
Windley (EW), Big Pine (EW, GU, MD), Long (MD), Bush (NC). Vouchers: ABS, NC,
Tapinoma melanocephalum (Fabricius). This exotic species is common in disturbed
habitats through southern Florida. Nests are under bark, at the bases of palm leaves,
and other similar cavities. Elliott (MD), Largo (MD), Plantation (DS), Upper
Matecumbe (MD), Grassy (MD), Long (MD), Vaca (GU), Big Pine (MD, GU),
Saddlebunch 2 (JW), Stock Island (MD), Key West (JW, MD), Dry Tortugas (CW, NC).
Vouchers: ABS, NC, GU.

Florida Entomologist 71(2)

Tapinoma sp. This species appears related to T. sessile (Say), but differs in a number
of morphological characters and its preference for marshes as nesting sites. Colonies
have been found in salt marshes and in Everglades sawgrass prairies. The colonies are
usually in tussocks or at the base of shrubs. Specimens found tending a lycaenid larva
in a coastal marsh on Sugarloaf by D. Harvey. Vouchers: ABS, Los Angeles Co.
Museum. Record courtesy of J. Longino, University of California, Santa Barbara.
Tetramorium bicarinatum (Nylander). Widely known in the literature as T.
guineense (Fabricius), this exotic species is widely dispersed in Florida, though seldom
very abundant. It is usually found in rather moist disturbed areas. Largo (EW), Bahia
Honda (MD). Vouchers: ABS.
Tetramorium caldarium (Roger). Records of this species were generally confused
with those of T. simillimum until Bolton's revision of the genus in 1979. This exotic
species occurs throughout southern Florida, usually in open disturbed areas. Upper
Matecumbe (MD), Stock Island (MD), Key West (MD), Dry Tortugas (CW). Vouchers:
Tetramorium simillimum (F. Smith). This exotic species occurs throughout Florida,
usually in disturbed open areas such as lawns and foundation plantings. Elliott (MD),
Upper Matecumbe (MD), Bala Honda (MD), Big Pine (MD), Key West (EW). Vouchers.
Trachymyrmex septentrionalis (McCook). This native species, occurs throughout
Florida in dry, open, sandy areas. Several colonies were found on Long Key (MD).
Vouchers: ABS.
Trachymyrmex sp. nr. jamaicensis (Andlre). The name jamaicensis has been
applied to a large dark brown species found in extreme southern Florida and in the
Keys. There appear to be differences, particularly a conspicuous carina on the antennal
scape, between the Florida populations and those of the West Indies. This species
occurs in tropical hammocks, and its nests are marked by a conspicuous thatched tur-
rent. Elliott (MD, NC), Largo (MD), Grassy (MD, JT), Indian (GU), Long (MD), Bahia
Honda (MD), Big Pine (GU, S&JP, MD), Shark (NC). Vouchers: ABS, JT, NC, GU.
Wasmannia auropunctata (Roger). We have found this exotic ant a dominant
species at some collection sites in southern Florida, including a Coccoloba stand on
Bahia Honda. This species has eliminated all native ants from parts of the Galapagos
Islands (Clark et al. 1982, Lubin 1984), but has not had a similar effect during its long
history in southern Florida. It was not reported by Wilson in his 1964 paper, and has
evidently become much more abundant since then. Populations may still be on the
increase in the Keys. Largo (MD), Plantation (MD), Upper Matecumbe (MD), Bahia
Honda (MD, JT), Big Pine (NC, GU, MD), Sugarloaf (S&JP), Middle Torch (S&JP),
Key West (JW, NC, MD). Vouchers: ABS, NC, GU.
Xenomyrmex floridanus Emery. This native species nests in hollow twigs. Elliott
(MD), Largo (EW, MD), Plantation (EW, MD), Grassy (MD), Long, (GU), Ohio (GU),
Big Pine (GU, MD), No Name (MD), Ramrod (NC), Key West (EW, MD). Vouchers:
Zacryptocerus varians (F. Smith). This native species nests in hollow twigs,
branches hollowed by termites, and, occasionally, weed stems. Elliott (MD), Largo
(EW), Plantation (EW), Ohio (GU), Big Pine (EW, GU, MD), No Name (MD, S&JP),
Big Torch (S&JP), Upper Matecumbe (MD), Stock Island (K&P), Key West (EW).
Vouchers: ABS, GU.


Diversity. The known diversity of the ant fauna has increased from 30 species to 83
species. The fauna no longer seems particularly depauperate, especially by Antillean

June, 1988

Deyrup et al.: Florida Ants 173

standards. Even some larger islands of the West Indies have produced shorter lists of
ants. M. R. Smith, who studied the ants of Puerto Rico for about a year, found 66
species (Smith 1936). Wheeler, drawing on the studies of a number of workers, lists 72
species from Trinidad (Wheeler 1916). Cuba has a more impressive fauna of about 137
species (Alayo 1974). Longer lists could undoubtedly be compiled from large West In-
dian islands today, but their fauna may still be depauperate compared with the Florida
Keys when the respective sizes of these islands are taken into account. The fauna of
the Florida Keys is, in turn, rather small when compared to that of the mainland. From
one site in southern Florida 102 species of ants have been reported (Deyrup & Trager
1986), to which we recently added 2 additional species. In one sense, therefore, the ant
fauna of the Florida Keys seems rather rich, but in another sense it seems rather
The richness of the Keys fauna is derived in part from its close association with the
mainland. This has allowed migration of about 31 widely distributed species of the
southeastern coastal plain. This immigration must have been facilitated by broad land
bridges that extended through the Keys in the past (Hoffmeister & Multer 1968). It
seems likely, however, that the Keys fauna would be richer still if the Keys were
attached to a large land mass with a tropical rather than a temperate climate.
The depauperate nature of the fauna relative to the mainland is probably due to the
habitats and climate of the Keys. A number of species that are associated with well-
drained sandy areas of the mainland seem to be missing from the Keys. The tropical
hammocks have a deep humus layer that would seem ideal for soil-dwelling species such
as dacetines, but the extreme dryness of this humus during much of the year appears
inimical to ants. Rotten wood, which harbors many specialized ants in wetter climates,
contains only a few generalists in south Florida in general and the Keys in particular.
Flooding during tropical storms may also take its toll of the ants, as suggested by
Wilson (1964), but the effects of the prolonged dry season are probably more important.
Exotic Species. The proportion of species introduced by man into the ant fauna of the
Keys is higher than in any other known area of the U.S. There are 27 recognized
exotics, 33% of the fauna, without counting any of the West Indian species that could
easily be unrecognized exotics. The proportion of exotic ants seems to decrease from
south to north in Florida. At the Archbold Biological Station in Highlands Co. there
are 20 species, or 19.6% of the total (Deyrup & Trager 1986), and in Alachua Co. there
are 17 species, or 15% of the total (Johnson 1987). The increase in exotics to the south
is partly due to heavy trade between the ports of tropical Florida and the rich faunas
of mainland tropical areas. It may also be partly due to an ecological vacuum caused by
the unsuitability of the climate and habitats of the Keys for many mainland species to
the north.
There has clearly been an increase in the populations of exotic ants since the 1958
survey. It would have been difficult to overlook such species as Paratrechina
guatemalensis, Pheidole megacephala, P. moerens, Tapinoma melanocephalum, and
Wasmannia auropunctata if these species had been as abundant as they are presently.
There is no clear indication, however, that the exotic species are displacing native
species. There are a few native species that are so scarce that it is difficult to avoid the
impression that they are being affected by the influx of exotics, or by other forms of
habitat modification. Such species include Crematogaster minutissima, Discothyrea
testacea, Myrmecina americana, Pachycondyla stigma, Smithistruma dietrichi, Parat-
rechina concionna, and P. wojciki. The exotic species themselves may not have reached
an equilibrium: Tetramorium bicarinatum, which was abundant 20 years ago (D. S.
Simberloff, personal communicationn, is now extremely scarce.
Endemic Species. One would not expect that an archipelago of recent origin closely
associated with the mainland would be home to many autochthonous species or sub-

Florida Entomologist 71(2)

June, 1988

species. There are endemic subspecies of mammals and reptiles in the Keys (Auffenberg
1982), as well as the butterfly, Papilio aristodemus ponceanus Schaus. The mammals
and reptiles, however, are even more strongly affected by barriers than the ants, and
both reptiles and butterflies are likely to show rather conspicuous geographic distinc-
tions in coloration caused by minimal genetic differences, which are eagerly seized upon
by avid collectors. Only two species of ants seem possible endemics. One is the uniden-
tified Leptothorax (Dichothorax), which is almost certainly derived from a more north-
ern Dichothorax. This species has not been found anywhere on the mainland. The
second species is the ambiguous Trachymyrmex sp., which seems to show at least
subspecific differences from West Indian populations of T. jamaicensis. This form has
also been found on the mainland in Dade County.
Unexpected Absences in the Fauna. There are a few additional species of south Florida
ants that we expected to find in the Keys. These include the exotics Strumigenys rogeri
Emery and Trichoscapa membranifera (Emery), which are widespread, but rather
localized, south Florida exotics. A native species, Leptogenys elongata manni Wheeler
should also occur in the Keys. Enclaves of sand-inhabiting ants might well occur in
relatively well-drained sandy sites in the Keys. An apparently isolated population of
Trachymyrmex septentrionalis has already been found on Long Key; other species with
similar edaphic requirements are Pheidole metallescens Emery, Monomorium viridum
Brown, Solenopsis pergandei Forel, and Paratrechina arenivaga (Wheeler). We had a
general expectation of finding more previously unreported West Indian species, but
succeeded in adding only two species to the fauna, Monomorium ebeninum and Lep-
tothorax torrei.
General Comments on the Fauna. In a biogeographical sense, the Florida Peninsula
resembles a mountain, whose peak is formed by the Keys. The apex of the mountain is
375 miles south of its origin and, like the peak of a mountain, projects into a climatic
zone unsuitable for most of the inhabitants of its base. A peculiar biota results from the
mingling of the more climatically adaptable of the species that have easy access from
the base, with a selection of species preadapted to the climate and habitat, and somehow
transported to the isolated, climatically appropriate zone. The young age of this isolated
zone in the Keys precludes the presence of many endemic or relict species. The age,
size, and isolation of the zone determines the establishment of species from other areas
of similar climate and habitat. The Keys are relatively young, but they offer ample land
masses for ant populations, and their isolation from other tropical areas is reduced by
human commerce.
In his earlier survey Wilson (1964) was most likely to collect the more abundant
species, which led to a view of the ant fauna as a depauperate Antillean fauna with a
strong component of recent tropical exotics. Our work strongly supports this view of
the dominant ants, so our ecological conception of the ants of the Keys has changed
little. From a biogeographical standpoint, the Keys now appear somewhat different, as
they clearly have an extensive fauna derived from the north.
The distribution of ant species among the Keys shows few, if any, effects of "equilib-
rium island biogeography." Most species apparently occur wherever their preferred
habitat is available. Relationships between island size and species diversity have been
applied to scarab beetles of the Florida Keys (Peck & Howden 1985), but in the case of
ants the larger number of species on the larger islands is most easily attributed to a
greater diversity of habitats on the larger islands. What evidence we have of species
turnover resembles that of mainland south Florida, caused by habitat changes and
recent invasions of exotics. Elliott Key, which is isolated, largely undeveloped, and free
from spraying for mosquitos, appears to lack 2 dominating exotics, Wasmannia au-
ropunctata and Paratrechina guatemalensis. Trachymyrmex sp. nr. jamaicensis is the
only native ant that appears unusually abundant. Elliott Key, insulated as much by


Deyrup et al.: Florida Ants 175

National Parks Service management as by its distance from the mainland, will probably
continue to diverge faunistically from most of the Florida Keys. We note that the key
with the richest ant fauna is Big Pine, from which 61 species have been collected. This
key has had large portions of natural habitat protected and maintained as part of the
National Key Deer Wildlife Refuge. We believe that this has been a major factor in
conserving the diversity of ant species on this island. Island biogeography is unfortu-
nately supplied with ever more examples from ever smaller fragments of natural
It seems inevitable that the known ant fauna of the Keys will continue to change.
We expect that exotic species will continue their trend toward ecological domination,
except in protected areas such as Elliott Key, Lignum Vitae Key, and parts of Largo,
Long, Bahia Honda, Big Pine, and No Name Keys. We also assume that the list of
species known from the Keys will increase to at least 90 species, including some native
species. A number of species on the present list are known only from one or a few
pockets of suitable habitat, and it seems unlikely that we have sampled the full diversity
of such sites. For example, a number of the islands are composed of porous rock and
underlain by a whole series of miniature caves and grottos that could have a cavernicol-
ous fauna. It is virtually certain that further myrmecological surprises await in the
Florida Keys.


We would like to acknowledge the collectors, listed in the Methods section, who
assisted in this project, and Stefan Cover (Harvard University) who assisted in identifi-
cations. The Florida Keys collecting program of Jan Klimaszewski, Stewart Peck, and
Jarmila Kukalova-Peck was supported by grants from the Natural Sciences and En-
gineering Research Council of Canada.


ALAYO, P. 1974. Introduction al studio de los Himenopteros de Cuba. Superfamlia
Formicoidea. Acad. Ciencias Cuba Ser. Biol. 53.
AUFFENBERG, W. 1982. Florida environments and their herpetofaunas. Part III.
Herpetogeography. Florida Herpetol. 4: 1-36.
BARONI URBANI, C. 1978. Material per una revision dei Leptothorax neotropicali
appartenenti al sottogenere Macromischa Roger, n. comb. (Hymenoptera:For-
micidae). Entomol. Basiliensia 3: 395-618.
BOLTON, B. 1979. The ant tribe Tetramoriini (Hymenoptera:Formicidae). The genus
Tetramorium Mayr in the Malagasy region and in the New World. Bull. Brit.
Mus. (Nat. Hist.) Entomol. Ser. 38: 1-181.
BROWN, W. L. AND W. W. KEMPF. 1960. A world revision of the ant tribe Basicero-
tini (Hymenoptera:Formicidae). Studia Entomol. 3: 161-249.
BRUES, C. T. 1952. Some evolutionary features inherent in the insect faunas of the
tropics. J. Florida Acad. Sci. 15: 149-154.
CARROLL, J. T. 1975. Biology and ecology of ants of the genus Aphaenogaster in
Florida. Unpublished Ph.D. Dissertation, Univ. Florida, Gainesville.
LACIS. 1982. The tramp ant Wasman:a auropunctata: autoecology and effects
on ant diversity and distribution on Santa Cruz Island, Galapagos. Biotropica
14: 196-207.
COLE, B. J. 1983. Assembly of mangrove ant communities: patterns of geographical
distribution. J. Anim. Ecol. 52: 339-347.
CREIGHTON, W. S. 1950. The ants of North America. Bull. Harvard Mus. Comp.
Zool. 104: 1-583.

Florida Entomologist 71(2)

June, 1988

DEYRUP, M. AND J. TRAGER. 1986. Ants of the Archbold Biological Station, High-
lands Co., Florida (Hymenoptera: Formicidae). Florida Entomol. 69: 206-228.
HOFFMEISTER, J. E. AND H. G. MULTER. 1968. Geology and origin of the Florida
Keys. Geol. Soc. Amer. Bull. 79: 1487-1502.
JOHNSON, C. 1986. A north Florida ant fauna. Insecta Mundi 1: 243-246.
LUBIN, Y. D. 1984. Changes in the native fauna of the Galapagos Islands following
invasion by the little red fire ant, Wasmannia auropuntata. Biol. J. Linn. Soc.
21: 229-242.
NAVES, M. A. 1985. A monograph of the genus Pheidole in Florida (Hymenopt-
era:Formicidae). Insecta Mundi 1: 53-90.
PECK, S. B. AND H. F. HOWDEN. 1985. Biogeography of scavenging scarab beetles
in the Florida Keys: post Pleistocene land-bridge islands. Canadian J. Zool. 63:
SMITH, D. R. 1979. Superfamily Formicoidea. Pp. 1312-1467 in K. V. Krombein, P.
D. Hurd, Jr., D. R. Smith, B. D. Burks (eds.). Catalog of Hymenoptera in
America north of Mexico. Smithsonian Institution Press, Washington, D.C.
SMITH, M. R. 1936. The ants of Puerto Rico. J. Agric. Univ. Puero Rico 20: 819-875.
SNELLING, R. R. In press. Taxonomic notes on Nearctic species of Camponotus,
subgenus Myrmentoma (Hymenoptera: Formicidae) in J. C. Trager, ed., Ad-
vances in Myrmecology. Florida and Fauna Publications, E. J. Brill Inc., NY.
SNELLING, R. R. AND G. C. Wheeler. 1979. The taxonomy, distribution and ecology
of California desert ants. Report Bur. Land Manage., U.S. Dept. Interior, River-
side, CA. 335 pp.
WARD, P. S. 1985. The Nearctic species of the genus Pseudomyrmex (Hymenoptera:
Formicidae). Quaest. Entomol. 21: 209-246.
WHEELER, W. M. 1916. Ants collected in Trinidad by Professor Roland Thaxter, Mr.
F. W. Urich, and others. Bull. Harvard Mus. Comp. Zool. 60: 323-330.
WILSON, E. 0. 1964. The ants of the Florida Keys. Breviora 210: 1-14.


Department of Entomology
Texas A&M University
College Station, Texas 77843-2475


Texas Agricultural Experiment Station
2415 East Highway 83, Weslaco, Texas 78597


A bibliographic revision of the neotropical cornstalk borer (NCB), Diatraea lineolata
(Walker) is presented. The bibliographical entries include information on distribution,
taxonomy, host plants, biology, damage, and control of the NCB. However, only a few
references cited include substantial and complete information on the NCB. Most of the
citations include brief local observations and preliminary information on this species.


Rodriguez-del-Bosque et al.: Cornstalk Borer Bibliography 177

The NCB has been poorly studied in spite of its wide distribution and importance as a
pest of corn in the neotropical region.


Se present una revision bibliografica sobre el barrenador neotropical del maiz
(BNM), Diatraea lineolata (Walker). Las citas bibliogrAficas incluyen informaci6n sobre
su distribuci6n, taxonomia, plants hospederas, biologia, daflos y control. Sin embargo,
pocas referencias presentan informaci6n substantial y complete sobre el BNM; la
mayoria de los reports incluyen breves observaciones locales e informaci6n preliminary
sobre este insecto. El BNM no se ha estudiado adecuadamente a pesar de su amplia
distribuci6n e importancia como plaga del maiz en la region neotropical.

The most economically important species of the genus Diatraea Guilding are sac-
charalis (Fabricius), grandiosella Dyar, and lineolata (Walker) because of their wide
distribution and injury to graminaceous crops, principally sugar cane, Saccharum of-
ficinarum L., corn, Zea mays L., and sorghum, Sorghum bicolor (L.) Moench. Morrison
et al. (1977) and Chippendale et al. (1985) prepared a bibliography for D. grandiosella,
and Roe et al. (1981) for D. saccharalis, however, no attempt has been made to locate,
summarize, and interpret the literature on D. lineolata.
The neotropical cornstalk borer (NCB), D. lineolata, described from Venezuela by
Walker (1856), is after D. saccharalis, the most widely distributed species of Diatraea.
It occurs in the Bahamas, Cuba, Grenada, Tobago, Trinidad, Mexico, Central America
and most of equatorial South America north of the Amazon River (Box 1950c). Within
the continental United States, it only occurs in the Rio Grande Valley, Texas (Anony-
mous 1966a).
The host plants of D. lineolata are more limited than D. saccharalis, but similar to
D. grandiosella. The larva is a destructive borer of corn, and is classified as a "domestic"
species since no true wild-grass host is known (Myers 1935b, Box 1951b). The NCB also
has been reported from teosinte, Euchlaena mexicana Schrad., Guatemala grass, Trip-
sacum laxum, wheat, Triticum aestivum L., sorghum, very rarely from sugar cane
(Box 1950a, 1950c, 1951b), johnsongrass, Sorghum halepense (L.) (Van Leerdam 1981),
and rice, Oryza sativa L. (Angeles et al. 1960). Damage to corn includes defoliation,
interference to the vascular system by tunnelling in the stalk, lodging due to stalk
weakening, and damage to the ear (Overman 1970).
Despite its wide distribution, the biology and ecology of D. lineolata have been
studied only in Trinidad (Hynes 1942, Kevan 1943, 1944), and Nicaragua (van Huis
1981). Some preliminary studies were conducted in Venezuela (Box 1950a) and Guate-
mala (Painter 1955). Oviposition by NCB occurs primarily on the upper leaves of corn
(Kevan 1944) at late whorl and tasseling (van Huis 1981). The average egg mass size
observed in the field is two (Overman 1970, van Huis 1981) although this number is
increased greatly under laboratory conditions (Kevan 1944, van Huis 1981). In Trinidad,
the duration from egg to adult was seven to nine weeks with six to eight instars (Kevan
1943). A larval aestivation occurs during the dry season with pupation commencing at
the onset of rain. The diapausing larva is characterized by the change from a spotted
to an immaculate morph (Hynes 1942, Kevan 1943, 1944, van Huis 1981). Aestivation
provides the species with a mechanism for survival during dry, host plant-free periods
(Kevan 1944).
Studies on suppression methods for NCB are limited, although some natural enemies
(parasites) and host plant resistance studies have been reported (Peairs & Saunders
1980). Van Huis (1981) reported a complete integrated pest management study on corn,
including D. lineolata, in Nicaragua.

Florida Entomologist 71(2)

June, 1988

Taxonomic confusion of NCB in the early literature led to several erroneous reports
(Box 1935b, 1949). Published references to D. lineolata in sugar cane in British Guiana
and Trinidad actually refer to D. impersonatella (e.g. Urich 1910, 1915, Wolcott 1913,
Bodkin 1913, Cleare 1922, 1923, Box 1926), in Mexico to D. grandiosella (e.g. Van
Zwaluwenburg 1923, 1926a, 1926b, Morrill 1925, Anonymous 1927, Van Dine 1929,
Osborn & Phillips 1946), and in corn and sugar cane in the United States to D. grand-
iosella (e.g. Dyar 1911, Barnes & McDunnough 1917, Morrill 1919, Holloway & Loftin
1919a,b, Vorhies 1919, USDA 1922, Howard 1923, Anonymous 1925). In addition, the
NCB has been mistakenly reported attacking sugar cane in Dutch Guiana by Van Dine
(1929), in Cuba by Van Dine (1926, 1929), and in Venezuela by Box (1927a). Finally,
the NCB has been confused with D. grandiosella attacking corn in northeastern Mexico
and south Texas, where the latter species has been erroneously reported by Elias (1970)
and Cevallos-Davila (1970). Examination of specimens from several comprehensive sur-
veys failed to detect D. grandiosella in this area (Agnew et al. in press).
NCB was referred to as Zeadiatraea lineolata in the literature after Box (1955)
established this new genus based on morphological characters. However, Bleszynski
(1966) returned lineolata and other species to Diatraea, their original genus.
Bibliographical entries were obtained from the following sources: Biological
Abstracts (1927, Vol. 1 through 1988, Vol. 84, No. 1), Entomology Abstracts (1969, Vol.
1 through 1987, Vol. 18, No. 11), Review of Applied Entomology, Series A (1913, Vol.
1 through 1987, Vol. 75, No. 11), and literature citations in the reviewed articles them-
selves. We also used the National Agricultural Library and Biosciences Information
Services computerized literature search system in the library of the Texas A&M Univer-
Most references cited are on file with the authors. The references are listed alphabet-
ically by authorss. Depending on the type of information given, a number after each
reference indicates the following categories:
(1) Distribution and catalog/listing.
(2) Taxonomy and morphology.
(3) Biology and ecology.
(4) Economic damage and host plants.
(5) Biological control.
(6) Host plant resistance.
(7) Chemical control.
(8) Erroneous reports for other species.
Only a small portion of the references cited here include substantial information on
NCB; the majority either make brief mention of NCB or are misidentities with other
species. We included many Mexican and Central American local reports since they are
not commonly cited in the literature. Unfortunately, with a few exceptions, most of
them either include general information or represent only preliminary observations on
NCB. Because D. lineolata has been poorly studied in spite of its wide distribution and
importance as a pest of corn, more detailed studies are needed on its biology, ecology,
and control.


Abarca, M., A. Cortes I. and S. Flores C. 1958. The sugarcane borers in Mexico;
an attempt to control them through parasites. Proc. 10th Int. Cong. Entomol. 4:827-34.
Agnew, C. W., L. A. Rodriguez-del-Bosgue and J. W. Smith, Jr. Misidentifications
of Mexican stalkborers in the subfamily Crambine (Lepidoptera: Pyralidae). Folia En-
tomol. Mex. (in press) (2)


Rodriguez-del-Bosque et al.: Cornstalk Borer Bibliography 179

Aguilar-Anzueto, L. 1985. Determinaci6n especifica de insects fit6fagos del maiz
(Zea mays L.), en dos municipios del Estado de Chiapas. Resimenes del XX Congreso
Nacional de Entomologfa. Cd. Victoria, Tamaulipas, M6xico. pp. 102-3. (1,4)
Aldrich, J. M. 1925. Two species of the Tachinidae genus Lixophaga, with notes and
key (Diptera). Proc. Ent. Wash. 26(6):132-6. (1,5)
Andrews, K. L. 1984. El manejo integrado de plagas invertebradas en cultivos
agron6micos, horticolas y frutales en la Escuela Agricola Panamericana. Proyecto Man-
ejo Integrado de Plagas en Honduras. E.A.P./A.I.D. Ministerio R.R.N.N. El
Zamorano, Honduras. 35 pp. (1,4)
Angeles, N., W. Szumkowski, and P. D. Paredes. 1960. Diatraea saccharalis (F.),
plaga del arroz en Venezuela. Agron. Trop. 9:127-32. (4)
Anonymous. 1925. Proceedings of the 6th Conference Western Plant Quarantine
Board. Denver, Colorado. Calif. Dept. Agric. Spec. Pub. 54, 100 pp. [Rev. Appl. En-
tomol. 13:637]. (8)
Anonymous. 1927. Insect pests of sugarcane (including utilization of parasites). Proc.
2nd Conf. Internat. Soc. Sugar Cane Technol. pp. 57-62. [Rev. Appl. Entomol. 17:464].
Anonymous. 1957. Pests of corn (Mex. Agr. Program). The Rockefeller Found.
Program in the Agricultural Sciences. Annual Report 1956-1957. pp. 264-9. (4,5)
Anonymous. 1959. Biological control studies (Mex. Agr. Program). The Rockefeller
Found. Program in the Agricultural Sciences. Annual Report 1958-1959. pp. 151-3. (5)
Anonymous. 1960. Pests of corn (Mex. Agr. Program). The Rockefeller Found.
Program in the Agricultural Sciences. Annual Report 1959-1960. pp. 76-7. (7)
Anonymous. 1966a. Neotropical corn borer, Zeadiatraea lineolata. USDA Coop.
Econ. Ins. Rep. 16(33):801. (1,4)
Anonymous. 1966b. Neotropical corn borer, Zeadiatraea lineolata. USDA Coop.
Econ. Ins. Rep. 16(36):876. (1,4)
Anonymous. 1966c. Outbreaks and new records. FAO Plant Prot. Bull. 14(5):124-5.
Anonymous. 1967a. Neotropical corn borer, Zeadiatraea lineolata. USDA Coop.
Econ. Ins. Rep. 17(2):15. (1,4)
Anonymous. 1967b. Neotropical corn borer, Zeadiatraea lineolata. USDA Coop.
Econ. Ins. Rep. 17(9):139. (1,4)
Anonymous. 1967c. Neotropical corn borer, Zeadiatraea lineolata. USDA Coop.
Econ. Ins. Rep. 17(28):615. (1,4)
Anonymous. 1968. Neotropical corn borer, Diatraea lineolata. USDA. Coop. Econ.
Ins. Rep. 18(42):978. (1,4)
Anonymous. 1969a. Neotropical corn borer, Diatraea lineolata. USDA. Coop. Econ.
Ins. Rep. 19(2):19. (1,4)
Anonymous. 1969b. Neotropical corn borer, Diatraea lineolata. USDA. Coop. Econ.
Ins. Rep. 19(34):660. (1,4)
Arnaud, P. H. Jr. 1978. A host-parasite catalog of north American Tachinidae (Dipt-
era). USDA Misc. Pub. No. 1319. p. 414 (1,5)
Barnes, W. and J. McDunnough. 1917. Check list of Lepidoptera of Boreal America.
Decatur, Illinois. p. 141. (1)
Beg, M. N. and F. D. Bennett. 1971. Accidental introduction of Diatraea centrella
(Moschi) into Abaco, Bahamas, and attempts at its control. Proc. Int. Soc. Sugar Cane
Technol. 14:418-23. (4)
Beg, M. N. and F. D. Bennett. 1973. Insects associated with sugarcane on Abaco
Island, The Bahamas. Proc. of the 1973 Meeting of the West Indies Sugar Technol.,
Barbados. West Indies Sugar Assoc. (Inc.) pp. 228-45. [Rev. Appl. Entomol. 64:1999].

Florida Entomologist 71(2)

June, 1988

Bennett, F. D. 1965. Tests with parasites of Asian graminaceous moth-borers on
Diatraea and allied genera in Trinidad. Commonwealth Ins. Biol. Control. Tech. Bull.
5:101-16. (5)
Bennett, F. D. 1966. Preliminary studies with Jaynesleskiajaynesi Aldrich, a poten-
tially important Tachinid parasite of Diatraea spp. Proc. of the 1966 Meeting of British
West Indies Sugar Technol. pp. 309-10. [Rev. Appl. Entomol. 60:210]. (5)
Bennett, F. D. 1969. Tachinid flies as a biological control agent for sugarcane moth
borers. pp. 117-48. In Pests of Sugarcane, J. R. Williams, J. R. Metcalf, R. W. Mungom-
ery, and R. Mathes (eds.). Elsevier Publ. Co., New York. 568 pp. (5)
Bennett, F. D. 1971. Current status of biological control of the small moth borers
of sugarcane Diatraea spp. (Lepidoptera-Pyralidae). Entomophaga. 16(1):111-24. (4,5)
Bennett, F. D., M. J. W. Cock, and F. A. Diaz. 1983. Allorhogas sp. n. (Braconidae)
a potential biological control agent for graminaceous stem borers from Mexico. ISSCT
Entomol. Newsl. 14:9-12 [Rev. Appl. Entomol. 72:733] (5)
Berry, P. A. 1959. Entomologia econ6mica de El Salvador. Bol. Tec. 24. Serv. Coop.
Agr. Salvadorefio Americano. Ministerio de Agr. y Gan. Santa Tecla, El Salvador. p.
120. (4)
Bleszynski, S. 1966. Studies on the Crambinae (Lepidoptera). Part 43. Further
taxonomic notes on some tropical species. Acta Zool. Cracov. 11:451-98. (2)
Bleszynski, S. 1967. Studies on the Crambinae (Lepidoptera). Part 44. New neotrop-
ical genera and species. Preliminary cheklist of neotropical Crambinae. Acta Zool.
Cracov. 12(5):39-100. (1)
Bleszynski, S. 1969. The taxonomy of the Crambinae moth borers of sugar cane. pp.
11-59. In Pests of Sugarcane, J. R. Williams, J. R. Metcalf, R. W. Mungomery, and R.
Mathes (eds.). Elsevier Publ. Co., New York. 568 pp. (2)
Bleszynski, S. and R. J. Collins. 1962. A short catalogue of the world species of the
family Crambidae (Lepidoptera). Acta Zool. Cracov. 7(12):197-389. (1)
Bodkin, G. E. 1913. Insects injurious to sugar-cane in British Guiana, and their
natural enemies. J. Board of Agric. Br. Guiana. 7(1):29-32. [Rev. Appl. Entomol. 1:521-
2]. (8)
Box, H. E. 1926. Sugar-cane moth borers (Diatraea spp.) in British Guiana. Bull.
Ent. Res. 16:249-66. (8)
Box, H. E. 1927a. Eleventh report upon entomological work. Central Aguirre Sugar
Co., Central Aguirre, P.R. 24 pp. [Rev. Appl. Entomol. 15:412-4]. (8)
Box, H. E. 1927b. Los parisitos conocidos de las species americanas de Diatraea
(Lepidoptera, Pyralidae). Rev. Ind. Agric. Tucuman. 18:53-61. (1,5)
Box, H. E. 1931. The Crambine genera Diatraea and Xanthopherene (Lep.,
Pyralidae). Bull. Ent. Res. 22:1-50. (1,2)
Box, H. E. 1933. Further observations on sugar-cane moth borers (Diatraea spp.)
in St. Lucia. Introduction of the Cuban parasite, Lixophaga diatraeae Townsend. Re-
port upon visit to St. Lucia, August-September, 1933. With an appendix on the recom-
mended biological control of the white coffee-leaf miner (Leucoptera coffeella) in St.
Lucia. 10 pp. [Rev. Appl. Entomol. 22:104]. (8)
Box, H. E. 1935a. New records and three new species of American Diatraea (Lep.:
Pyral.). Bull. Ent. Res. 26:323-33. (1,4)
Box, H. E. 1935b. The species of Diatraea attacking sugar-cane in the New World.
Proc. Int. Soc. Sug. Cane Technol. 5:470-6. (1,4)
Box, H. E. 1935c. The food plants of American Diatraea species. Imp. Col. Trop.
Agric. Trinidad, Memoir.11 pp. [Rev. Appl.Entomol. 24:22-23]. (1,4)
Box, H. E. 1938. Observations on sugar-cane moth borers (Diatraea spp.) in St.
Lucia. III. The introduction and establishment of the Amazon fly (Metagonistylum
minense Townsend) and control of Diatraea saccharalis Fabricius by means of this

Rodriguez-del-Bosque et al.: Cornstalk Borer Bibliography 181

parasite. Report upon a visit to St. Lucia, March-Abril, 1938. 25 pp. [Rev. Appl. En-
tomol. 27:182-3]. (1)
Box, H. E. 1947. Informe preliminary sobre los taladradores de la cafa de azicar
(Diatraea spp.) en Venezuela. Bol. Tec. Dep. Ent. Minist. Agric. Venezuela. 117 pp.
[Rev. Appl. Entomol. 37:273-4]. (4)
Box, H. E. 1948a. Notes on the genus Diatraea Guilding (Lepid., Pyralidae). Intro-
duction and parts I, II and III. Bol. Ent. Venez. 7:26-59. (1)
Box, H. E. 1948b. Report upon specimens of Diatraea Guild. in the Paris museum
with the description of a new species from Brazil (Lep., Pyral.). Rev. de Entomol.
19(3):419-22. (1)
Box, H. E. 1949. Notes on the genus Diatraea Guilding (Lepid., Pyral.). Parts IV
and V. Rev. de Entomol. 20:541-55. (1,4)
Box, H. E. 1950a. Investigaciones sobre los taladradores de la cafia de azfcar (Diat-
raea spp.) en Venezuela. Informe de progress durante 1947-49. Bol. Tec. Div. Ent.
Minist. Agric. Venezuela 2: 1-60 [Rev. Appl. Entomol. 39:92-4]. (3,4,5)
Box, H. E. 1950b. Report upon specimens of Diatraea Guilding (Lepidoptera,
Pyralidae) in the Cornell University collection. J.N.Y. Entomol. Soc. 58:241-5. (1)
Box, H. E. 1950c. The geographical and ecological distribution of some neotropical
species of Diatraea Guild. (Lep.: Pyralidae) and certain of their parasites. Int. Cong.
Ent. 8:351-7. (1,4)
Box, H. E. 1951a. Informe preliminary sobre los barrenadores o "borers" de la cafia
de azfcar (Diatraea, Chilo) en M6xico, a base de un viaje de reconocimiento efectuado
durante marzo-abril, 1951, a las regions cafieras: I Sinaloa, II Nayarit y XIV Huas-
tecas, con observaciones complementarias. Un. Nal. Prod. Azuc. Mexico, D.F. 93 pp.
[Rev. Appl. Entomol. 39:431-2]. (1,4,5)
Box, H. E. 1951b. New species and records of Diatraea Guild. from northern Ven-
ezuela (Lepid., Pyral.). Bull. Ent. Res. 42:379-98. (1,4)
Box, H. E. 1952. Investigaciones sobre los taladradores de la cafia de azfcar (Diat-
raea spp.) en Venezuela. El proyecto del combat biol6gico. Informe del progress
durante 1949-1951. Min. Agric. y Cria, Inst. Nac. Agric., Maracay. Bol. Tec. 5:1-52.
[Rev. Appl. Entomol. 42:213-4]. (4,5)
Box, H. E. 1953a. Informe sobre las plagas insectiles que atacan a a cafia de azfcar
en Mexico, a base de un viaje de recorrido efectuado durante mayo-julio, 1952, a las
regions cameras: I Sinaloa, VI Balsas, VII Tehuacin, VIIIb Papaloapan, XII Veracruz-
Central, XIII Costa de Veracruz y XIV Huasteca. Bol. Azuc. Mex. 44 suppl. M6xico,
D.F. 26 pp. [Rev. Appl. Entomol. 43:220-2]. (1,4,5)
Box, H. E. 1953b. The control of sugarcane moth borers (Diatraea) in Venezuela; a
preliminary account. Trop. Agric. 30:97-113. (4,5)
Box, H. E. 1953c. The history and changing status of some neotropical insect pests.
Trans. Int. Cong. Ent. 9(2):254-9. (1,4)
Box, H. E. 1955. New Crambine genera allied to Diatraea Guilding (Lepidoptera:
Pyralidae). III. Proc. R. Ent. Soc. Lond. (B) 24:197-200. (2)
Box, H. E. 1956. New species and records of Diatraea Guilding and Zeadiatraea
Box from Mexico, Central and South America (Lepid., Pyral.). Bull. Ent. Res.
47:(4):755-76. (1)
Box, H. E. 1960. The species of Diatraea and allied genera attacking sugarcane.
Proc. Int. Soc. Sug. Cane Technol. 10:870-6. (1,4)
Box, H. E. and P. Guagliumi. 1953. The insects affecting sugarcane in Venezuela.
Proc. Int. Soc. Sug. Cane Technol. 8:553-9. (4)
Cardin, P. 1915. Cafia de azfcar (Saccharum officinarum). Inf. Depto. Patol. Veg.
& Entomol., Est. Exp. Agron. Santiago de las Vegas, Cuba. 3(1909-1915):112-7. (4)
Centro Internacional de Mejoramiento de Maiz y Trigo. 1969. Stem borer resistance.

Florida Entomologist 71(2)

June, 1988

Report 1968-1969, CIMMYT (International Maize and Wheat Improvement Center). El
BatAn, M6xico. pp. 43-45. (6,8)
Centro Internacional de Mejoramiento de Matz y Trigo. 1970. Stem borer resistance.
Report 1969-1970, CIMMYT (International Maize and Wheat Improvement Center). El
Batan, M6xico. p. 38. (6)
Cevallos-DAvila, A. 1970. Reacci6n varietal del maiz a la infestaci6n artificial de
Diatraea saccharalis (Fabricius) y manejo del insecto en el laboratorio. Tesis M.C.
ITESM. Monterrey N.L., M6xico. 60 pp. (8)
Chippendale, G. M. 1979. The southwestern corn borer, Diatraea grandiosella: case
history of an invading insect. Univ. Missouri. Agric. Exp. Station. Res. Bull. 1031. 52
pp. (1)
Chippendale, G. M. and K. L. Cassatt. 1985. Case history of the southwestern corn
borer, Diatraea grandiosella. II. Annotated bibliography, 1977 to 1985. Misc.Publ.En-
tomol. Soc. Am. No. 60. 30 pp.
Cleare, L. D. 1922. Notes on small moth-borers of sugarcane in British Guiana. J.
Bd. Agric. British Guiana. 15(4):163-184 [Rev. Appl. Entomol. 11:113]. (8)
Cleare, L. D. 1923. Notes on small moth-borers on sugarcane in British Guiana.
Bull. Ent. Res. 13:457-68. (8)
Cock, M. J. W. 1982. Telenomus sp. (ex ova Diatraea rufescens) from Bolivia, a
potential biological control agent for Diatraea centrella. Entomology Newsletter. 12:6-7
[Rev. Appl. Entomol. 71:111]. (5)
Cock, M. J. W. 1983. Host range testing of the tachinid Diatraea parasites Pal-
pozenilia diatreaea Tns. and Paratheresia claripalpis (Wulp) (from Brazil and
Ecuador). Entomology Newsletter. 15:8-15. [Rev. Appl. Entomol. 72] (5)
D'Aure, N. and J. L. Fontenia-Rizo. 1986. Zoogeografia del g6nero Diatraea
(Lepidoptera: Pyralidae) y su situaci6n en Cuba. Ciencias Biol6gicas. 16:101-10. [En-
tomol. Abst. 18:118] (1)
Davis, E. G., J. R. Horton, C. H. Gable, E. V. Walter, and R. A. Blanchard. 1933.
The southwestern corn borer. USDA Tech. Bull. 388. 61 pp. (1)
Diaz-Palma, R. 0. 1957. Susceptibilidad de algunos mestizos de maiz al ataque del
barrenador Diatraea saccharalis Fab. y D. lineolata Walker. Tesis Ing. Agr. ITESM.
Monterrey, N.L., M6xico. 22 pp. (6)
Dyar, H. G. 1911. The American species of Diatraea Guilding (Lepid., Pyralidae).
Ent. News. 22:199-207. (8)
Dyar, H. G. and C. Heinrich. 1927. The American moths of the genus Diatraea and
allies. U.S. Nat. Mus. Proc. 71(19), 48 pp. (1,2)
Elias, L. A. 1970. Maize resistance to stalk borers in Zeadiatraea Box, and Diatraea
Guilding (Lepidoptera: Pyralidae) at five localities in Mexico. Dissertation. Kansas State
University, Manhattan. 172 pp. [Diss. Abstr. Intl. 31(3):1331-2B]. (1,6,8)
Estrada, F. A. 1960. Lista preliminary de insects asociados al maiz en Nicaragua.
Turrialba 10:68-73. (1,4)
Flores C., S. 1955. Combate biol6gico del barrenador de la cafia de azfcar. Agric.
Tec. Mex. 1(1):16-37. (5)
Flores C., S. and M. Abarca R. 1961. Principales plagas de la cafia de azfcar en
Mexico. IMPA Boletin de Divulgaci6n 4:47-58. (1)
Frohlich, G. and W. Rodewald. 1970. Pests and diseases of tropical crops and their
control. Pergamon Press. Oxford. 371 pp. (4)
Fuchs, T. W. and J. A. Harding. 1979. Seasonal abundance of the sugarcane borer,
Diatraea saccharalis, on sugarcane and other hosts in the Lower Rio Grande Valley of
Texas. Southwes. Entomol. 4(2):125-31. (4)
Holloway, T. E. and U. C. Loftin. 1919a. Insects attacking sugar cane in the United
States. J. Econ. Entomol. 12:448-50. (8)

Rodriguez-del-Bosque et al.: Cornstalk Borer Bibliography 183

Holloway, T. E. and U. C. Loftin. 1919b. The sugar-cane moth borer. USDA Bull.
796. pp 10-11. (8)
Howard, L. O. 1923. Report (1922-23) of the entomologists. U.S. Dept. Agric. 37
pp. [Rev. Appl. Entomol. 12:165-7]. (8)
Hynes, H. B. N. 1942. Lepidopterous pests of maize in Trinidad. Trop. Agric. 19:194-
202. (3,4,5)
Jepson, W. F. 1954. A critical review of the world literature on the Lepidopterous
stalk borers of graminaceous crops. Commonwealth Inst. Entomol. London. 127 pp.
Kaye, W. J. and N. Lamont. 1927. A catalogue of the Trinidad Lepidoptera Heteroc-
era (moths). Mem. Dept. Agric. Trinidad & Tobago. 3:128-9. (8)
Kevan, D. K. M. 1943. The neotropical cornstalk borer, Diatraea lineolata Walker
and the sugarcane moth borer, Diatraea saccharalis (F.) as maize pests in Trinidad
(B.W.I), with notes from Grenada. Trop. Agric. (Trinidad) 20:167-74. (3,4,5)
Kevan, D. K. M. 1944. The bionomics of the neotropical cornstalk borer, Diatraea
lineolata Wlk.(Lep., Pyral.) in Trinidad, B.W.I. Bull. Ent. Res. 35:23-30. (3)
King, A. B. S. and J. L. Saunders. 1984. Las plagas invertebradas de cultivos
alimenticios anuales en Am4rica Central. Overseas Development Administration, Lon-
don. pp. 53-54. (1,3,4,5,7)
Kirkland, R. L. 1982. Biology of Iphiaulex kimballi (Hym.: Braconidae), a parasite
of Diatraea grandiosella (Lep.: Pyralidae). Entomophaga. 27(2):129-34. (5)
Lacayo, L. 1977. Especies parasiticas de Spodoptera frugiperda (Smith), Diatraea
lineolata (Wlk.) y Trichoplusia ni (Hbn.) en zonas de Managua y Masatepe. XXIII
Reuni6n Anual del PCCMCA, Panama, PanamA. pp. 1-28. (5)
Loera-Gallardo, J. 1986. El gusano barrenador Diatraea lineolata (Walker) y su
efecto en el rendimiento del maiz. Resimuenes del XXI Congreso Nacional de En-
tomologia. Monterrey, N.L., M6xico. pp. 177-8. (4)
Mahadeo, C. R. 1980. Minor pests of sugar cane in Trinidad. J. Agric. Soc. Trinidad
Tobago. 80(2):120-8. (3,4)
Manrique-G6mez, F., A. I. Galindo-R., and N. G. Gonzalez-Hernandez. 1979. Fluc-
tuaci6n de las poblaciones de algunos insects de importancia econ6mica en la Comarca
Lagunera de los Estados de Coahuila y Durango. VII Reuni6n Nacional de Control
Biol6gico. Veracruz, Mexico. pp. 10-38. [Rev. Appl. Entomol. 68:212]. (3)
Mihm, J. A. 1984. T6cnicas eficientes para la crianza masiva e infestaci6n de insects,
en la selecci6n de las plants hospedantes para resistencia a los taladradores del tallo
del maiz Diatraea sp. Centro Internacional de Mejoramiento de Maiz y Trigo (CIM-
MYT), El BatAn, M6xico. 23 pp. (3,6)
Morrill, A. W. 1919. Report of the entomologists. Ann. Rept. Arizona Comiss. Agric.
& Hortic. 1917-1918, 10:29-73. [Rev. Appl. Entomol. 9:405-7]. (8)
Morrill, A. W. 1925. Commercial entomology on the west coast of Mexico. J. Econ.
Entomol. 18:707-16. (8)
Morrison, W. P., D. E. Mock, J. D. Stone, and J. Whitwort. 1977. A bibliography
of the southwestern corn borer, Diatraea grandiosella Dyar (Lepidoptera: Pyralidae).
Bull. Entomol. Soc. Am. 23(3):185-90.
Myers, J. G. 1931. Descriptions and records of parasitic Hymenoptera from British
Guiana and the West Indies. Bull. Ent. Res. 22:267-77. (5)
Myers, J. G. 1932a. Biological observation on some neotropical parasitic Hymenopt-
era. R. Entomol. Soc. Lond. Trans. 80:121-36. (5)
Myers, J. G. 1932b. The original habitat and hosts of three major sugar cane pests
of tropical America (Diatraea, Castnia, and Tomaspis). Bull. Ent. Res. 23(2):257-71. (4)
Myers, J. G. 1935a. Second report on an investigation into the biological control of
West Indian insect pests. Bull. Ent. Res. 26(2):181-252. (4)

Florida Entomologist 71(2)

June, 1988

Myers, J. G. 1935b. The ecological distribution of some South American grass and
sugar cane borers (Diatraeo spp. Lep., Pyralidae). Bull. Ent. Res. 26:335:42. (4)
Ortega, A. 1974. Maize diseases and insects. Proc. World Wide Maize Improvement
in the 70's and the role for CIMMYT. Centro Internacional de Mejoramiento de Maiz y
Trigo (CIMMYT). El Batan, Mexico. 41 pp. (1,4)
Ortega, A. and C. de Leon. 1971. Plant protection. Proc. of the 1st Maize Workshop.
CIMMYT (International Maize and Wheat Improvement Center). El Batan, Mex. pp.
95-102. (1,4)
Ortega, A., S. K. Vasal, J. A. Mihm, and C. Hershey. 1980. Breeding for insect
resistance in maize. pp. 371-419. In Maxwell, F. G. and P. R. Jennings (eds.). Breeding
plants resistance to insects. John Wiley and Sons. N.Y. 683 pp. (1,4,6)
Osborn, H. T. and G. R. Phillips. 1946. Chilo loftini in California, Arizona, and
Mexico. J. Econ. Entomol. 39:755-9. (8)
Overman, J. L. 1970. Relationship of resistance in maize (Zea mays L.) to two
related species of Pyralidae: Diatraea saccharalis (F.) and Zeadiatraea lineolata
(Wlk.). Dissertation. The University of Florida. 105 pp. [Diss. Abst. Intl. 31:6667B]. (6)
Painter, R. H. 1955. Insects on corn and teosinte in Guatemala. J. Econ. Entomol.
48:36-42. (3,4,5)
Peairs, F. B. and J. L. Saunders. 1980. Diatraea lineolata y D. saccharalis: una
revision en relaci6n con el maiz. Agron. Costarric. 4(1):123-35. (3,4,5,6)
Pschorn-Walker, H. and F. D. Bennett. 1970. Host suitability experiments with
three tachinids parasites of Diatraea spp. in Barbados and Trinidad, W.I. Proc. Int.
Soc. Sug. Cane Technol. 13:1331-41. (5)
Quezada, J. R. 1979. Poblaciones remanentes de barrenadores en cafias de maiz.
Bol. 1. Fac. de Ciencias y Humanidades, Univ. de el Salvador. San Salvador, El Sal-
vador. 22 pp. (3,5)
Reed-Gil, C. G. Estudio sobre la biologia de campo, hospederas y poblaciones del
barrenador Diatraea saccharalis Fabricius y D. lineolata Walker, en maiz. Tesis Ing.
Agr. ITESM. Monterrey, N.L. M6xico. 41 pp. (3,4,5)
Riess H., C. M. and S. Flores C. 1976. Catalogo de plagas y enfermedades de la
calfa de azicar en Mexico. Serie Divulgaci6n T6cnica IMPA II. CNIA. 177 pp. (1,4)
Risco, S. H. 1960. La situaci6n actual de los barrenadores de la cafia de azacar del
g6nero Diatraea y otros taladradores en el Peri, Panama y Ecuador. Rev. Peru. En-
tomol. Agric. 3(1):6-10. (4,5)
Rodriguez-del-Bosque, L. A., J. W. Smith,, Jr., and H. W. Browning. 1988. Damage
by stalkborers (Lepidoptera: Pyralidae) to corn in northeastern Mexico. J. Econ. En-
tomol. (in press). (3,4)
Roe, R. M., A. M. Hammond Jr., T. E. Reagan, and S. D. Hensley. 1981. A bibliog-
raphy of the sugarcane borer, Diatraea saccharalis (Fabricius), 1887-1980. USDA,
ARS, Agric. Rev. and Manuals S-20. 101 pp.
Rolston, L. H. 1955. The southwestern corn borer in Arkansas. Ark. Agric. Exp.
Stn. Bull. 553. 40 pp. (1)
Saenz, L. and F. Sequeira. 1972. Especies parasiticas del gusano cogollero, Spodopt-
era frugiperda (J. E. Smith) y de barrenador del tallo del maiz, Diatraea lineolata
(Wlk.), encontradas en los diferentes Campos Experimentales del PMMYSN. XVIII
Reuni6n Anual del PCCMCA, Managua, Nicaragua. 6 pp. (5)
Scaramuzza, L. C. 1933. Prospects for the control of the sugar cane moth stalkborer
(Diatraea saccharalis Fab.) in Cuba by means of natural enemies. Proc. 6th Conf. Asoc.
Tec. Azuc. Cuba. pp. 87-93. [Rev. Appl. Entomol. 22:185-6]. (5)
Scaramuzza, L. C. 1939. The introduction of Theresia claripalpis V.D.W., into
Cuba, and its artificial multiplication. Proc. Int. Soc. Sug. Cane Technol. 6:589-95. (5)
Scaramuzza, L. C. 1945. Biological control of the sugarcane borer in Cuba by means
of the fly Lixophaga. Mem. Asoc. Tec. Azuc. Cuba. 19:11-6. (5)
Scaramuzza, L. C. 1956. Achievements in the biological control of the sugarcane
borers Diatraea spp. (Lepidoptera: Pyralidae) in the Americas. Proc. Int. Cong. En-
tomol. 4:845-50. (5)
Sequeira, R. A. 1987. Studies on pests and their natural enemies in maize and
sorghum in Honduras. M.S. Thesis. Texas A&M University, College Station, Tx. 296
pp. (3,4,5)


Rodriguez-del-Bosque et al.: Cornstalk Borer Bibliography 185

Sequeira, R. A., F. E. Gilstrap, K. L. Andrews, D. H. Meckenstock, and H.
Fuentes. 1986. DinAmica de poblaciones de Diatraea lineolata (Walker) en sistemas de
cultivo de pequefios agricultores del sur en Honduras. XXXII Reuni6n Anual del
PCCMCA, San Salvador, El Salvador. (3)
Sequeira, R. A., F. E. Gilstrap, K. L. Andrews, D. H. Meckenstock, and H.
Fuentes. 1986. Importancia de la hormiga brava, Solenopsis geminata en maiz y sorgo
sembrados en cultural mixta en Choluteca, Honduras. XXXII Reuni6n Anual del
PCCMCA, San Salvador, El Salvador. (5)
Serrano-Cervantes, L., G. Henriquez-Martinez, J. A. Najera-Montes, Reyes, and
R. A. Sequeira. 1986. Determinaci6n de la occurrencia de barrenadores, Diatraea
(Pyralidae, Lepidoptera) y del nivel de control biol6gico native en El Salvador. XXXII
Reuni6n Anual del PCCMCA, San Salvador, El Salvador. (3,5)
Simmonds, F. J. 1958. The successful breeding of Palpozenillia palpalis (Ald.)
(Diptera, Tachinidae) a parasite of Diatraea spp. Trop. Agric. 35(3):218-224. (5)
Simmonds, F. J. 1963. Genetics and biological control. Canad. Entomol. 95(6):561-7.
Todd, C. J. and F. L. Thomas. 1930. Notes on the southwestern corn borer, Diatraea
grandiosella Dyar. J. Econ. Entomol. 23:118-21. (1)
Urich, F. W. 1910. Sugar cane insects in Trinidad. West India Bull. 12: 388-91. (8)
Urich, F. W. 1915. Insects affecting the sugar cane in Trinidad. Bull. Dept. Agr.
Trinidad & Tobago. 14:156-61. (8)
U.S. Department of Agriculture. 1922. The Insect Pest Survey Bulletin 2(1):1-32.
[Rev. Appl. Entomol. 10:331-2]. (8)
Valle-Duarte, G. A. 1958. Biologia de los barrenadores del maiz, Diatraea sac-
charalis (F) y D. lineolata (Walker) y experiments de control. Tesis Ing. Agr. ITESM.
Monterrey, N.L., M6xico. 105 pp. (3,4,7)
Van Dine, D. L. 1926. The sugar cane moth stalkborer. Trop. Plant Res. Found.
Bull. 2, 11 pp. [Rev. Appl. Entomol. 14:542]. (8)
Van Dine, D. L. 1929. Parasites of sugar cane moth borers. J. Econ. Entomol.
22:248-68. (8)
Van Dine, D. L. and L. D. Christenson. 1932. A revised list of the insects affecting
sugar cane in Cuba. Int. Soc. Sug. Cane Technol. 4th Congress. Bull. 116. 3pp. (4)
Van Emden, F. I. 1949. The scientific name of the common tachinid parasite of
Diatraea spp. (Lep., Pyral.) in Central and South America, with notes on related species
(Dipt.). Rev. Entomol. 20:499-508. (5)
van Huis, A. 1975. Diatraea lineolata (Wlk.): Diapause and spatial distribution in
sorghum stubbles in Nicaragua. Project FAO/UNDP/Nic/70/002, Progress Report No.
1. Managua. 12 pp. (3)
van Huis, A. 1977. Duty travel report: Guatemala, El Salvador, Honduras, Costa
Rica, Panama. Project FAO/UNDP/Nic/70/002. Report to FAO. Managua. 21 pp. (3)
van Huis, A. 1981. Integrated pest management in the small farmer's maize crop in
Nicaragua. Meded Landbouwhogesch Wageningen. 81(6):1-221. (3,4,5,7)
Van Leerdam, M. B. 1981. Parasitism of Diatraea saccharalis (F.) infesting Johnson
grass, by the braconid parasite Apantelesflavipes (Cameron). M.S. Thesis. Texas A&M
University, College Station, Tx. 67 pp. (4,5)
Van Zwaluwenburg, R. H. 1923. Tachinids and Sarcophagids established in Mexico.
J. Econ. Entomol. 16:227. (8)
Van Zwaluwenburg, R. H. 1926a. Insects enemies of sugarcane in western Mexico.
J. Econ. Entomol. 19:664:69. (8)
Van Zwaluwenburg, R. H. 1926b. Some sugar cane insects of the Pacific Coast of
Mexico. Bernice P. Bishop Mus. Spec. Pub. 11:51-52. [Biol. Abstr. 2: 1104]. (8)
Velazquez-Hernandez, A. M. 1957. Zonas preferentes de ataque de Diatraea sac-
charalis F. y D. lineolata W. y experiments de control quimico. Tesis Ing. Agr.
ITESM. Monterrey, N.L., M6xico. 59 pp. (4,7)
Vesey-Fitzgerald, D. 1935. Progress report for February and March, 1935. I. Divi-
sion of Entomology, Sugar-cane Investigation Commitee, Trinidad. (4)
Vignes, W. G. des. 1983a. Rearing, release and recovery of Allorhogas sp. n.
(Hymenoptera: Braconidae)-a potential biological control agent of Diatraea spp. in

186 Florida Entomologist 71(2) June, 1988

Trinidad. Ent. Newsl., Int. Soc. Sug. Cane Technol. 14:34. [Rev. Appl. Entomol.
72:214]. (5)
Vignes, W. G. des. 1983b. Laboratory hosts for rearing Allorhogas sp. n., a potential
biocontrol agent of Diatraea spp. on sugarcane in Trinidad. Ent. Newsl. 15:13. [Rev.
Appl. Entomol. 72:327]. (5)
Vorhies, C. T. 1919. Entomology. Ann. Rept. Arizona Agric. Expt. Sta. 30: 437-8.
[Rev. Appl. Entomol. 9:119]. (8)
Walker, F. 1856. List of the specimens of Lepidopterous insects in the collection of
the British Museum. Part IX. London, 252 pp. (2)
Wolcott, G. N. 1913. Report on a trip to Demerara, Trinidad and Barbados during
the winter of 1913. J. Econ. Entomol. 6:443-57. (8)
Youm, 0. 1984. Stemborers attacking Sorghum bicolor (L.) Moench and Zea mays
L. in the Lower Rio Grande Valley. M.S. Thesis. Texas A&M University, College
Station, Tx. 84 pp. (3,4,5)


Approved by the Texas Agricultural Experiment Station as TA 22815.


Department of Zoology,
National University of Singapore,
Kent Ridge, Singapore 0511

Florida Medical Entomology Laboratory,
University of Florida,
200 9th Street S.E.,
Vero Beach, FL 32962


A new species of Atrichopogon (Diptera: Ceratopogonidae), whose immature stages
are found on leaves of the water lettuce, Pistia stratiotes L., is described in all stages.
The adults have a glossy black head and thorax, white abdomen, and dark terminal
tarsal segments. The pupa possesses large, elongate tubercles and bears dark brown
mediodorsal and dorsolateral pigmented spots on the first six abdominal segments. The
larva is atypical, resembling those of the subgenus Forcipomyia. The larva does not
resemble any of the Atrichopogon species figured by Ewen and Saunders (1958). The
species is quite distinct and different from the seven described Florida species of At-
richopogon listed by Wilkening et al. (1985).

Se describe en todas sus etapas una nueva especie de Atrichopogon (Diptera:
Ceratopogonidae), cuya etapa inmadura se encuentra en hojas de la lechuga Pistia

Chan & Linley: New Florida Ceratopogonid


stratiotes L. Los adults tienen la cabeza y el t6rax negro lustroso, el abdomen blanco,
y obscuros los segments terminales del tarso. La pupa posee tub6rculos largos, elon-
gados, y tiene manchas pigmentadas pardas obscuras en el medio dorso y en el dorso
lateral de los seis primeros segments abdominales. La larva es atipica, parecida a
aquellas del subg6nero Forcipomyia. La larva no se parece a ninguna de las species
de Atrichopogon figuradas por Ewen y Saunders (1985). La especie es bien distinta y
diferente de las siete species descritas de la Florida por Wilkening et al. (1985).

Atrichopogon Kieffer is one of two genera in the ceratopogonid subfamily For-
cipomyiinae Lenz, the other genus being Forcipomyia Meigen. The type species, At-
richopogon levis, was originally described from a single male specimen by Coquillett as
Ceratopogon exilis. Known as the "grass punkie", it is widely distributed in the United
States (Wilkening et al. 1985) and its immature stages are commonly found in grassy
areas such as pastures, parks and lawns (Boesel & Snyder 1944). The immature stages
figured as A. levis by Ewen & Saunders (1958) are, in fact, those ofA. geminus Boesel.
The adults of Atrichopogon are notoriously difficult to separate to species. Even the
male genitalia do not offer useful distinct characters, or combinations of characters,
owing to extreme similarity between species. The immature stages, however, particu-
larly the larvae, are excellent for species recognition (Nielsen 1951, Ewen & Saunders
1958). The larval head and bodily chaetotaxy are specifically unique. We have therefore
followed Nielsen (1951) and Ewen & Saunders (1958) in defining this new species on the
basis of the larva and, accordingly, have designated a larva as the holotype.
The species is named in honor of Dr. Willis W. Wirth, whose eminent contributions
to ceratopogonid taxonomy are well known. Dr. Wirth cooperated with us in determin-
ing the status of this new species.


Eggs, larvae and pupae were obtained from the mature leaves of water lettuce
plants collected from Chinese Farm, a location adjacent to Old Dixie Highway, about 3
miles south of the Florida Medical Entomology Laboratory, Vero Beach, Indian River
County, Florida. Adults were reared from immatures. Larvae from hatched eggs were
reared in petridishes on pieces of fresh Pistia leaf cut from areas where most of the
older larvae are found. The leaf substrate, with its natural microfauna and flora served
as the rearing medium. Larvae hatched from eggs were also reared by supplementing
the natural food with small quantities of lactalbumin/brewer's yeast (1:1) mixture,
sprinkled sparingly on the leaves. A thick layer of wet paper towel or filter paper was
placed in each petridish to provide a moist micro-environment. After pupation, pupae
with associated larval skins were placed individually on small strips of moist paper
applied to the inside walls of small specimen tubes, until adult emergence.
For killing, storing and examining all stages, the techniques essentially followed
those of Chan & LeRoux (1965). Larvae were killed in a small quantity of water in a
4.8 x 1.5 cm tube held over a flame to induce extension of the pseudopods and anal
papillae. Some larvae killed directly in 80% ethanol also extended their pseudopods and
papillae. Living pupae were killed by immersion in 80% ethanol, but had first to be
taken out of their larval exuviae by carefully cutting away the larval skins from the
terminal appendages (sexual processes). This tedious procedure was accomplished using
minute nadeln for cutting the larval skin along the sides of the terminal processes,
while the pupa was pushed away from its exuvium with a very fine brush. The numerous
minute, anteriorly-directed spines of the two terminal processes (Fig. 16) prevented the
larval skin from being dislodged easily. Living pupae often writhed about when dis-
turbed unduly, or when accidentally pricked by the needle. Such movements assisted

Florida Entomologist 71(2)

June, 1988

disengagement from the larval skin, but removal could, nonetheless, require as long as
half an hour. Adults were killed after they had assumed full coloration and their integu-
ment had hardened (12-24 hr after emergence), by touching them with an ethanol-wet-
ted brush and then immersing them in a tube of 80% ethanol. Pupae and genitalia of
adult males were cleared in hot KOH for 30 sec., or by leaving them in the KOH solution
overnight. Cleared specimens were washed in several changes of distilled water, then
stained in alcoholic fast green solution (2 gm fast green FCF in 2 ml glacial acetic acid
and 100 ml absolute ethanol). After staining, specimens were transferred to lactophenol
(phenol, lactic acid, glycerine, distilled water 3:1:2:1) and examined under a compound
The terms used in the diagnosis follow those of Chan & LeRoux (1965), except for
the male genitalia, which follow Wirth et al. (1977), where "claspers" are synonymous
with "parameres" of Snodgrass (1957) and "parameres" are synonymous with "claspet-
tes" of Snodgrass (1957).
Examination of all stages and measurements of key characters were done with a
Zeiss Universal research microscope at various magnifications. Measurements are pre-
sented in the text as the mean, followed by the range in parentheses.
The terminology used in the description of the larval and pupal appendages follows
that of Ewen & Saunders (1958). Adults are described using the terminology of Wirth

K /-



Metathorax J

Figs. 1-2. Atrichopogon wirthi sp. n. 1, egg. 2, 4th instar larva, lateral view.




Thorax Head



I I~

Chan & Linley: New Florida Ceratopogonid

(1952) and Wirth et al. (1977). Adult length was measured from the vertex of the head
to the wing tip (Tables 1, 2), rather than to the abdominal tip. Measurement to the wing
tip is more accurate because the abdomen in both sexes, prior to post-emergence
diuresis (Linley 1984), is longer than in older individuals.

Atrichopogon wirthi Chan and Linley, new species

Egg (Fig. 1)
Material: 14 eggs
Shape: banana-shaped (Fig. 1), anterior end blunt and rounded, posterior end more
Length: 429.5 pm (418.5-441.8 iJm).
Width: 92.2 jim (90.7-93.0 jm).
Color: grey, pinkish-red eyes of larva visible at anterior end of mature eggs.



anterior row
of 6 hooklets

Figs. 3-6. Atrichopogon wirthi sp. n., 4th instar larva. 3, head, lateral view. 4, head,
dorsal view. 5, prothorax, lateral view. 6, prothoracic pseudopod, ventral view.

Florida Entomologist 71(2)

June, 1988

7 9

.. h; A ,


8 b

x Vill

Figs. 7-9. Atrichopogon wirthi sp. n. 4th instar larva. 7, abdominal segment 1,
lateral view, showing chaetotaxy. 8, abdominal segments VIII and IX, lateral view,
showing chaetotaxy, anal papillae and posterior pseudopod. 9, posterior pseudopod,
ventral view.


Chan & Linley: New Florida Ceratopogonid

Larva, 4th instar (Fig. 1-9)
Material: 37 specimens
Total length: 2.20-3.79 mm.
Color: pale white (living specimens).
Head: (Fig. 3, 4), length 307-335 mr, depth 221-246 im, width about 246 Jim, eye a
prominent dome, moderately large, raised above surface, bearing two pinkish anterior
lobes, set high. Antenna long, socle about as long as width at base. Head setae as in
Fig. 3 and 4.
Thorax: (Fig. 2, 5, 6), chaetotaxy similar in prothorax and mesothorax, different in
metathorax. Prothorax with 3 short, stout setae (a, b, c) on dorsum (Fig. 5); setae d,
e, f simple, setae g minute, paired. Mesothorax with same setae as prothorax, but setae
a, b, c simple, not stout. Metathorax without setae c and f, otherwise similar to
mesothorax. Prothoracic pseudopod (Fig. 5, 6) bilobed, each bearing 2 rows of hooklets,
anterior (6 hooklets) hyaline, slender, longer; posterior (4 hooklets) dark colored, stout
and shorter. Common base of the pseudopod with about 15 transverse rows of extremely
fine and minute shagreened hooklets ventrally (Fig. 6). Posterior to 20 hooklets on both
lobes, are 2 patches of fine, minute and shagreened hooklets.
Abdomen: (Fig. 2, 7-9), segments not flattened (segment side not extended laterally to
form processes, as in typical Atrichopogon larva). Chaetotaxy of segments I-VII similar,
as figured (Fig. 2,7). Chaetotaxy of segments VIII and IX different (Fig. 8), and both
differ from segments I-VII. Segment VIII lacks setae c, d and f, but setae a and b are
stout as in segments I-VII. Segment IX with chaetotaxy as figured (Fig. 8), posterior
end bearing 4 anal papillae, each bilobed apically, anal pseudopod bearing 10 shorter,
stouter and dark hooklets in anterior row, and 10 longer, more slender and hyaline
hooklets in posterior row (Fig. 9).

Pupae: (Fig. 10-16)
Material: 4 specimens (3 d 1 9).
Color: white in living state, tinged with green and with dark brown pigmented spots
on mediodorsum and dorsolateral positions of abdominal segments I-VI, as figured (Fig.
Total length: 6 2.36 mm (2.26-2.52 mm); 9 2.02 mm.
Cephalothorax: cephalothoracic dorsal length, 6 0.67 mm (0.65-0.70 mm), 9 0.55 mm.
Cephalothoracic ventral length, S 1.13 mm (1.12-1.14 mm), 9 1.01 mm. Cephalothoracic
width, & 0.54 mm (0.52-0.56 mm), 9 0.50 mm.
Head: (Fig. 10-12), eye large, raised above surface, set high. Frons (Fig. 10) bears 3
small frontal processes, 2 anterior ones with setae, short, and one posterior one without
setae, low (Fig. 10, 11). Epicranium bears a pair of small setae on a low process, the
epicranial process (Fig. 10). Ventral posterior region of head as figured (Fig. 12).
Thorax: prothoracic respiratory horn (Fig. 10) small, narrow, with angular base, boot-
shaped bearing bout 8 spiracular papillae in continuous row over top. Mesothoracic
dorsum furnished with 6 pairs of cuticular processes, 4 (ml, m2, m3, m4) are elongate,
styliform and state and covered basally with short spinules (Fig. 10, 11). Mesothoracic
processes m5 and m6 reduced to inconspicuous nodules. Metathorax bisected by poste-
rior median point of mesothorax and bears a pair of small, lateral knobs.
Abdomen: (Fig. 10-16), length 6 1.69 mm (1.60-1.85 mm), 9 1.40 mm. Abdominal
height, S 0.38 mm (0.37-0.39 mm), 9 0.31 mm. Abdominal segment I bears, in common
with segments II-VI a pair of large, elongate, state mediodorsal processes and a pair
of small lateral processes, each bearing 2 setae (Fig. 10). Abdominal segments III-VI
with similar chaetotaxy and dark brown pigmentation. Each segment bears a pair of
elongate, state mediodorsal processes and a similar pair of lateral processes, as well
as a pair of small lateroventral setae and 2 pairs of small, closely approximating ventral
setae (Fig. 13). Segment II lacks the 2 pairs of closely approximating ventral setae, but
is otherwise similar in chaetotaxy to segments III-VI. Caudal segments as shown (Fig.
15, 16).

Florida Entomologist 71(2)

respiratory horn
\ eF^.

icranial process

m /mediodorsal
-*-./ process

posterior median
point of mesotho

Fig. 10. Atrichopogon wirthi sp. n., pupa, dorsal view, showing chaetotaxy.

Adult Male (Fig. 17-24)
Material: 10 specimens
Color: in both living and preserved specimens, head and thorax glossy black, terminal
tarsal segments of legs also dark. Abdomen white. Abdominal segments I-VI with dark
spots on mediodorsum and dorsolateral edges. These spots correspond to the dark
brown pigment spots in same segments of pupa.
Total length: 1.90 mm (1.85-1.99 mm) from head to wing tip.
Head: (Fig. 17, 18), eyes black, frons pale white, proboscis dark brown. Antenna (Fig.
17) black, segments 12-15 elongate, segment 12 only slightly elongate, segment 11 not

June, 1988


Chan & Linley: New Florida Ceratopogonid



process .


Figs. 11-12. Atrichopogon wirthi sp. n., pupa, 11, lateral view, showing chaetotaxy.
12, posterior region of head, ventral view.

elongate, AR 1.73 (1.64-1.86). Maxillary palp (Fig. 18) with 3rd segment slender, bear-
ing a shallow cylindrical sensory pit, PR 3.30 (3.00-3.75).
Thorax: (Fig. 19-22), wing length 1.16 mm (1.10-1.21 mm), width 0.32 mm (0.29-0.34
mm), CR 0.66 (0.64-0.67). Legs with terminal tarsal segments dark, fore TR 3.17 (3.05-
3.37), mid TR 3.42 (3.14-3.55), hind TR 2.44 (2.28-2.57). Foreleg with apical tibial comb
(a grooming apparatus for the head and anterior thoracic region) bearing a closely touch-
ing row of about 14 small, short spines, a few adjacent longer and thicker setae and 1
large, thick spur (Fig. 19). Mid leg lacks grooming organ, but bears about 5 large setae
near apex (Fig. 20). Hind leg with apical tibial comb (an organ for grooming wings and
abdomen) bearing 2 rows of spines, anterior row composed of about 6 longer and thicker
spines and posterior row consisting of about 20 closely touching, shorter and thinner


Florida Entomologist 71(2)

June, 1988

" -. ., V.

.:- ".-.j.
..- -. t- .-

Lpigmented brown spot

lateral process

--lateroventral seta

ventral setae




Figs. 13-16. Atrichopogon wirthi sp. n., pupa. 13, 4th abdominal segment (male).
14, last 3 abdominal segments, normally inside last larval exuvium. 15, abdominal seg-
ment IX (female). 16, abdominal segment IX (male), showing long sexual processes
bearing forwardly directed spinules.




Chan & Linley: New Florida Ceratopogonid


17 18


0 2 21


Figs. 17-22. Atrichopogon wirthi sp. n., adult male. 17, antenna. 18, maxillary palp.
19, tibio-tarsal grooming apparatus of foreleg. 20, tibio-tarsal joint of midleg. 21, tibio-
tarsal grooming organ of hind leg. 22, scutellum.

spines (Fig. 21). Claws on all legs bifid at the tip (Fig. 28). Scutellum glossy black as
thoracic dorsum, with 4 long, marginal bristles and 3 shorter ones (Fig. 22).
Abdomen: (Fig. 23, 24), genitalia (Fig. 23, 24) as follows. Ninth sternite bearing about
8-10 setae, 9th tergite rounded, 1.18 (1.06-1.30) times as long as wide. Claspers with
basistyles slender, extending to posterior tip of 9th tergite, dististyles long and slender,
tapering to pointed and incurved, heavily sclerotized apices, basistyles 106.58 pLm

Florida Entomologist 71(2)

9th Sternite



-9th Sternite



Figs. 23-24. Atrichopogon wirthi sp. n., Male genitalia. 23, ventral view showing
details of aedeagus (coarsely stippled), claspers, and parmeres (finely stippled). 24,
lateroventral view, showing juxtaposition of aedeagus (coarsely stippled) and parameres
(finely stippled).

(99.96-117.60 im) long, CLR clasperr ratio = length of dististyle/length of basistyle)
0.82 (0.75-0.86). Aedeagus typical of genus, trilobed (Fig. 24), simple, shield-shaped
(Fig. 23), with basal arms stout, basal arch deep, caudolateral shoulders broad. Mesal
point of aedeagus funnel-shaped basally and broadly pointed at tip, which bears 2 tiny,
recurved hooklets (Fig. 23, 24). Parameres (basistylar apodemes) a broad, somewhat
kidney-shaped structure (Fig. 23, 24). Posterior to tip of aedeagus on ventral side of
9th tergite is a heart-shaped patch of minute setae (Fig. 23). Summary of measurements
of key characters in 10 specimens shown in Table 1.

Adult Female (Fig. 25-29)
Material: 10 specimens.
Color: as in male.
Total length: 1.77 mm (1.68-2.00 mm) from front of head to wing tip.
Head: (Fig. 25, 26), eyes black, frons pale white, proboscis dark brown. Antenna (Fig.


June, 1988

Chan & Linley: New Florida Ceratopogonid 197

w 00 00 ^

- O1-
c4o E

z 4



o C) U t:
1- rl0_

m cq
F- ^g *s^ s^

S ~ r3 *C <

a .- ts o ^ M



o 'wI1 t

z a-
0 ^

o o~
g ~H M

F. | ^ I e
Cc'3 a0
co aoir


0 '
0 ry* ^" ^

Florida Entomologist 71(2)

June, 1988

25 27



S \ gland
I/ spermathecal
fat layer gland duct



sternite _

pedicel cercus

Figs. 25-29. Atrichopogon wirthi sp. n., adult female. 25, antenna, showing scape,
pedicel and flagellum bearing 13 flagellomeres. 26, maxillary palp. 27, wing, showing
venation. 28, last tarsal segment of foreleg, showing bifid claws. 29, spermatheca and
spermathecal gland attached to last abdominal segment.

25) black, flagellomeres 9-13 elongate, flagellomeres 1-8 short and globular, AR 2.14
(1.92-2.27). Maxillary palp (Fig. 26) with 3rd segment scarcely swollen, bearing a
sunken, cylindrical sensory pit, PR 2.60 (2.33-2.80).
Thorax: (Fig. 27, 28), wing (Fig. 27) length 1.02 mm (0.96-1.11 mm), width 0.37 mm
(0.35-0.41 mm), CR 0.69 (0.67-0.70). Legs with terminal tarsal segments dark, fore TR
3.18 (2.94-3.38), mid TR 3.30 (3.11-3.46), hind TR 2.33 (2.22-2.46). Apical tibial combs

Chan & Linley: New Florida Ceratopogonid






w C.J
C d


c- as





Q 0


ia eo


Florida Entomologist 71(2)

June, 1988

in fore and hind legs as in male. Claws on all legs bifid at tip (Fig. 28). Scutellum as in
the male (Fig. 22).
Abdomen: (Fig. 29), one spermatheca, globular, about 1.5x as long as wide, length 86.44
Vm (70.56-99.96 pm) width 61.15 Rm (48.52-67.62 jLm). Spermathecal duct covered with
layer of fat cells along entire length, but these cells not enveloping spermatheca. Sper-
mathecal gland finger-like, long, narrow, with duct arising a short distance from gono-
pore and joining spermathecal duct (Fig. 29). Summary of measurements of key charac-
ters in 10 specimens shown in Table 2.
Holotype: whole 4th instar larva slide preparation, collected from leaves of water let-
tuce, Pistia stratiotes, between July and September 1987, at Chinese Farm, Ft. Pierce,
St. Lucie County, Florida.
Paratypes: 3 slide preparations of 4th instar larvae, and about 80 larvae, pupae and
adults (both sexes) with associated pupal exuviae, in ethanol. All type specimens depo-
sited in U.S. National Museum collection, Washington, D.C. Also, in Dr. Wirth's collec-
tion are one 4th instar larva, 1 male and 2 females, with associated pupal skins.


The adults of Atrichopogon are difficult to separate to species. Even the male
genitalia of different species are very similar and many species share the common,
typical, shield- or cooking-pot-shaped aedeagus, as seen in corpulentus, caribbeanus,
remigatus, maculosus, saundersi, bifidus, meloesugans (of the subgenus Melohelea),
inconspicuous, geminus (as levis), fuscus, polydactylus and crinitus, all figured in Ewen
& Saunders (1958). Atrichopogon wirthi is no exception in this respect. Furthermore,
in all these species, including A. wirthi, the apex of the aedeagus is ventral to a heart-
shaped patch of fine setae (Fig. 23). The parameres, too, are very similar in all these
species and, likewise, the dististyles are not distinctly longer than the basistyles, except
for A. fuscus and A. crinitus. Thus, it is inadvisable to describe species from the adults
We agree with Nielsen (1951) and Ewen & Saunders (1958) that Atrichopogon
species are more easily recognized on the basis of the larvae and/or pupae. We have,
accordingly, designated a 4th instar larva as the holotype. Based on larval morphology,
A. wirthi may be easily mistaken for a species of Forcipomyia. Unlike typical and
unmistakable Atrichopogon larvae, which are flattened in appearance owing to lateral
extension of the thoracic and abdominal segments, the A. wirthi larva has a long,
cylindrical body and a head very much resembling the subgenus Forcipomyia.
Of the 19 species of Atrichopogon described as larvae by Ewen & Saunders (1958),
only 1, A. crinitus (from rotting wood in water at Nanaimo, Canada), has larvae re-
sembling Forcipomyia. Atrichopogon wirthi, however, differs from A. crinitus in hav-
ing much shorter a, b and c setae in the larva. Atrichopogon crinitus has seta a and
seta c on prothorax to abdominal segment VII 4x as long as the segment, and seta b
on prothorax to abdominal segment VIII about 1.5x as long as the segment. Atrichopo-
gon wirthi larvae also differ from those of A. crinitus in the number of hooklets on the
prothoracic pseudopod. On each lobe of the deeply cleft pseudopod A. crinitus has 2
anterior hooklets and 3 posterior hooklets, as opposed to 6 and 4, respectively, in A.
Wilkening et al. (1985) listed 7 described Florida species of Atrichopogon. They state
that there are at least 30 undescribed Nearctic Atrichopogon species in the collections
of the National Museum of Natural History in Washington, D.C. They further state
that "this genus is in need of comprehensive study and revision". It is hoped that this
paper will stimulate interest in this project.


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