morales-ramos et al.: two populations of c. grandis 137...

Morales-Ramos et al.: Two populations of

C. grandis 137

DIFFERENCES IN REPRODUCTIVE POTENTIAL OFTWO POPULATIONS OF

CATOLACCUS GRANDIS

(HYMENOPTERA: PTEROMALIDAE) AND THEIR HYBRIDS

J

UAN

A. M

ORALES

-R

AMOS

, M. G

UADALUPE

R

OJAS

AND

E

DGAR

G. K

ING

USDA-ARSSouthern Regional Research Center

New Orleans, Louisiana

A

BSTRACT

A new colony of the boll weevil ectoparasitoid

Catolaccus grandis

was introducedfrom Guasave, Sinaloa, Mexico to improve vigor of a 12-year-old laboratory rearedstock in Weslaco, Texas. The biological characteristics of the introduced colony werecompared to those of the Weslaco colony and a crossbreed of these 2 colonies. Devel-opmental time was not significantly different among the 3 colonies, but the preovipo-sitional period of the Sinaloa females was 3 times as long compared to the other 2colonies. The fecundity, net reproductive rate (

R

o

), and intrinsic rate of increase (

r

m

) offemales from Sinaloa were significantly lower than those of females from Weslaco andthe hybrid colony. Generation time (

G

) and doubling time (

DT

) were significantlylonger in the Sinaloa colony. These characteristics make the Sinaloa population lessdesirable for mass propagation and release to control boll weevil populations than theWeslaco colony. The biological and population parameters of the hybrid colony werenot significantly different from those of the Weslaco population. The implications ofthe observed results on the mass propagation and release strategies against the bollweevil are discussed and recommendations are presented.

Key Words: Boll weevil, parasitoid, population, variability, biological parameters, bi-ological control

R

ESUMEN

Una nueva colonia del parasitoide del picudo del algodonero

Catolaccus grandis

deGuasave, Sinaloa, México, fue introducida a Texas con el objetivo de mejorar el vigorde la colonia madre de este parasitoide. Las características biológicas de la colonia in-troducida fueron comparadas con aquellas de la colonia de Weslaco, Texas y el híbridoentre estas dos colonias. No hubo diferencias significativas en el tiempo de desarrolloentre las tres colonias, pero el período de preoviposición de las hembras de Sinaloa fueel triple del observado en las otras 2 colonias. La fecundidad, tasa reproductiva neta(

R

o

) y tasa intrinseca de crecimiento (

r

m

) de las hembras de Sinaloa, fueron significan-temente bajas comparadas con las de las hembras hibrídas y de Weslaco. Los tiemposde generación (G) y de duplicación (DT) fueron significantemente mas largos en la co-lonia de Sinaloa. Estas características indican que el uso de la colonia de Sinaloa enun programa de propagación en masa y liberación contra el picudo del algodonero po-dria ser desventajoso. Sin embargo, las características biológicas de la colonia híbridano fueron significativamente diferentes a las observadas en la colonia de Weslaco. Sediscuten las implicaciones de estos resultados en las estrategias de propagación en

masa y liberación contra el picudo del algodonero y se presentan recomendaciones.

Augmentative releases of the exotic ectoparasitoid

Catolaccus

grandis

(Burks)(Hymenoptera: Pteromalidae) have proven effective at biologically controlling the boll

138

Florida Entomologist

83(2) June, 2000

weevil,

Anthonomus

grandis

grandis

Boheman (Coleman et al. 1996, King et al. 1995,Morales-Ramos & King 1991, Morales-Ramos et al. 1994, 1995, Summy et al. 1992,1993, 1994, 1995, Vargas-Camplis et al. 1997). Rearing methods for this parasitoidhave been improved over the years (Cate 1987, Morales-Ramos et al. 1992, 1994,1997, Palamara 1995, Roberson & Harsh 1993). The development of an artificial diet(Rojas et al. 1996, 1997) significantly advanced the mass propagation technology forthis parasitoid. The effectiveness of diet-reared

C.

grandis

has been established bylaboratory and field evaluations (Morales-Ramos et al. 1995, 1998, R. J. Coleman, un-published). All these advances have made the mass propagation of

C. grandis

econom-ically feasible according to an economic analysis done by Ellis et al. (1997).

The process of colonization and mass rearing over long periods of time may havedetrimental effects in reared species (Bartlett 1984). Initial loss of genetic variabilityand subsequent selection tend to induce domestication and adaptation to laboratoryenvironments (Bartlett 1984, van Lenteren 1991, Wajnberg 1991, Lappla 1993). After12 years of constant laboratory rearing of

C. grandis

, there is no evidence of loss ofsearching capacity or field adaptation in this parasitoid (Morales-Ramos et al. 1995,Vargas-Camplis et al. 1997); however, the potential for loss of genetic variability andsubsequent adaptation to laboratory environments is a constant concern. Some meth-ods for avoiding such problems include 1) pooling multiple-founder colonies, 2) main-taining variable laboratory environments, and 3) regular infusion of wild geneticstock (Joslyn 1984).

The first method has been used with the colony of

C. grandis

that is currentlymaintained in Weslaco, TX. This colony is the result of systematic cross breedingamong 3 distinct populations of

C. grandis

from Tabasco and Oaxaca Mexico, and fromEl Salvador (J. A. M., unpublished data) (see Materials and Methods). The secondmethod is difficult to apply because it requires large supplies of equipment, space, andpersonnel. These resources have not been available to the present research program.This study is an attempt to apply the third method by introducing wild

C. grandis

from Sinaloa, Mexico to prevent the loss of genetic variability from the Weslaco colony.The objectives were to evaluate the reproductive potential of the newly introducedparasitoids to determine the desirability of cross breeding the wild population fromSinaloa with the Weslaco population.

M

ATERIALS

AND

M

ETHODS

Boll weevil larvae used in this study were reared on artificial diet at the USDA-ARS, Biological Control and Mass Rearing Research Unit at Mississippi State, Mis-sissippi (Roberson & Wright 1984). Rearing techniques for

C. grandis

were as re-ported by Morales-Ramos et al. (1992) using the Parafilm® encapsulation methoddeveloped by Cate (1987). Parasitoid colonies and experiments were held at constant27

±

1

°

C, 50

±

10% R.H., and a photoperiod of 14:10 (L:D).

Biological Materials Origin

A

C. grandis

colony was established in Guasave, Sinaloa, Mexico in early June,1996 using parasitoids isolated from boll weevil infested cotton squares and bolls andfrom net captures of adult

C. grandis

females. The infested cotton material and adultparasitoids were collected from a commercial cotton field located near Guasave. Thecolony was maintained and increased using Cate’s (1987) method of encapsulation.

Two shipments of

C. grandis

were received at the APHIS quarantine facility locatedat the Biological Control Center in Mission, TX (importation permit No. 31352). The ship-


C. grandis 139

ments consisted of 120 females and 136 males on August 1st and 144 females and 106males on August 8, 1996. The

C. grandis

colony was held in quarantine for one generationwhile being screened for purity and microbial contamination. The Sinaloa colony of

C.grandis

(Sinaloa population) was released from quarantine on October 25, 1996 (permitNo. 31669) to the USDA-ARS Subtropical Agricultural Research Center in Weslaco, TX.The parasitoids used for this study were from the 2

nd

generation after introduction.The mother colony maintained at Weslaco, TX (Weslaco population) originated

from 2 different localities in Southeast Mexico and one locality from El Salvador. Thelocalities in Mexico were Cardenas, Tabasco from

Hampea nutricia

(29 males and 13females); La Ventosa, Oaxaca from

Cienfuegosia rosei

(1 male and 7 females); and inEl Salvador, The University of El Salvador, from cultivated and wild cotton (2 malesand 2 females). Samples of plant buds and fruits infested by boll weevil were hand-carried to the quarantine facility at Texas A&M University in College Station, TX.These colonies were reared independently for 3 years at the Department of Entomol-ogy, Texas A&M University, from 1985 to 1988.

A crossbred colony was produced in 1989 from a systematic cross between the 3 pop-ulations. This was accomplished by placing 300 newly emerged females from one colonyand 300 newly-emerged males from another in a new cage. A total of 6 combinationswere initially created. The progeny of all the 6 combinations were then mixed andreared as a single colony. This colony was successfully transferred to Weslaco, TX in Feb-ruary 1990 and it has been in constant culture since (approximately 153 generations).Several permits have been issued to release this particular population in the state ofTexas. The most recent one was issued on September 21, 1994 (permit No. 944483).

The third

C. grandis

colony tested was a crossbreed (hybrid) of the Sinaloa (12

th

generation) and Weslaco (generation 169) populations. The hybridization was accom-plished by placing 100 female pupae of each population in separated cages made of2.8-liter Rubbermaid® containers (No. 6 Rubbermaid, Wooster, OH). The pupae wereallowed to emerge at 27

±

1

°

C, 60

±

10% RH, and a photoperiod of 14:10 (L:D). One dafter emergence, the 100 females from the Weslaco population received 50 newly-emerged males from the Sinaloa population. Similarly, the 100 females from the Si-naloa population were provided with 50 newly emerged males from the Weslaco pop-ulation. Each cage was provided with distilled water, honey and 120 encapsulated bollweevil larvae, which were replaced daily for a period of 20 d. The parasitized boll wee-vils were maintained at the conditions described above for 10 d to allow the parasi-toids to develop to mid-age pupae. Eighteen female parasitoid pupae were randomlyselected from weevils parasitized by each of the 2 groups when the females were 8 dold. This procedure was repeated again using weevils parasitized when the femaleswere 12 d old to obtain a total of 72 hybrid

C. grandis

pupae.

Pupal Weight

A total of 72 female parasitoid pupae of each population (3 d-old) were weighed in-dividually using a Mettler H51 precision balance. The weights of the different groupswere analyzed by Analysis of variance (ANOVA) and the means were compared byTukey’s test using SigmaStat

(

software (Jandel Corporation 1995). The female para-sitoid pupae were placed individually in plastic square Petri dishes (9

´

9 cm) wherethey completed development at the conditions described above.

Fecundity and Progeny Sex Ratio

Once the female parasitoids completed development and emerged, two males fromthe same population were placed in each of the Petri dishes to ensure fertilization.

140


83(2) June, 2000

Each female was provided daily with 12 encapsulated boll weevils, water and honeyaccording to evaluation methods reported by Morales-Ramos and Cate (1992). Deadfemales were not replaced, but dead males were replaced during the first 15 d. Eachday, the Parafilm® capsules enclosing the parasitized weevils were opened to countthe number of eggs oviposited per female. They were resealed and returned to the en-vironmental chamber for parasitoid development. Nine d later, the Parafilm® cap-sules were reopened to count and sex the parasitoid pupae. The number of eggsoviposited per female per day and the number and sex of developing progeny were re-corded over a 45-d period.

The sample size needed to estimate the population mean (

m

) of eggs/female andeggs/female/d for confidence intervals (E) of

±

20 and 1.5, respectively, with

a

= 0.05,was determined by using the equation:

where

n

is the sample size,

Z

a

/2

= 1.96 (from tables), and

s

is the population standarddeviation (estimated from sample ‘

s

’). This equation determines the adequate samplesize for a given value of

E

based on the standard deviation of a preliminary samplesfrom each population (Ott 1984).

The total number of eggs oviposited by each female during the 45-d period and themean number of eggs oviposited per day during the fecundity plateau period (as de-fined by Morales-Ramos and Cate 1992) were used to compare the fecundity of fe-males from each of the 3 populations studied. The starting age of the fecundityplateau period was determined according to the criteria used by Morales-Ramos andCate (1992). The sex of each of the female’s progeny was recorded and the sex ratio ofthe progeny was calculated. The Analysis of variance was used to compare fecundityand progeny sex ratio between the three populations studied and Tukey’s test wasused to compare the means using SigmaStat® software.

R

ESULTS

AND

D

ISCUSSION

Biological Parameters

Developmental time of

C. grandis

females was not significantly different amongthe Sinaloa and Weslaco populations and their hybrid (Table 1). The preovipositionalperiod, on the other hand, was significantly longer (16 days) in the Sinaloa populationthan in the Weslaco and hybrid populations (4.3 and 3.6 d respectively) (

F

= 64.4,

df

= 2, 126,

P

< 0.001) (Table 1). The difference observed between the Weslaco and hybridpopulations was not significant, showing that the hybrids tend to resemble more theWeslaco population for this parameter instead of showing an intermediate value be-tween the Weslaco and Sinaloa populations as expected.

A longer preovipositional period is considered to be an undesirable trait. In manyparasitoid species, the females do not respond to host cues during this period (Vinson1981, 1984). Evidence from field studies on searching capacity indicated that this maybe the case in

C. grandis

; no parasitism of boll weevil larvae was observed in an ex-perimental field in Ricardo, TX up to 5 d after the release of newly emerged females(J. A. M. unpublished data). However, a release of 5-d old parasitoids produced highrates of parasitism during the same period of time in the same experimental field(J. A. M., unpublished data).

No significant differences in female longevity and pupal weight were observedamong the Sinaloa and Weslaco populations and their hybrids (Table 1). Parasitoid

n Za 2¤( )2 s2( ) E2¤=


C. grandis 141

females from the Weslaco and hybrid populations oviposited a significantly highernumber of eggs (554 and 600 respectively) than that of females from the Sinaloa pop-ulation (242) (

F

= 37.6,

df

= 2, 213,

P

= 0.001) (Table 1). Total fecundity was not sig-nificantly different between the Weslaco and hybrid populations, again, showingresemblance of the hybrids to the Weslaco population.

Parasitoids from the Weslaco and hybrid populations produced most of their prog-eny earlier in their life cycle than those of the Sinaloa population (Fig. 1A). The fecun-dity plateau period (period of highest fecundity) started earlier in the Weslaco andhybrid populations (9 and 8 d of age, respectively) than in the Sinaloa population (23 dof age). The daily oviposition of Weslaco and hybrid parasitoids during the fecundityplateau period was significantly higher (22.0 and 21.6 eggs/d, respectively) than thatof parasitoids from the Sinaloa population (14.5 eggs/d) (

F

= 95.3,

df

= 2, 2103,

P

<0.001) (Table 1). The daily oviposition rate during this period was not significantly dif-ferent between the Weslaco and hybrid populations.

A reduced oviposition rate is another undesirable trait. The effectiveness of

C. gran-dis

in controlling the boll weevil depends on the rate at which the females find and par-asitize host larvae. Females from the Sinaloa population seem to parasitize at nearlyhalf the rate as females from the Weslaco population, and at more than twice the age.

There was no significant difference in progeny sex ratio among the 3 populations(Table 1). This indicates that the rates of fertilization and host acceptance were not af-fected by hybridization.

These results indicate that the undesirable characteristics of the Sinaloa popula-tion were not manifested after hybridization with the Weslaco population. However,we recommend caution in introducing the Sinaloa strain to the mother colony(Weslaco). The detrimental characteristics observed in the Sinaloa population may re-surface in subsequent generations in the hybrid population. Since the only apparentpotential advantages of introducing the Sinaloa strain to the mother colony is gain ofgenetic variability, we recommend the use of a different strain to achieve this purpose.

T

ABLE

1. B

IOLOGICAL

PARAMETERS

OF

3

C.

GRANDIS

POPULATIONS

.

Parameter

Population

Weslaco Hybrid Sinaloa

Developmental time (females)

1

13.4

±

0.6 13.5

±

0.7 13.8

±

1.3

Preovipositional period

1

4.3

±

3.4b 3.6

±

4.3b 16.0

±

10.1aLongevity

1

43.2

±

20.7a 44.1

±

16.4a 35.9

±

22.6a

Fecundity

Total eggs 554.0

±

247a 600.1

±

253a 242.0

±

300bEggs/F./d

2

22.0

±

8.8a 21.6

±

9.8a 14.5

±

13.5bPupal weight

3

6.4

±

1.1a 6.3

±

1.0a 6.6

±

1.5aProgeny sex ratio

4

5.1

±

4.4a 4.5

±

3.6a 4.6

±

4.3a

X

±

SD, means with the same letter are not significantly different after ANOVA Tukey test

a

= 0.05.

1

In days.

2

During the fecundity plateau period.

3

In mg.

4

In females per male.

142


83(2) June, 2000

Fig. 1. Age-dependent fecundity and survival of females of 2 C. grandis popula-tions and their hybrid. A) Age-dependent fecundity expressed as eggs oviposited perfemale per day; B) age-dependent survival expressed as lx which is the proportion ofindividuals surviving from eclosion to the beginning of age x (in days); and C) Age-de-pendent reproductive value (Vx), which is the potential female progeny of a female ofage x (in days).


C. grandis 143

A

CKNOWLEDGMENTS

We thank Ma. Teresa Chávez technical director of Asesoria Biológica Integral fromGuasave, Sinaloa, México for her assistance in collecting, rearing, and shipping of theSinaloa specimens of

C. grandis

. We also thank Nina M. Barcenas from the Colegio dePostgraduados, Montecillo, México for her help in providing training and materials tothe contacts in Sinaloa. This was accomplished with financial support of USDA-RSED-FAS Grant No. FG-Mx-101 Project No. MX-ARS-2.

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♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

and Plants, Wageningen, Netherlands.

POTENTIAL OF FIELD CORN AS A BARRIER CROP AND EGGPLANT AS A TRAP CROP FOR MANAGEMENT OF BEMISIA ARGENTIFOLII (HOMOPTERA: ALEYRODIDAE) ON COMMON

BEAN IN NORTH FLORIDA

HUGH ADAM SMITH AND ROBERT MCSORLEY

Department of Entomology and NematologyUniversity of Florida

Gainesville, FL, 32611

ABSTRACT

Trap crops and barrier crops are among the cultural control methods promoted formanagement of Bemisia argentifolii Bellows & Perring, particularly for small farmersin the tropics. In 1996 eggplant, Solanum melongena L., was tested as a trap crop, andin 1996 and 1997 corn, Zea mays L., was tested as a barrier crop for management ofB. argentifolii on bean, Phaseolus vulgaris L. In 1996 treatments were compared bysampling immature B. argentifolii on bean leaves. Neither egg nor nymphal densitieswere reduced by eggplant or corn treatments in 1996. In the 1997 corn barrier trialplot size was increased and the orientation of barrier row to wind direction was eval-uated. A dust-and-release procedure was used to measure entry of greenhouse-rearedadult B. argentifolii into experimental plots. Counts from yellow sticky traps in 1997indicated that migration by adult whiteflies into plots was determined primarily byair currents and was only marginally influenced by the presence of a corn barrier. Theresults indicate that barrier crops and certain trap crops may have limited value forwhitefly management.

Key Words: Intercropping, polyculture, vector management, wind dispersal, pestmanagement

Smith & McSorley: Trap and Barrier Crops for Whitefly

145

V

AN

L

ENTEREN

, J. C. 1991. Quality control of natural enemies: hope or illusion?, pp.1-14

in

F. Bigler (ed.), Proceedings of the Fifth Workshop of the IOBC GlobalWorking Group “Quality Control of Mass Reared Arthropods”. InternationalOrganization for Biological Control of Noxious Animals and Plants, Wagenin-gen, Netherlands.

V

ARGAS

-C

AMPLIS

, J., R. J. C

OLEMAN

, J. G

ONZALEZ

,

AND

L. R

ODRIGUEZ

D

EL

B. 1997.Life table analysis of cotton boll weevil in the tropics of Tamaulipas, Mexico af-ter

Catolaccus grandis

releases, pp. 1194-1197

in

Proc. Beltwide Cotton Con-ferences 1997 Vol. 2. National Cotton Council of America, Memphis, TN.

V

INSON

, S. B. 1981. Habitat location, pp. 51-77

in

D. A. Norlund, R. L. Jones, and W.J. Lewis (eds.), Semiochemicals their role in pest control. John Wiley & Sons,New York, NY.

V

INSON

, S. B. 1984. Parasitoid-host relationship, pp. 205-233

in

W. J. Bell and R. T.Cardé (eds.), Chemical Ecology of Insects. Sinauer Associates Inc. Sunderland,MA.

W

AJNBERG

, E. 1991. Quality control of mass-reared arthropods: a genetical and sta-tistical approach, pp. 15-25

in

F. Bigler (ed.), Proceedings of the Fifth Workshopof the IOBC Global Working Group “Quality Control of Mass Reared Arthro-pods”. International Organization for Biological Control of Noxious Animals

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

and Plants, Wageningen, Netherlands.

POTENTIAL OF FIELD CORN AS A BARRIER CROP AND EGGPLANT AS A TRAP CROP FOR MANAGEMENT OF

BEMISIA ARGENTIFOLII

(HOMOPTERA: ALEYRODIDAE) ON COMMON BEAN IN NORTH FLORIDA

H

UGH

A

DAM

S

MITH

AND

R

OBERT

M

C

S

ORLEY

Department of Entomology and NematologyUniversity of Florida

Gainesville, FL, 32611

A

BSTRACT

Trap crops and barrier crops are among the cultural control methods promoted formanagement of

Bemisia argentifolii

Bellows & Perring, particularly for small farmersin the tropics. In 1996 eggplant,

Solanum melongena

L., was tested as a trap crop, andin 1996 and 1997 corn,

Zea mays

L., was tested as a barrier crop for management of

B. argentifolii

on bean,

Phaseolus vulgaris

L. In 1996 treatments were compared bysampling immature

B. argentifolii

on bean leaves. Neither egg nor nymphal densitieswere reduced by eggplant or corn treatments in 1996. In the 1997 corn barrier trialplot size was increased and the orientation of barrier row to wind direction was eval-uated. A dust-and-release procedure was used to measure entry of greenhouse-rearedadult

B. argentifolii

into experimental plots. Counts from yellow sticky traps in 1997indicated that migration by adult whiteflies into plots was determined primarily byair currents and was only marginally influenced by the presence of a corn barrier. Theresults indicate that barrier crops and certain trap crops may have limited value forwhitefly management.

Key Words: Intercropping, polyculture, vector management, wind dispersal, pestmanagement

146


83(2) June, 2000

R

ESUMEN

El uso de cultivos trampa y cultivos de barrera se está promoviendo como medidade control de la mosquita blanca (

Bemissia argentifolii

Bellows & Perring), principal-mente entre pequeños agricultores de los trópicos. En 1996 se probó el cultivo de be-renjena (

Solanum melongena

L.) como planta trampa y en 1996 y 1997 se utilizó maíz(

Zea

mays

L.) como cultivo barrera para el control de

B

.

argentifolii

en un campo de fri-jol,

Phaseolus vulgaris

L. En 1996 se colectaron muestras de

B

.

argentifolii

en fase in-madura de hojas de frijol para comparar el efecto de los tratamientos. En 1996 no selogró reducir la densidad de huevecillos o ninfas en el frijol al emplear berenjena omaíz. En 1997 se aumentó el tamaño de la parcela experimental de maíz y se evaluó elefecto de la orientación del surco barrera en relación a la dirección del viento. La can-tidad de adultos de

B

.

argentifolii

que entraron a los lotes experimentales se cuantificómediante un procedimiento de espolvoreo y liberación (dust-and-release). En 1997 seencontró que la migración de adultos de mosquita blanca hacia los lotes experimenta-les fue determinada principalmente por corrientes de aire y que la presencia de maízbarrera tuvo muy poco efecto en controlar su entrada. Los resultados indican que los

cultivos barrera y trampa fueron poco efectivos en el control de la mosquita blanca.


Bellows & Perring, also known as the B strain of

B. tabaci

(Gennadius), causes significant economic damage to agronomic and horticulturalcrops throughout warm regions of the world (Brown et al. 1995).


is a phloem-feeder which vectors numerous geminiviruses and inflicts a variety ofplant disorders as well as mechanical damage (Byrne et al. 1990, Hiebert et al. 1996,Shapiro 1996). It has demonstrated resistance to most classes of pesticides (Denholmet al. 1996), forcing growers and researchers to evaluate alternative methods of con-trol. Attempts to manage whiteflies by cultural means have included the use of trapcrops (Al-Musa 1982, Ellsworth et al. 1994, McAuslane et al. 1995, Schuster et al.1996) and barrier crops (Sharma & Varma 1984, Fargette & Fauquet 1988, Rataul etal. 1989, Morales et al. 1993).

Trap crops are preferred host plants which are used to draw an herbivore awayfrom a less-preferred main crop (Vandermeer 1989).


has been ob-served to oviposit heavily on eggplant,

Solanum melongena

L. (Tsai & Wang 1996),leading researchers to suggest eggplant as a promising trap crop (Faust 1992).

Whiteflies are weak fliers, relying on air currents for both short and long distancemigration (Byrne & Bellows 1991, Byrne et al. 1996). Several tall-growing non-hostplants, primarily in the family Gramineae, have been tested as barrier crops or inter-crops to reduce whitefly colonization and virus transmission among main crops. Re-sults have been mixed. Morales et al. (1993) reported that a sorghum,

Sorghumbicolor

(L.) Moench, barrier reduced

B.

tabaci

densities and transmission of virus, ontomatoes,

Lycopersicon esculentum

Mill. A pearl millet,

Pennisetum typhoides

(Burm.f.) Stapf & Hubbard, barrier reduced whitefly virus transmission on cowpea,

Vignaunguiculata

(L.) Walp. (Sharma & Varma 1984) and on soybean,

Glycine max

(L.) Mer-rill (Rataul et al. 1989). Gold et al. (1990) found reduced densities of

Aleurotrachelissocialis

Bondar and

Trialeurodes variabilis

(Quaintance) on cassava,

Manihot escu-lenta

Crantz, intercropped with maize,

Zea mays

L., and cowpea, but attributed thisin part to reduced host quality due to intercrop competition. Fargette & Fauquet(1988), whose study included the effect of wind direction, found densities of

B. tabaci

and virus incidence were sometimes higher on cassava intercropped with maize thanon monocropped cassava.


147

These studies have been carried out primarily in the tropics, where safe, inexpen-sive cultural control measures are a priority for low resource farmers. Extension ma-terial from Central America promotes the use of crop barriers as a component ofwhitefly management programs (Salguero 1993). The present study was undertakenin 1996 to test the usefulness of eggplant as a trap crop and field corn as a barrier cropfor management of

B. argentifolii

on common bean,

Phaseolus vulgaris

L. It was con-tinued in 1997 focusing only on the barrier crop treatment and including the effectsof wind direction and barrier row orientation.

M

ATERIALS

AND

M

ETHODS

Research Design and Plot Management, 1996

The experiment was carried out at the University of Florida Green Acres AgronomyResearch Farm, northwest of Gainesville, FL (29

°

40’N, 82

°

30’W). Four treatmentswere compared: 1) bean planted in monoculture, 2) bean intercropped with eggplant,3) bean intercropped with field corn, and 4) bean monoculture treated with imidaclo-prid (Provado 1.6F, Bayer, Kansas City, MO), a systemic insecticide. The imidaclopridtreatment was included for yield comparison only. It was not sampled for whiteflies.

‘Espada’ bean (Harris Seed, Rochester, NY) was used in the monoculture and inter-crop treatments. ‘Black Beauty’ eggplant (Ferry-Morse Seed, Fulton, KY) was testedas a trap crop in one intercrop treatment. The subtropical field corn hybrid Howard II-IST (Gallaher et al. 1998) was tested as a barrier crop in the other intercrop treat-ment. Plant spacing within the row was 10 cm for bean, 15 cm for corn, and 46 cm foreggplant. Each plot contained 14 rows which were 6.1 m in length with 0.9 m betweenrows. Intercropped plots were planted in a 2:4:2:4:2 pattern, with corn or eggplant inthe outermost and central 2 rows, surrounding 2 four-row patches of bean. Each treat-ment was replicated 5 times and arranged in a randomized complete block design.

Corn was planted on 26 July and fertilized with 0.68 kg 15-0-14 (N-P

2

O

5

-K

2

O) perrow. Corn received 0.3 kg 15-0-14 per row on 9 August. Heavy

Spodoptera frugiperda

(J. E. Smith) damage threatened the barrier crop treatment in August. Corn wastreated with 1.74 liter/ha methomyl (Lannate, DuPont Corp., Wilmington, DE) on 9August and 29 August. Eggplant was transplanted on 22 August when 3 wks old. Egg-plant received 0.23 kg per row 15-0-14 fertilizer 27 August, and 0.8 kg on 27 Septem-ber and 10 October. Beans were planted on 15 September and fertilized with 0.37 kg15-0-14 per row on 23 September and 12 October.

The experimental area was treated with 0.19 liter/ha paraquat (Gramoxone, Zen-eca) on 26 July. Subsequent weed control was mechanical or by hand. The imidaclo-prid-treated beans received 52.6 g/ha ai imidacloprid (Provado 1.6 F, Bayer) on 4October and 12 October. This is the rate recommended on the label for most vegetables(Bayer, Kansas City, MO). Provado 1.6F was applied with a backpack sprayer. Imida-cloprid is not registered for use on beans but was included so that yield from inter-cropping treatments could be compared with yield from chemically-protected beans.

Sampling

Whole plant examinations were made of 1 or 2 bean plants per plot each week from22 September through 11 November except for 29 September. Only the underside ofthe leaf was examined. The area of each leaf was recorded using a LI-COR portableleaf area meter (model LI-3000A, LI-COR, Lincoln, NE). Bean treatment comparisonswere made on the basis of whole plant counts. Leaf counts from upper, middle, and

148


83(2) June, 2000

lower plant strata were used for comparison with eggplant on 21 October and 4 No-vember. On 29 September bean and eggplant comparisons were based on the averageof counts taken from one 3.35 cm

2

disc from a leaf in the upper and lower strata of twoplants per plot (McAuslane et al. 1995).

Whole plant examinations were made of 1 to 3 eggplants per block each week from25 August through 8 October. After that time, plants became too large for whole plantexaminations. Whole leaf counts from upper, middle, and lower strata were made ofeggplant on 21 October and 4 November.

Leaves were examined using a stereoscope and fiber-optic light. Total number of

B.argentifolii

eggs, nymphs, parasitized nymphs, and red-eyed nymphs (also called ‘pu-pae’) was recorded for each leaf. Leaves with nymphs showing symptoms of parasit-ism were placed in unwaxed cylindrical 0.95 liter cardboard cartons (Fonda Group,Union, NJ) to allow parasitoids to emerge.

The height of five corn plants per row was measured on 4 October to assess the bar-rier effect. Beans were harvested from two 1.8-m sections from each plot on 22 No-vember and fresh weight was recorded.

Research Design and Plot Management, 1997

In 1997 the corn barrier treatment was repeated on a larger scale. Three treat-ments were compared to evaluate the influence of the barrier crop and the effect ofbarrier row orientation to wind direction on adult whitefly movement. Prevailing windsin August in the area tend to be from the east. The treatments were 1) bean plantedin monoculture (bean alone), 2) alternating rows of bean and corn planted north tosouth (barrier), and 3) alternating rows of bean and corn planted east to west (open).

Treatments were arranged in a randomized complete block strip split plot design.Each treatment was replicated four times. The four blocks were arranged in pairs oneither side of a 12 m-wide path running north to south. Treatment plots were 15.25 m

´

30.5 m, with the shorter side parallel to the central path. This design was used toallow for a release of whitefly adults from points spaced evenly along the central path.

Corn was planted on 25 March. It was fertilized with 67 kg/ha 15-0-14 (N-P

2

O

5

-K

2

O) on 1 April, 26 April, and 14 May. Bean was planted on 1 July and fertilized with33 kg/ha 15-0-14 (N-P

2

O

5

-K

2

O) at planting, 10 July, and 20 July. Overhead irrigationwas used to supplement rainfall. Plots were weeded mechanically and by hand.

Mass-rearing of

B. argentifolii

About 30 senescing broccoli,

Brassica olerecea

L., plants infested with

B. argenti-folii

were removed from an organic farm near Gainesville between 1-6 June. Theywere potted and placed with 36 flowering hibiscus,

Hibiscus rosa-sinensis

L., plants ina greenhouse at the Department of Entomology and Nematology at the University ofFlorida. Hibiscus plants were watered regularly and fertilized with Purcell’s Sta-Green® plant food (18-6-12 N-P

2

O

5

-K

2

O, Purcell Industries, Sylacauga, AL). By earlyAugust, the hibiscus plants were heavily infested with whiteflies.

Trap Preparation

Yellow sticky traps have been used in many instances to monitor and samplewhitefly adults (Ekbom & Xu 1990). In the evening of 7 August 180 plastic yellow 710-ml cups (Solo Cup Company, Urbana, IL) were coated with an aerosol adhesive (prod-uct 95010, Tanglefoot Company, Grand Rapids, MI) for use as whitefly traps. Thetraps were arranged in 5 rows within each plot at 1.5, 7.6, 14, 20, and 26 m from the


149

edge of the plot bordering the central path. Three traps were placed in each row. Onetrap was placed 3.8 m in from either side of the plot, and one was placed 7.6 m withinthe plot, at the center of the row.

Dust-and-release Procedure

Byrne et al. (1996) developed a method of dusting whitefly adults with a fluores-cent pigment in the field and trapping them at a distance as a means to monitor move-ment. We modified this method to distinguish the released whitefly adults from thetrapped field population.

Before dawn on 8 August the infested hibiscus plants were enclosed in 113.5 literplastic leaf litter bags. The nozzle of a technical duster (product 1964, Lesco, Cleve-land, OH) was forced through the plastic and approximately 8.5-14 g orange fluores-cent AX-14-N pigment (Fire Orange, Day-Glo Color, Cleveland, OH) was puffed fromthe duster into the bag onto the infested plants. The hibiscus plants were transportedto the experimental area enclosed in plastic bags and arranged in 6 clusters of 6 plantsalong the central path and between pairs of treatment plots. The plastic bags were re-moved between 7:30 and 7:50 AM to allow a unified release of dyed whitefly adults.The traps were removed and replaced at dusk. The second set of traps was removedat dusk on 9 August. After removal, traps were kept refrigerated until examined.

On 10 August, the hibiscus plants were returned to the greenhouse. Traps wereplaced in the plots from 8:00 AM to 5:00 PM on 14 August to determine that whiteflyadults from the first release were no longer measurably present in the area. On 24August the dust-and-release procedure was repeated. Traps were set out from 8:00 AMto 8:00 PM on 24 August, and replaced with traps that were recovered at dusk on 25August. Hibiscus plants were removed after the second set of traps had been retrieved.

Traps were examined using a Spectroline 365 nm black light (model B-14N, Spec-tronics, Westbury, NY). The number of fluorescing whitefly adults on each trap was re-corded.

The height of 15 corn plants per plot was measured on 27 August to evaluatethe barrier effect.

Statistical Analysis

In 1996, densities of

B. argentifolii

eggs, nymphs, parasitized nymphs, and red-eyed nymphs were compared among bean treatments using analysis of variance(PROC GLM, SAS Institute 1996). Densities of whitefly immatures on bean and egg-plant in the trap crop test were compared using the same test, as was bean yield. Forthe 1997 study, the effect of treatment, block, and trap position on trap count was an-alyzed using analysis of variance. Orthogonal contrasts were then used to comparetrap counts in the same treatment east and west (upwind and downwind) of the re-lease point, and to compare trap counts among treatments in blocks west of the re-lease point. Wind direction data collected at the site were provided by Dr. E. B. Whitty,Agronomy Department, University of Florida, Gainesville, FL.

R

ESULTS

AND

D

ISCUSSION

Whitefly Densities, 1996

Densities of eggs were highest on bean when sampling began and declined oversubsequent weeks (Table 1). Nymphal densities were highest during weeks 3 and 4.Observations of parasitized nymphs and red-eyed nymphs were low throughout, al-though parasitism increased slightly over time.

150

Florid

a En

tomologist

83(2)Ju

ne, 2000

T

ABLE

1. M

EAN

(

±

SD)

NUMBER

OF

IMMATURE

B

EMISIA

ARGENTIFOLII

/

CM

2

FOLIAGE

ON

BEAN

, 1996.

Date Treatment Egg Nymph Para. nymph

1

REN

2

22 Sept. Bean aloneBean w/ cornBean w/ eggplant

0.79

±

0.581.04

±

0.731.27

±

0.68

000

000

000

8 Oct. Bean aloneBean w/ cornBean w/ eggplant

0.62

±

0.400.93

±

0.261.00

±

0.58

0.64

±

0.290.86

±

0.331.31

±

0.87

00.002

±

0.0040.006

±

0.01

000.004

±

0.008


0.40

±

0.300.67

±

0.490.60

±

0.27

0.79

±

0.301.10

±

0.650.80

±

0.25

0.010

±

0.0080.010

±

0.0040.004

±

0.005

0.010

±

0.020.002

±

0.0040


0.36 ± 0.200.39 ± 0.100.44 ± 0.15

0.48 ± 0.300.80 ± 0.510.61 ± 0.33

0.006 ± 0.0050.016 ± 0.0150.006 ± 0.008

0.006 ± 0.0050.008 ± 0.0130.004 ± 0.005


0.41 ± 0.340.43 ± 0.160.46 ± 0.14

0.46 ± 0.230.58 ± 0.220.67 ± 0.26

0.004 ± 0.0050.020 ± 0.0150.010 ± 0.007

0.012 ± 0.0110.018 ± 0.0160.016 ± 0.011

4 Nov. Bean aloneBean w/ cornBean w/ eggplant

0.54 ± 0.580.22 ± 0.180.34 ± 0.32

0.51 ± 0.260.44 ± 0.150.41 ± 0.22

0.010 ± 0.010.016 ± 0.0150.036 ± 0.027

0.010 ± 0.0100.016 ± 0.0130.014 ± 0.008

11 Nov. Bean aloneBean w/ cornBean w/ eggplant

0.26 ± 0.060.06 ± 0.040.11 ± 0.16

0.45 ± 0.350.31 ± 0.200.33 ± 0.24

0.046 ± 0.0490.052 ± 0.0430.024 ± 0.018

0.006 ± 0.0080.008 ± 0.0080.002 ± 0.004

1Parasitized nymphs.2Red-eyed nymphs.


There were no differences (p > 0.10) in egg density among treatments during thefirst six weeks of sampling. Egg densities on bean alone were higher (p < 0.05) thanon bean intercropped with corn or eggplant during weeks 7 and 8. No differences (p >0.10) in nymphal densities occurred among treatments. Densities of red-eyed nymphswere higher (p < 0.05) on bean alone than on the corn and eggplant treatments duringweek 4. During week 7, parasitism was more than twice as high in the eggplant treat-ment as in the other two treatments.

Whitefly adults were observed on eggplant the day following transplanting on 22August. Egg densities on eggplant foliage ranged from 0.66 ± 0.46/cm2 on 25 Augustto 3.53 ± 0.72/cm2 on 16 September, and declined over the following weeks. Nymphaldensities on eggplant foliage were 1.31 ± 1.60/cm2 on 1 September and peaked at 2.39± 0.33 on 16 September, declining on subsequent sampling dates. When bean plantswere emerging, eggplants were quite large; they had an average of 7.0 ± 1.3 branches,a mean height of 17.33 ± 0.28 cm, and mean leaf area of 485 ± 156 cm2 (n = 5).

Bean vs. Eggplant

On all sampling dates after the first week, egg densities were higher (p < 0.05) onbean than on eggplant (Table 2). During the week that nymphs were first observed onbean, densities were lower (p < 0.05) on bean than on eggplant. During subsequentsampling dates, nymphal densities were either higher (p < 0.05) on bean or not statis-tically different. Observations of parasitized and red-eyed nymphs were either higheron eggplant than on bean or not statistically different on the two hosts.

Parasitoid Species

All parasitoids reared from bean and eggplant were hymenopterans from the familyAphelinidae. Thirty-nine parasitoid individuals were recovered from bean leaves. Thirty-two of these were Encarsia nigricephala Dozier (82%), 4 were Eretmocerus sp. (10.3%),and 3 were Encarsia pergandiella Howard (7.7%). Among the 121 parasitoid individualsreared from eggplant leaves, 51 were E. pergandiella (42.1%), 48 were E. nigricephala(39.7%), 13 were Eretmocerus sp. (10.7%), 6 were E. transvena (Timberlake) (5%), and 3were Encarsia sp. (2.5%). The greater parasitism and variety of parasitoid species on egg-plant may be due to the greater number of weeks that eggplant was in the field.

Bean Yield

There was an average of 37.30 ± 5.88 bean plants per 3.6 m of row in all treat-ments. Bean yield per 3.6 m of row was not different among the three treatments andthe imidacloprid-treated bean plants (imidacloprid: 0.95 kg ± 0.71; bean: 0.87 kg ±0.58; corn: 0.47 kg ± 0.28; eggplant: 1.14 kg ± 0.77).

Eggplant as a Trap Crop

Eggplant did not reduce oviposition on adjacent bean early in the season, and sodid not function as a trap crop. Oviposition was not consistently higher on eggplantthan on bean as reported elsewhere (Tsai & Wang 1996). Eggplant leaves may havebeen less suitable for oviposition because they were several weeks older than the beanleaves. A concurrent test of squash, Cucurbita pepo L., as a trap crop for whitefliesalso produced negative results (Smith 2000).

It is possible that host-finding mechanisms used by whitefly adults prevent themfrom being drawn away from one host plant by the presence of another. Bemisia tabaci

152F

lorida E

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ologist 83(2)Ju

ne, 2000

TABLE 2. IMMATURE BEMISIA ARGENTIFOLII (MEAN ± SD/CM2 FOLIAGE) ON BEAN AND EGGPLANT, 1996.

Date

Egg Nymph Parasitized nymph Red-eyed nymph

Bean Eggplant Bean Eggplant Bean Eggplant Bean Eggplant

22 Sept. 1.66 ± 1.67 2.74 ± 1.72 0 1.84 ± 1.72* 0 0 0 0

29 Sept. 5.52 ± 3.44 1.68 ± 1.72*1 0.88 ± 0.62 2.13 ± 1.78* 0 0 0 0.031 ± 0.104

8 Oct. 0.65 ± 0.31 0.24 ± 0.41* 1.59 ± 0.83 0.29 ± 0.19* 0.005 ± 0.016 0.035 ± 0.037* 0.005 ± 0.016 0.012 ± 0.015

21 Oct. 0.64 ± 0.54 0.23 ± 0.22* 0.45 ± 0.35 0.49 ± 0.65 0.009 ± 0.014 0.025 ± 0.038 0.003 ± 0.009 0.046 ± 0.078

4 Nov. 0.26 ± 0.26 0.02 ± 0.03* 0.28 ± 0.19 0.11 ± 0.10* 0.024 ± 0.043 0.069 ± 0.067 0.006 ± 0.012 0.048 ± 0.043*

1*Indicates that numbers on bean and eggplant are significantly different on a given date according to analysis of variance at a = 0.05.


apparently does not respond to host-specific visual or olfactory cues (Mound 1962).Elucidation of the precibarial and cibarial chemosensilla of B. tabaci by Hunter et al.(1996) indicates that B. tabaci may be able to evaluate plant sap before ingesting it.It has been demonstrated that Trialeurodes vaporariorum (Westwood) relies on gus-tatory information to accept or reject a host (van Lenteren & Noldus 1990), and thismay also be true for B. argentifolii. In addition, whitefly adults tend to leave some hostplant species more quickly than others (Costa et al. 1991, Verschoor-van der Poel1978). The observed differences in host-specific oviposition density by B. argentifoliimay be due in part to length of tenure on the plant rather than to some preference ex-pressed in the host-finding stage (Bernays 1999).

Many trap crop studies have not resulted in consistent reductions of whitefly den-sities on the main crop (Ellsworth et al. 1994, McAuslane et al. 1995, Perring et al. 1995,Puri et al. 1996, Schuster et al. 1996). However, Al-Musa (1982) and Schuster et al.(1996) reported a reduction in virus incidence on tomato using cucumber, Cucumis sa-tivus L., and squash, respectively, as trap crops. Power (1990) suggests that crop com-binations which cause virus vectors to probe for briefer periods may reduce theincidence of persistent viruses such as geminiviruses. Bernays (1999) demonstratedthat B. tabaci tends to move more often and spend less time on certain plants whenthey were grown in combination than when they were grown in pure stands. The cropcombinations and densities employed by Al-Musa (1982) and Schuster et al. (1996)may have led to reduced probing by the vector, and so reduced incidence of virus.

Corn as a Barrier Crop

The corn did not grow well in 1996 due to insufficient fertilizer. It attained a meanheight of 1.18 m ± 0.34 (n = 150) and a density of 27 ± 7 plants per 6.1m row (n = 30).We re-evaluated the barrier effect in 1997 with larger, properly fertilized plots. Egg-plant did not appear to be a promising trap crop, and so was not included in the fieldexperiment the following year.

Release of Adult Whiteflies, 1997

Average corn height was 2.45 ± 1.97 m when whitefly releases were made. The ef-fect of treatment on trap count was not significant (p > 0.10) on any of the four collec-tion dates. Wind direction was from the east or northeast during the 4 days thatcollections were made (Table 3). Trap counts in plots to the west of the release pointwere significantly higher than trap counts in plots to the east of the release point foreach treatment on each collection date (Table 3). When treatments were compared onthe basis of downwind plots only, counts were lower (p < 0.05) in the barrier treatmentthan the monocropped bean treatment on 9 August and 25 August. Trap counts werelower in the downwind barrier plots than in the downwind ‘open’ plots on 24 August(p < 0.05) and 25 August (p < 0.1) (Table 3).

Wind direction appeared to be the primary factor determining where whiteflyadults were trapped. This is consistent with observations that whitefly adults movepassively with wind currents as ‘aerial plankton’ (Byrne & Bellows 1991). Amongdownwind plots, the barrier treatment tended to have the lowest counts, indicatingthat the arrangement of corn rows perpendicular to the prevailing wind direction didhave some effect on the movement of adults within the plot. However the overall trapcounts in this study were low. The contribution made by corn barriers to reducingwhiteflies may depend on the density of the whitefly population. Crop barriers such ascorn may be more effective when used with other control measures. Short of employ-

154F

lorida E

ntom

ologist 83(2)Ju

ne, 2000

TABLE 3. WHITEFLY ADULTS (MEAN ± SD) PER TRAP UNDER 3 CROPPING SYSTEMS, AUGUST 1997.

Date Row1

Bean alone Corn: barrier to wind Corn: open to wind

Downwind Upwind Downwind Upwind Downwind Upwind

Release 12

8 Aug. 1 1.67 ± 2.25 0.33 ± 0.52 2.33 ± 1.03 0.33 ± 0.52 2.33 ± 1.21 0.50 ± 0.842 1.33 ± 1.97 0 1.00 ± 0.63 0 0.33 ± 0.52 0.17 ± 0.413 0.67 ± 0.52 0 1.33 ± 1.51 0 0.17 ± 0.41 0.17 ± 0.414 0.50 ± 0.55 0.33 ± 0.33 ± 0.52 0 0.67 ± 0.82 05 0.33 ± 0.52 0 0.17 ± 0.41 0.16 ± 0.41 0.67 ± 1.03 0x–1 0.90 ± 1.40 0.13 ± 0.35*3 1.03 ± 1.16 0.10 ± 0.31* 0.83 ± 1.12 0.17 ± 0.46*

9 Aug. 1 2.00 ± 1.79 0.17 ± 0.41 1.17 ± 0.75 0 1.83 ± 1.47 0.33 ± 0.522 1.67 ± 0.82 0.33 ± 0.52 1.00 ± 0.89 0.17 ± 0.41 1.17 ± 1.17 03 1.50 ± 1.22 0 0.83 ± 0.98 0 0.67 ± 1.21 04 0.50 ± 0.55 0 0.50 ± 0.84 0 1.00 ± 0.89 05 0.50 ± 0.84 0.17 ± 0.41 0.67 ± 0.82 0 0.50 ± 0.84 0x–1 1.23 ± 1.22a4 0.13 ± 0.35* 0.83 ± 0.83b 0.03 ± 0.18* 1.03 ± 1.16ab 0.07 ± 0.25*

Release 22

24 Aug. 1 3.00 ± 2.00 0.33 ± 0.52 2.83 ± 3.25 0 3.50 ± 2.17 0.17 ± 0.412 1.67 ± 1.21 0.17 ± 0.41 1.50 ± 1.05 0.17 ± 0.41 2.50 ± 1.05 0.17 ± 0.413 0.83 ± 0.75 0.17 ± 0.41 0.50 ± 0.84 0 1.83 ± 1.33 04 0.50 ± 0.55 0.17 ± 0.41 0.17 ± 0.41 0 1.17 ± 0.41 0

1Row refers to trap location (1 = nearest, 5 - farthest from release point; see text). (x– = mean across all 5 row locations.)2Wind direction on release dates: 8 Aug.: 75°; 9 Aug.: 97°; 24 Aug.: 61°; 25 Aug.: 55°.3*Indicates mean trap counts in the same treatment upwind and downwind of the release point are significantly different at p < 0.05 according to F-test for contrasts.4Different letters indicate that mean trap counts among treatments downwind of release point are significantly different at p < 0.05 according to F-test for contrasts.

Sm

ith & M

cSorley: T

rap and Barrier C

rops for Whitefly

155

5 0.33 ± 0.52 0.17 ± 0.41 0 0 1.33 ± 1.21 0x–1 1.27 ± 1.46b 0.20 ± 0.41* 1.00 ± 1.82b 0.03 ± 0.18* 2.10 ± 1.52a 0.07 ± 0.25*

25 Aug. 1 3.33 ± 1.97 0.17 ± 0.41 0.33 ± 0.52 0 1.17 ± 1.17 02 1.00 ± 1.10 0 1.00 ± 1.10 0 1.00 ± 0.89 03 1.17 ± 0.98 0 0.17 ± 0.41 0 0.50 ± 0.55 04 0.67 ± 0.82 0 0.33 ± 0.82 0 0.83 ± 1.17 05 0.33 ± 0.52 0 0.17 ± 0.41 0 1.00 ± 0.89 0x–1 1.30 ± 1.53a 0.03 ± 0.18* 0.40 ± 0.72b 0 0.90 ± 0.92 0*

TABLE 3. (CONTINUED)WHITEFLY ADULTS (MEAN ± SD) PER TRAP UNDER 3 CROPPING SYSTEMS, AUGUST 1997.

Date Row1

Bean alone Corn: barrier to wind Corn: open to wind

Downwind Upwind Downwind Upwind Downwind Upwind

1Row refers to trap location (1 = nearest, 5 - farthest from release point; see text). (x– = mean across all 5 row locations.)2Wind direction on release dates: 8 Aug.: 75°; 9 Aug.: 97°; 24 Aug.: 61°; 25 Aug.: 55°.3*Indicates mean trap counts in the same treatment upwind and downwind of the release point are significantly different at p < 0.05 according to F-test for contrasts.4Different letters indicate that mean trap counts among treatments downwind of release point are significantly different at p < 0.05 according to F-test for contrasts.


ing manufactured barriers such as floating row covers or fine mesh screens, whiteflyadults probably cannot be excluded from a cropped area (Norman et al. 1993).

Trap position had a significant effect on trap count (ANOVA, d.f. = 4; 8 August, F =4.67, p < 0.05; 9 August, F = 4.65, p < 0.05; 24 August, F = 2.99, p < 0.1; 25 August,F = 2.86, p < 0.1). The number of whiteflies caught decreased as trap distance from therelease point increased. The interaction of treatment and trap position interactionwas not significant, suggesting that this decline was not different among treatments.

Data derived from attractive traps may be ambiguous. A gravid or hungry whiteflyadult which is surrounded by non-hosts, such as corn, may be more sensitive to a dis-tant patch of bright yellow than an adult in similar condition surrounded by accept-able hosts, such as bean. It is conceivable that the whitefly adults in the corntreatments spent more time searching and so were drawn from a greater area thanthe whitefly adults trapped in the monocropped bean treatments. It is possible thatfewer whitefly adults entered the corn treatments than the monocropped bean, butthat a higher proportion of those entering the corn treatments were trapped. How-ever, these considerations do not alter the overall impression that where air currentscan enter, whitefly adults can follow.

ACKNOWLEDGMENTS

The authors would like to thank to R. Wilcox, without whom this research wouldnot have been possible. We are grateful to D. C. Beitrusten, D. W. Dickson, R. N. Gal-laher, and C. W. Scherer for field assistance, and to J. L. Stimac for sampling advice.G. A. Evans kindly identified whitefly parasitoids, and J. M. Harrison gave invaluablestatistical guidance. We thank E. B. Whitty for wind direction data. Thanks to R. N.Gallaher and H. J. McAuslane for improving earlier versions of this manuscript. Thisresearch was made possible by support from the Ruth Freeman and Anselm FisherFoundation. Florida Agricultural Experiment Station Journal Series No. R-06937. Noendorsements or registrations are implied herein.

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Davidson et al.: Bacteria in

Bemisia argentifolii 159

CULTURABLE BACTERIA ASSOCIATED WITH THE WHITEFLY,

BEMISIA ARGENTIFOLII

(HOMOPTERA: ALEYRODIDAE)

E

LIZABETH

W. D

AVIDSON

1,

R

OSEMARIE

C. R

OSELL

2

AND

D

ONALD

L. H

ENDRIX

3

1

Department of Biology, Arizona State University, Tempe, AZ 85287-1501

2

Department of Biology, University of St. Thomas, Houston, TX 77006

3

USDA-ARS, Western Cotton Research Laboratory, Phoenix, AZ 85040

A

BSTRACT

Several different types of bacteria were cultured from surface-sterilized

Bemisiaargentifolii

Bellows, Perring, Gill and Hedrick 1994 (Homoptera: Aleyrodidae) adultsand nymphs, including

Bacillus

spp., Gram-variable pleomorphic rods and Gram-pos-itive cocci. Two of the isolates were capable of being ingested by adults and passed intothe honeydew. One of these,

Enterobacter cloacae

, was found within the gut cells ofadult whiteflies and was mildly pathogenic. This isolate represents the first bacte-rium with potential as a pathogen of whiteflies. Bacteria which were not capable of be-ing ingested, may have been located in structures which were protected from surfacesterilization, such as the lingula or the female reproductive tract.

Key Words:

Bemisia tabaci

B-biotype,


,

Bacillus

sp., symbioticbacteria

R

ESUMEN

Diferentes tipos de bacterias fueron aisladas de adultos y ninfas de

Bemisia argen-tifolii

Bellows et al., 1994 (Homoptera: Aleyrodidae) esterilizados superficialmente.Entre las bacterias aisladas se encontró

Bacillus

spp., bacilos pleomórficos Gram-va-riables y cocos Gram-positivos. Los adultos fueron capaces de ingerir a dos de los or-ganismos aislados y de transferirlos a la mielecilla que secretan. Uno de éstos,


, fué encontrado dentro de células intestinales de mosquitasblancas adultas y fué moderadamente patogénico. Esta bacteria representa el primerorganismo aislado que posee potencial patogénico para el control de la mosquitablanca. Otras bacterias aisladas no fueron capaces de ser ingeridas, lo cual se atribuyóa que se ubicaron en estructuras protegidas de la esterilización superficial, como la lí-

gula o el tracto reproductivo femenino.

The silverleaf whitefly,


(=

B. tabaci

B biotype) (Bellows, Perring,Gill and Hedrick, 1994), is one of the most damaging insects to agriculture in the south-ern United States and in warmer regions of many other countries.

B. argentifolii

feedson a wide variety of plant species, including cotton, melons, brassicas, tomato, peppers,and ornamentals such as poinsettia (Cock 1986, De Barro 1995). This insect causes eco-nomic damage in four ways: by removing photosynthetic products during phloem feed-ing; by contaminating cotton fiber and ornamental plants with sticky excretedhoneydew; by inducing physiological responses in the host such as squash silverleaf andtomato irregular ripening; and by vectoring viruses (Byrne & Bellows 1991).


possesses pleomorphic and coccoid obligate intracellular bac-terial endosymbionts, housed in mycetomes, which are transmitted from the female to

160


83(2) June, 2000

her eggs (Costa et al. 1993a). Growth and development of the nymph and induction ofthe squash silverleaf disorder, for which the species is named, are retarded if the sym-biotic bacteria are reduced or eliminated by feeding antibiotics to the adult female be-fore oviposition or if fed to the nymph via the leaf (Costa et al. 1993b, 1997). Theseendosymbionts have not reportedly been cultured on laboratory media.

The objectives of this study were to investigate whether other bacteria are presentin whiteflies, and modes of entry of these bacteria into the whiteflies. We have isolateda variety of bacteria from surface-sterilized

B. argentifolii

adults and nymphs, someof which have previously been implicated in the production of medium-length oli-gosaccharides in the honeydew (Davidson et al. 1994). Two of the isolated bacteriawere shown to be ingested by the whitefly and one of these was mildly pathogenic.

M

ATERIALS

AND

M

ETHODS

Whiteflies

Whitefly adults, nymphs and eggs were collected from cotton, cabbage, cucumber,squash, lantana, pepper or melon plants and surface sterilized with ethanol andhousehold chlorine bleach as described previously (Davidson et al. 1994). Because ofthe small size of these insects, samples were processed in groups of ca. 50-200. Adults,nymphs and eggs were processed separately. Surface sterilized insects were inocu-lated into liquid nutrient broth-yeast extract-salts medium (Myers and Yousten 1978)or brain heart infusion broth, either whole or homogenized in sterile 0.9% saline. Ho-mogenized insects were also plated directly on microbiological media including nutri-ent agar, brain heart infusion agar, tryptose agar, Luria agar, purple agar, andchocolate agar (Sigma, St. Louis, MO) and incubated aerobically at 25

°

C.

Bacterial Cultures

When bacterial growth was observed in liquid or agar bacterial media, cultureswere streaked for purity on agar media of the same type on which positive growth hadbeen observed. Cultures were preserved by freezing at -70

°

C in 20% glycerol in liquidmedium which best supported growth of each culture.

Bacteria were identified according to microscopic appearance, Gram’s stain,anaerobic growth, catalase production, colony morphology, and reactions on API 20Eand API CH identification strips (bioMerieux Vitek, Hazelwood, MO). Two isolates,designated WFA73 and WFN29, were also identified by gas chromatography (GC)fatty acid profiles using the MIDI identification system, by Dr. Joel Siegel, IllinoisNatural History Survey.

Whitefly Ingestion of Bacteria

Eight different bacterial strains, representative of the morphological and Gram-stain groups isolated from whiteflies (Table 3), were suspended in 30% sucrose andgreen food coloring at ca. 10

9

cells/ml, and fed to adult whiteflies through parafilm sa-chets. Control whiteflies were fed on sucrose alone. After 48 hr, mortality was re-corded, and all insects were surface sterilized, homogenized, and plated toappropriate agar medium. Sterile petri dishes were used to collect honeydew fromcontrol whiteflies and those fed bacteria. The dishes were rinsed with sterile salineand the saline plated to agar medium. Bacterial colonies recovered were comparedmicroscopically and in colony morphology to those originally fed to the whiteflies. Bio-



assays of strains WFA73 and WFN29 were repeated twice and % mortality reportedas the mean of two replicates; at least 100 insects were included in each treatment.

Electron Microscopy

Adult whiteflies were fed strain WFA73, which had been found to be ingested andmildly pathogenic in experiments described above, in green sucrose solution at ca.10

9

cells/ml in parafilm sachets. Control insects were fed on green sucrose only. After24 hr, insects with green digestive tracts were prepared for electron microscopy. In-sects were gently pierced in the thorax region while immersed in fixative, which con-sisted of 4% glutaraldehyde in 0.05M cacodylate buffer. Insects were fixed for 4 hr,postfixed in 0.5% osmium tetroxide in 0.05M cacodylate buffer for 1-2 hr, dehydratedin ethanol, and embedded in Spurr’s resin or LR White resin. Thin sections from 2 con-trol and 4 experimental whiteflies were observed by transmission electron microscopyusing a Philips EM 200 TEM (Eindhoven, Netherlands).

R

ESULTS

Bacteria from Whiteflies

Bacteria were cultured from surface-sterilized whitefly adults and nymphs collectedfrom all host plant species and from all groups of whiteflies collected at 30 differenttimes over a period of five years. A greater variety of bacteria was recovered from adultwhiteflies than from nymphs, however none were cultured from surface-sterilized eggs.A total of 80 isolates were preserved from adults and 29 from nymphs; representativeexamples are shown in Tables 1 and 2. There was no relationship between the hostplant species and the type of bacteria isolated from the whiteflies (Tables 1 and 2).

Four major types of bacteria were cultured from

B. argentifolii

using standard mi-crobiological media. These included: 1. Gram-positive sporeforming aerobic rods,

Ba-cillus

spp.; 2. Gram-positive cocci; 3. Gram-variable short pleomorphic rods producingvery short rods to cocci in older cultures; 4. Gram-variable long, thin, highly pleomor-phic rods forming cocci in older cultures (Tables 1 and 2).

There was a strong correlation between the insect life stage and the types of bac-teria isolated. Most isolates of Gram-positive or Gram-variable rod-shaped bacteriawere obtained from adult whiteflies, whereas most isolates of cocci were producedfrom nymphs (Tables 1 and 2).

Bacterial Identification

Sporeforming aerobic bacteria were identified as

Bacillus licheniformis

(Weig-man)

, B. megaterium

deBary

, B. amyloliquefasciens

Fukumoto

,

and

B. subtilis

(Ehrenberg), based upon microscopic examination and the results of API diagnostictests. The

B. subtilis,

B. licheniformis

and

B. megaterium

isolates were found to pro-duce medium-length sugars from sucrose in an earlier study (Davidson et al. 1994).

Spherical cells, forming diads and tetrads and occasionally chains, were frequentlyisolated from

B. argentifolii

nymphs. Isolates which were Gram-positive, catalase pos-itive, facultatively anaerobic, and capable of growing in the presence of 10% NaCl,were identified as

Staphylococcus

spp., close to

S. aureus

Rosenbach

, S. sciuri

Kloos,Schleifer & Smith and

S. epidermidis

(Winslow & Winslow). Larger gram-positivecocci forming diads and tetrads resembled

Sporosarcina

spp. (Orla-Jensen) althoughspores were not observed in these cultures (Tables 1 and 2).

162


83(2) June, 2000

On the basis of API tests, short, Gram-negative or Gram-variable pleomorphicrods were identified as


(Jordan),

Flavimonas oryzihabitans

Holmes et al.

, Citrobacter

sp. Workman and Gillen,

Cellulomonas

sp. Bergey et al.,

Chryseomonas luteola

Holmes et al.,

Acinetobacter lwoffsii

Brisou & Prevot or

A. bau-manni

(Deacon)

.

Some strains produced no reaction on API tests and therefore re-main unindentified (Tables 1 and 2). Two strains which were found to be ingested byadult whiteflies (below) were further identified by MIDI-GC.

Whitefly Ingestion of Bacteria

To determine whether bacteria isolated from whiteflies could have entered the in-sects during feeding, we fed eight different bacterial strains to adults (Table 3). Bac-teria strains designated WFA73 and WFN29 were recovered in large quantity fromsurface-sterilized adults and their honeydew following feeding on these bacteria. Bac-terial colonies, morphologically and microscopically identical to WFA73 or WFN29,were not found in homogenates or honeydew of control insects. WFA73 is a short ple-omorphic Gram-variable rod, identified by GC analysis of fatty acids and API analysis

T

ABLE

1. M

ORPHOLOGY

,

HOST

AND

IDENTIFICATION

OF

BACTERIA

RECOVERED

FROMADULT

B.

ARGENTIFOLII

. *I

DENTIFICATION

BY

MIDI;

ALL

OTHERS

IDENTIFIEDBY

API. ND =

NOT

DETERMINED

.

Designation Description Host Identification

WFA1 sporeforming rod cotton

Bacillus amyloliquefasciens

WFA2 spheres in diads cotton NDWFA3 sporeforming rod cucumber

B. licheniformis

WFA5 sporeforming rod cucumber

B. megaterium


B. licheniformis

WFA9, 10, 11, 12 sporeforming rod melon

Bacillus

spp.WFA12, 14, 35,36, 40, 45, 46 sporeforming rod cotton

Bacillus

spp.WFA37 short pleomorphic

rod, yellow colony cotton NDWFA41 cocci, diads cotton NDWFA43,44 small pleomorphic

rods, clear colonies cotton

Chryseomonas luteola

WFA52 diplococci, tetrads cotton NDWFA56, 59 very short rods,

thick at one end cotton

Acinetobacter lwoffsii

WFA67 short rods, yellowcolonies cotton

Citrobacter sp.

WFA68 short pointed rods cotton

Flavomonas oryzihabitans

WFA69, 70 short curved rods cotton

Acinetobacter baumanii?


Bacillus

sp.WFA 73 short pleomorphic rods cotton


*WFA74 short rod with inclusions cotton NDWFA80 short pleomorphic rods cotton

Flavomonas oryzihabitans



as

E. cloacae

(0.70 agreement). Isolate WFN29, which formed bent rods in young cul-tures and cocci in older cultures, was identified by GC and API analysis as

Cellulomo-nas (Oerskovia) turbata

(Erikson) (0.78 agreement).

Bacillus

spp., Gram-positivecocci and the other Gram-negative or Gram-variable isolates were not recovered fromadults fed these isolates (Table 3).

T

ABLE

2. B

ACTERIA

ISOLATED

FROM

B.

ARGENTIFOLII

NYMPHS

. M

ORPHOLOGY

,

HOSTAND

IDENTIFICATION

OF

BACTERIA

RECOVERED

FROM

ADULT

B.

ARGENTIFOLII

.*I

DENTIFICATION

BY

MIDI;

ALL

OTHERS

IDENTIFIED

BY

API. ND =

NOT

DE-TERMINED

.

Designation Description Host Identification

WFN1 spheres, diads cotton NDWFN3 sporeforming rod cotton

Bacillus

sp.WFN4 large diplococcus cotton

Sporosarcina

sp.?WFN6 sporeforming rod cabbage

Bacillus

sp.WFN7 coccus cabbage

Staphylococcus epidermidis

WFN10, 11, 13 cocci cabbage

Staphylococcus epidermidis

WFN12 cocci cabbage

Staphylococcus aureus

WFN14 large cocci cabbage

Sporosarcina

sp.?WFN15 short pleomorphic rods cabbage NDWFN17A very thin rods cabbage

Agromonas

sp.WFN28 sporeforming rod cabbage

Bacillus licheniformis

WFN29 pleomorphic rods forming cocci

cabbage

Cellulomonas turbata

*

T

ABLE

3. M

ORPHOLOGY

,

IDENTIFICATION

AND

RECOVERY

OF

BACTERIA

FED

TO

ADULT B.ARGENTIFOLII.

Straindesignation Morphology Identification

Recoveryfrom insect

or honeydew

WFA5 Gram-positive Bacillus Bacillus megaterium noWFA11 Gram-positive Bacillus Bacillus subtilis noWFN12 Gram-positive Coccus Staphylococcus aureus noWFN7 Gram-positive Coccus Staphylococcus epidermidis noWFA73 Short pleomorphic

Gram-variable rod Enterobacter cloacae yesWFN29 Pleomorphic

Gram-negative rod Cellulomonas turbata yesWFA69 Short curved or branched

rod Acinetobacter baumanii? noWFA74 Short rod with inclusions No API reaction, Gram-

variable no


Bioassay Results

Strain WFA73 (E. cloacae) produced an average 34% mortality (s.d. = 1.41) of adultwhiteflies at ca. 109 bacterial cells/ml after 24 hr in two experiments (control mortality= 4.0%; s.d. = 2.83). Mortality increased to 75% (s.d. = 0) after 48 hr in adult whitefliesfed WFA73 (control mortality = 9.5%; s.d.= 6.36). Bacteria identical to E. cloacae werealso recovered from honeydew of adults fed these bacteria but not from the honeydewof control adults fed only on sucrose. Strain WFN29 (C. turbata) produced only 4.5%mortality (s.d. = 2.12) of adult B. argentifolii at 109 cells/ml after 48 hr, similar to con-trol mortality (4.0%; s.d. = 1.41), but was recovered from homogenized adults fed thisstrain and from their honeydew.

Electron Microscopy

Bacteria were not observed in the digestive system of control whiteflies in thesestudies (not shown).

In adults fed strain WFA73 (E. cloacae), large numbers of bacteria were seenthroughout the digestive tract (Fig. 1). The lumen of the entire midgut was filled withrod-shaped bacteria (WFA73). Bacteria adhered to the apical portion of the descend-

Fig. 1. After feeding on bacteria, both the ascending and descending portions of themidgut of B. argentifolii contained high numbers of WFA73 (E. cloacae) bacteria (B)in the lumina (L). Thorax appears to the left and abdomen to the right. Note the pres-ence of engulfed bacteria (EB) in the apical portion of the epithelial cells. Nu, nucleusof midgut cell; FB, fat body; E, egg. Bar = 10 mm.

Davidson et al.: Bacteria in Bemisia argentifolii 165

ing and ascending midgut epithelial cells and were also found within the epithelialcell cytoplasm in membrane bound vesicles (Figs. 2 and 3). Cells in the descendingmidgut appeared to be taking up the bacteria by phagocytosis as evidenced by bacte-ria in various stages of engulfment (Figs. 3 and 4). Descending midgut cells of infectedinsects also exhibited poorly developed microvilli, numerous spherical vesicles, eachwith a small amount of electron dense material, many large electron dense lysosomal-like vesicles, and small electron dense residual bodies (Fig. 4). Vacuolation of mito-chondria was observed both in control and bacteria-fed whiteflies, probably a result ofslow fixative penetration.

We observed bacteria-like organisms in the reproductive tracts of two femalewhiteflies (Fig. 5). The bacteria in the female reproductive tracts were readily distin-guishable from sperm, which were observed in the spermatheca and appeared as elec-tron-dense structures which lacked notable internal structure, and were aligned inpackets (not shown). The identity of the bacteria found in the female reproductivetracts is not currently known.

DISCUSSION

Some of the bacteria cultured from whiteflies may be involved in a mutualistic re-lationship with the insects, contributing to the digestion and nutrition of the insect,while obtaining access to the high sugar content of phloem sap in the gut of the insectand honeydew. As described earlier (Davidson et al. 1994), Bacillus spp. associatedwith B. argentifolii may produce long-chain sugars which contribute to the stickinessof the honeydew of this insect. During a study of digestive tract ultrastructure (Rosellet al., unpubl.), bacteria were observed in the esophagus of adult Bemisia spp. takenfrom a laboratory colony reared on cotton.

Similar gut bacteria have been isolated from other Homoptera. Staphylococcus sci-uri and S. epidermidis, and Gram-negative rods, close to Pseudomonas fluorescensMigula, have been isolated from the pea aphid, Acythosiphon pisum (Harris) (Grenieret al. 1994). The flora in the aphid gut was assumed to have been acquired duringprobing on the leaf surface, as is probably the case with most of the bacteria isolatedfrom whiteflies as well. Srivastata and Rouatt (1963) isolated Sarcina, Micrococcus,Achromobacter and Flavobacterium from aphids. Bacteria have also been reported inthe hemocoel of aphids and leafhoppers (Grenier et al. 1994, Purcell et al. 1986). Atthis time we cannot rule out the possibility that some of the bacteria isolated from B.argentifolii are also occasional residents in the hemocoel.

We do not currently know the significance of bacteria present in the female repro-ductive tract. Morphologically, they resemble the short rod-shaped bacteria commonlyisolated from adult whiteflies (Table 1). Following electron microscope observation ofbacteria in the female reproductive tract, two collections of whiteflies were separatedinto males and females, surface sterilized and processed separately for bacterial iso-lation. Bacteria were cultured only from females. Their presence suggests possibletransmission of these bacteria to offspring, however bacteria were not cultured fromsurface-sterilized eggs.

The results presented here confirm that B. argentifolii adults and nymphs can in-gest certain culturable bacteria and contain these bacteria in their digestive tracts. Asthese bacteria were ingested through Parafilm, and were recovered from honeydew af-ter feeding, the bacteria were truly ingested and not simply resident on the externalsurface or mouthparts of the insect. Our results are in agreement with those of Zeidanand Czosnek (1994) who found that B. tabaci could ingest Agrobacterium. The failureto culture similar bacteria from surface sterilized whitefly eggs, suggests that these


Fig. 2. Descending midgut from whitefly fed WFA73 bacteria. Note bacterium (B)present in the lumen (L) that is being engulfed and bacteria (EB) present in mem-brane bound vesicles. M, mitochondrion. Bar = 1 mm.

Fig. 3. WFA73 bacterial cell (EB) actively being engulfed at apical surface of de-scending midgut epithelia in this whitefly fed the bacteria. Extracellular bacteria (B)are present in the lumen (L). M, mitochondrion. Bar = 1 mm.


bacteria were obtained from the leaf surface during probing prior to feeding. Thesebacteria are not transovarially transmitted, as the same bacteria were not isolatedfrom all samples and bacteria were not isolated from homogenized surface-sterilizedeggs. These culturable bacteria are therefore not obligate symbionts.

Rosell et al. (1995) demonstrated ultrastructurally that the adult B. tabaci styletfood canal is 0.65 mm in diameter. Using fluorescent beads, we have recently shownthat 0.2 mm beads are ingested and pass in honeydew, but 0.5 mm beads do not enterthe insect (Rosell et al., in prep). Therefore in order for bacteria to enter the food ca-nal, they must be less than 0.5 mm in diameter, or be pleomorphic with some membersof the population smaller than 0.5 mm. The bacteria which successfully entered thefood canal, strains WFA73 and WFN29, fulfilled these requirements. In contrast,Gram-positive bacteria including Bacillus spp. and Staphylococcus spp., which havegenerally larger diameters, entered the food canal poorly or not at all. As several Ba-cillus spp. and Staphylococcus spp. were isolated from surface-sterilized whiteflies,these organisms may have been located under the lingula at the posterior of the gut,where honeydew accumulates prior to excretion, or in the female reproductive tractsas seen in electron microscopy (Fig. 5).

Fig. 4. Descending midgut from whitefly fed WFA73 showing electron lucentspherical vesicles (Sv), electron dense lysosomal-like vesicles (Lv) and small dense re-sidual bodies (Rb). Bacteria are present in the lumen (L). Bar = 1 (m.


Isolate WFA73 (E. cloacae) appears to be mildly pathogenic to B. argentifoliiadults, and represents the first bacterial pathogen reported from whiteflies. In speci-mens fed this bacterium, midgut cells were massively invaded by bacteria, likely lead-ing to loss of part or all of gut function (Fig. 1). Phagocytosis of bacteria by insectmidgut cells has been observed during the pathogenesis of both American foulbrooddisease of honey bees (Davidson 1973), and milky disease of beetles (Kawanishi et al.1978, Splittstoesser et al. 1978). Enterobacter cloacae was described as a pathogen ofgrasshoppers under its original name, Coccobacillus acridiorum. Enterobacter (Aero-bacter) aerogenes is a pathogen of lepidoptera in association with Proteus mirabilis(Tanada and Kaya 1993, Wysoki and Raccah 1980) and occurs in the gut flora of theNew Zealand grass grub (Stucki et al. 1984).

Isolate WFA73 (E. cloacae), which was readily ingested by adults, originated fromwhiteflies with enhanced resistance to the insecticide Danitol® (Valent, USA) and is ca-pable of precipitating Danitol in vitro (E. Davidson, L. Williams and D. Alexander, unpubl.results). Therefore culturable bacteria associated with B. argentifolii may be important toinsecticide degradation in the phylloplane, and perhaps in the insect as well.

The relationship of E. cloacae to B. argentifolii is similar in several aspects to thebacterium designated BEV in leafhoppers. BEV can be cultured on bacteriological me-

Fig. 5. Bacteria-like organisms (BL) are found in the female reproductive tractwhich is convergent with the ovipositor canal (OV). C, cuticle; Mu, muscle; nu, nucleusof epidermal cell. Posterior of the whitefly is to the right. Bar = 5 mm.


dia, is mildly pathogenic to its normal host, Euscelidius variegatus Kirshbaum, andpenetrates gut cells in a manner ultrastructurally similar to E. cloacae in the whitefly.However BEV is both transmitted transovarially and acquired from the plant (Purcellet al. 1984, Purcell and Suslow 1987, Cheung and Purcell 1993).

Clark et al. (1992) examined the endosymbionts of B. argentifolii and B. tabaciusing 16S rDNA analysis, and found the secondary endosymbiont of Bemisia is re-lated to Enterobacteriaceae. Results presented here confirm that Enterobacteri-aceae are commonly present in B. argentifolii, and E. cloacae can enter the cells ofthe insect, suggesting that such bacteria could have been ancestors of whitefly endo-symbiotic bacteria, as suggested by Harada et al. (1996) for an endosymbiont of thepea aphid. Gut bacteria also present contaminating foreign bacterial DNA whichmay confuse genetic analysis of endosymbionts, as pointed out by Grenier et al.(1994) for aphids.

While it is clear that mycetome endosymbionts are critical to the development ofB. argentifolii (e.g. Costa et al. 1993b), the presence of other bacteria must be takeninto consideration in studies of the physiology of this insect. Although WFA73 (E. clo-acae) is only mildly pathogenic to B. argentifolii, its ability to penetrate whitefly gutcells suggests that this microorganism could be genetically modified to enhance its ef-fectiveness as a biological control agent. Transformation of a cotton phyllosphere bac-terium, B. megaterium, with Bacillus thuringiensis Berliner toxin genes for control oflepidoptera (Bora et al. 1994) is an example of such manipulation. Finally, geneticmodification of gut symbionts of Rhodnius prolixus to interfere with vectoring of Cha-gas disease has been reported (Beard et al. 1992, 1993). Similar modification of gutbacteria in other insects, such as whiteflies, to alter their ability to vector plant vi-ruses or other characteristics of these insects, is an intriguing possibility (Richards1993).

ACKNOWLEDGMENTS

Research at ASU was funded by a Cooperative Agreement with the USDA-ARSWestern Cotton Laboratory, Phoenix, AZ, and USDA CSREES 97-35316-5139. TheASU Life Sciences Electron Microscopy Laboratory and the University of Arizona Di-vision of Biotechnology Electron Microscopy facilities were used in this study, and weare grateful to William Sharp, Gina Zhang, Leah Kennaga and David Bentley for as-sistance. Identification of bacteria using fatty acid analysis was performed by Dr. JoelSiegel, Illinois Natural History Survey, Urbana, IL.

REFERENCES CITED

BEARD, C. B., P. W. MASON, S. AKSOY, R. B. TESH, AND F. F. RICHARDS. 1992. Trans-formation of an insect symbiont and expression of a foreign gene in the Chagas’disease vector, Rhodnius prolixus. Am. J. Trop. Med. Hyg. 46: 195-200.

BEARD, C. B., S. L. O’NEILL, R. B. TESH, F. F. RICHARDS, AND F. AKSOY, 1993. Modifi-cation of arthropod vector competence via symbiotic bacteria. Parasitol. Today9: 179-183.

BELLOWS, T. S. JR., T. M. PERRING, R. J. GILL, AND D. H. HEADRICK. 1994. Descriptionof a species of Bemisia (Homoptera: Aleyrodidae). Ann. Entomol. Soc. Am. 87:195-206.

BORA, R. S., M. G. MURTY, R. SHENBAGARATHAI, AND V. SEKAR. 1994. Introduction ofa lepidopteran-specific insecticidal crystal protein gene of Bacillus thuringien-


sis subsp. kurstaki by conjugal transfer into a Bacillus metagerium strain thatpersists in the cotton phyllosphere. Appl. Environ. Microbiol. 60: 214-222.

BYRNE, D. N., AND T. S. BELLOWS, JR. 1991. Whitefly biology. Annu. Rev. Entomol. 36:431-457.

CHEUNG, W. W. K., AND A. H. PURCELL, 1993. Ultrastructure of the digestive systemof the leafhopper Euscelidius variegatus Kirshbaum with and without congen-ital bacterial infections. Int. J. Insect Morphol. & Embryol. 22: 49-61.

CLARK, M. A., L. BAUMANN, M. A. MUNSON, P. BAUMANN, B. C. CAMPBELL, J. E. DUF-FUS, L. S. OSBORNE, AND N. A. MORAN. 1992. The Eubacterial endosymbiontsof whiteflies constitute a lineage distinct from the endosymbionts of aphids andmealybugs. Curr. Microbiol. 25: 119-123.

COCK, M. J. W. 1986. Bemisia tabaci, a literature survey on the cotton whitefly withan annotated bibliography. FAO/CAB, Ascot, UK.

COSTA, H. S., D. M. WESTCOT, D. E. ULLMAN, AND M. W. JOHNSON. 1993a. Ultrastruc-ture of the endosymbionts of the whitefly, Bemisia tabaci and Trialeurodes va-porariorum. Protoplasma 176: 106-115.

COSTA, H. S., D. E. ULLMAN, M. W. JOHNSON, AND B. E. TABASHNIK. 1993b. Antibioticoxytetracycline interferes with Bemisia tabaci oviposition, development andability to induce squash silverleaf. Ann. Entomol. Soc. Am. 86: 740-748.

COSTA, H. S., T. J. HENNEBERRY, AND N. C. TOSCA. 1997. Effects of antibacterial mater-ials on Bemisia argentifolii oviposition, growth, survival and sex ratio. J. Econ.Entomol. 90: 333-339.

DAVIDSON, E. W. 1973. Ultrastructure of American foulbrood disease pathogenesis inlarvae of the worker honey bee, Apis mellifera. J. Invertebr. Pathol. 21: 53-61.

DAVIDSON, E. W., B. J. SEGURA, T. STEELE, AND D. L. HENDRIX. 1994. Microorganismsinfluence the composition of honeydew produced by the silverleaf whitefly, Be-misia argentifolii. J. Insect Physiol. 40: 1069-1076.

DE BARRO, P. J. 1995. Bemisia tabaci biotype B: a review of its biology, distributionand control. CSIRO Div. Entomol. Tech. Paper No. 33, Canberra, Australia.

GRENIER, A.-M., C. NARDON, AND Y. RAHBE. 1994. Observations on the micro-organ-isms occurring in the gut of the pea aphid Acyrthosiphon pisum. Entomol. Exp.Appl. 70: 91-96.

HARADA, H., H. OYAIZU, AND H. ISHIKAWA. 1996. A consideration about the origin ofaphid intracellular symbiont in connection with gut bacterial flora. J. Gen.Appl. Microbiol. 42: 17-26.

KAWANISHI, C. Y, C. M. SPLITTSTOESSER, H. TASHIRO, AND K. H. STEINKRAUS. 1978.Infection of the European chafer, Amphimallon majalis by Bacillus popilliae:Ultrastructure. J. Invertebr. Pathol. 31: 91-102.

MYERS, P., AND A. A. YOUSTEN. 1978. Toxic activity of Bacillus sphaericus SSII-1 formosquito larvae. Inf. Immun. 19: 1047-1053.

PURCELL, A. H., T. STEINER, F. MEGRAUD, AND J. M. BOVE. 1986. In vitro isolation ofa transovarially transmitted bacterium from the leafhopper Euscelidus varie-gatus. J. Invertebr. Pathol. 48: 66-73.

PURCELL, A. H., K. G. SUSLOW, AND M. KLEIN. 1984. Transmission via plants of an in-sect pathogenic bacterium that does not multiply or move in plants. Microb.Ecol. 27: 19-26.

PURCELL, A. H., AND K. G. 1987. Pathogenicity and effects on transmission of a myco-plasma-like organism of a transovarially infective bacterium on the leafhopperEuscelidius variegatus. J. Invertebr. Pathol. 50: 285-290.

RICHARDS, F. F. 1993. An approach to reducing arthropod vector competence. ASMNews, 59: 509-514.

ROSELL, R. C., J. E. LICHTY, AND J. K. BROWN. 1995. Ultrastructure of the mouthpartsof adult sweetpotato whitefly, Bemisia tabaci. Int. J. Insect Morphol. & Em-bryol. 24: 297-306.

SPLITTSTOESSER, C. M., C. Y. KAWANISHI, AND H. TASHIRO. 1978. Infection of the Eu-ropean chafer, Amphimallon majalis by Bacillus popilliae: light and electronmicroscope observations. J. Invertebr. Pathol. 31: 84-90.

Epler et al.: Redescription of Cricotopus lebetis 171

SRIVASTAVA, P. N., AND J. W. ROUATT. 1963. Bacteria from the alimentary canal of thepea aphid, Acyrthosiphon pisum. J. Insect Physiol. 9: 435-438.

STUCKI, G., T. A. JACKSON, AND M. J. NOONAN. 1984. Isolation and characterizationof Serratia strains pathogenic for larvae of the New Zealand grass grub, Cost-elytra zealandica. N.Z. J. Science 27: 255-260.

TANADA, Y., AND H. KAYA. 1993. Insect Pathology. pp. 147-151. Academic Press, NY.WYSOKI, M., AND B. RACCAH. 1980. A synergistic effect of two pathogenic bacteria

from the Enterobacteriaceae on the geometrid Boarmia selenaria. J. Invertebr.Pathol. 35: 209-210.

ZEIDAN, M., AND H. CZOSNEK. 1994. Acquisition and transmission of Agrobacterium

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

by the whitefly Bemisia tabaci. Molec. Plant-Microbe Interactions 7: 792-798.

REDESCRIPTION OF CRICOTOPUS LEBETIS (DIPTERA: CHIRONOMIDAE), A POTENTIAL BIOCONTROL AGENT OF THE AQUATIC WEED HYDRILLA (HYDROCHARITACEAE)

J. H. EPLER1, J. P. CUDA2 AND T. D. CENTER3

1461 Tiger Hammock Road, Crawfordville, FL 32327

2Entomology and Nematology Department, University of FloridaPO Box 110620, Gainesville, FL 32611-0620

3Fort Lauderdale Research and Education Center, University of Florida3205 College Avenue, Fort Lauderdale, FL 33314-7799

ABSTRACT

The adult male and female of Cricotopus lebetis Sublette are redescribed and thepupa and larva described for the first time. Larvae of C. lebetis mine in the stems ofthe submersed aquatic weed hydrilla, Hydrilla verticillata (L.f. Royle), causing suffi-cient damage to the apical meristem to preclude further growth. The species is verysimilar to C. tricinctus (Meigen) but can be distinguished from that species in theadult male by the broader, more rounded inferior volsella; in the female by the lowernumber of sensilla chaetica on the mid and hind basitarsi; in the pupa by the fusiformthoracic horn; and in the larva by the simple S I and long setal tufts on abdominal seg-ments I-VII.

Key Words: Chironomidae, Cricotopus, taxonomy, Hydrilla verticillata, aquatic weed,biocontrol

RESUMEN

Se redescriben el adulto macho y hembra de Cricotopus lebetis Sublette y se des-criben por primera vez la larva y pupa de esta especie. Las larvas de C. lebetis minanlos tallos de la hierba acuática sumergida Hydrilla verticillata (L.f. Royle), causandosuficiente daño al meristemo apical como para impedir su crecimiento. C. lebetis esmuy similar a C. tricinctus (Meigen), pero se distingue de esta especie en que el machoadulto posee una volsella inferior más redondeada y ancha, mientras que la hembra

Epler et al.: Redescription of

Cricotopus lebetis 171

S

RIVASTAVA

, P. N.,

AND

J. W. R

OUATT

. 1963. Bacteria from the alimentary canal of thepea aphid,

Acyrthosiphon pisum.

J. Insect Physiol. 9: 435-438.S

TUCKI

, G., T. A. J

ACKSON

,

AND

M. J. N

OONAN

. 1984. Isolation and characterizationof

Serratia

strains pathogenic for larvae of the New Zealand grass grub,

Cost-elytra

zealandica

. N.Z. J. Science 27: 255-260.T

ANADA

, Y.,

AND

H. K

AYA

. 1993. Insect Pathology. pp. 147-151. Academic Press, NY.W

YSOKI

, M.,

AND

B. R

ACCAH

. 1980. A synergistic effect of two pathogenic bacteriafrom the Enterobacteriaceae on the geometrid

Boarmia selenaria

. J. Invertebr.Pathol. 35: 209-210.

Z

EIDAN

, M.,

AND

H. C

ZOSNEK

. 1994. Acquisition and transmission of

Agrobacterium

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

by the whitefly

Bemisia tabaci

. Molec. Plant-Microbe Interactions 7: 792-798.

REDESCRIPTION OF

CRICOTOPUS LEBETI

S (DIPTERA: CHIRONOMIDAE), A POTENTIAL BIOCONTROL AGENT OF THE AQUATIC WEED HYDRILLA (HYDROCHARITACEAE)

J. H. E

PLER

1

, J. P. C

UDA

2

AND

T. D. C

ENTER

3

1

461 Tiger Hammock Road, Crawfordville, FL 32327

2

Entomology and Nematology Department, University of FloridaPO Box 110620, Gainesville, FL 32611-0620

3

Fort Lauderdale Research and Education Center, University of Florida3205 College Avenue, Fort Lauderdale, FL 33314-7799

A

BSTRACT

The adult male and female of

Cricotopus lebetis

Sublette are redescribed and thepupa and larva described for the first time. Larvae of

C. lebetis

mine in the stems ofthe submersed aquatic weed hydrilla,

Hydrilla verticillata

(L.f. Royle), causing suffi-cient damage to the apical meristem to preclude further growth. The species is verysimilar to

C. tricinctus

(Meigen) but can be distinguished from that species in theadult male by the broader, more rounded inferior volsella; in the female by the lowernumber of sensilla chaetica on the mid and hind basitarsi; in the pupa by the fusiformthoracic horn; and in the larva by the simple S I and long setal tufts on abdominal seg-ments I-VII.

Key Words: Chironomidae,

Cricotopus

, taxonomy,


, aquatic weed,biocontrol

R

ESUMEN

Se redescriben el adulto macho y hembra de

Cricotopus lebetis

Sublette y se des-criben por primera vez la larva y pupa de esta especie. Las larvas de

C. lebetis

minanlos tallos de la hierba acuática sumergida


(L.f. Royle), causandosuficiente daño al meristemo apical como para impedir su crecimiento.

C. lebetis

esmuy similar a

C. tricinctus

(Meigen), pero se distingue de esta especie en que el machoadulto posee una volsella inferior más redondeada y ancha, mientras que la hembra

172


83(2) June, 2000

posee menos sénsulos tipo chaetica en los basitarsi medio e inferior. Asmísmo, la pupade

C. lebetis

presenta un cuerno torácico fusiforme y la larva tiene un S I simple y lar-

gos pinceles de cerdas en los segmentos abdominales I-VII.

The chironomid genus

Cricotopus

van der Wulp is common, widespread and speci-ose. Hirvenoja (1973) revised the Palaearctic species but in the Nearctic the taxonomyof the genus remains in less than satisfactory condition. Many undescribed species ex-ist and the conspecificity of some Nearctic taxa with species originally described fromthe Palaearctic is uncertain.

One such species is

Cricotopus lebetis

Sublette, a member of the

sylvestris

group ofthe subgenus

C. (Isocladius)

. Interest in the taxonomy of

C. lebetis

has been recentlystimulated by the discovery of the larvae of this species feeding within the stems ofhydrilla,


(L.f. Royle) (Hydrocharitaceae), a well known pestaquatic plant that was introduced into Florida in the 1950’s (Schmitz et al. 1991). Lar-vae of

C. lebetis

mine in the stems of hydrilla, causing sufficient damage to the plant’sapical meristem to preclude further growth of the plant. This natural growth controlmay prevent hydrilla from reaching the water’s surface, eliminating the dense surfacemats which reduce biodiversity and interfere with navigation.

In this paper the adult male and female of

C. lebetis

are redescribed and the pupaand larva are described for the first time. Information on the midge’s life history andpotential use as a biocontrol agent for hydrilla is discussed in Cuda et al. (1999).

M

ATERIALS

AND

M

ETHODS

Morphological terminology and abbreviations follow Sæther (1980), Oliver & Dil-lon (1989), Epler (1988) and Sublette, et al. (1998). Measurements are in

m

m, unlessotherwise stated, and consist of the range followed by the mean if three or more spec-imens were measured.

For the descriptions below, the majority of the adult male material and all of theadult female, pupal and larval material was from the F3 generation of laboratoryreared midges originally collected from the Plantation Inn Canal, Crystal River inCitrus Co., Florida, on 23 September 1997 (see Cuda et al. 1999); data from twoparatype males are included in the adult male description.

S

YSTEMATICS

Sublette (1964) described

Cricotopus lebetis

from adult male and female speci-mens collected in Louisiana in 1957-1959. He noted that this new species would keyto

C. tricinctus

(Meigen)

in Johannsen and Townes (1952) and that it was difficult toseparate

C. lebetis

from Palaearctic material of

C. tricinctus

(Meigen) on the basis ofcolor pattern. He also stated (1964: 118) that the “strong, almost right angled basallobe [= inferior volsella of Sæther (1980) and Oliver & Dillon (1989)] on the basistyle[= gonocoxite] as well as the shape of the dististyle” [= gonostylus] seemed to be dis-tinctive for

C. lebetis

.Beck & Beck (1966: 131) listed

Cricotopus lebetus

[sic] as a “recently substitutedAmerican name” for

C. tricinctus

, but did not give any references or reason for thisplacement.



Hirvenoja (1973: 304), in the section Ökologie und Verbreitung (“Ecology and Dis-tribution”) under

C. tricinctus

, mentioned Beck and Beck’s (1966) listing of

C. lebetis

as a synonym of

C. tricinctus

with some doubt as indicated by a “?” before his listing;he did not list

C. lebetis

as a synonym of

C. tricinctus

.Only Boesel (1983) formally listed

C. lebetis

as a new synonym of

C. tricinctus

, aconcept followed by Oliver et al. (1990).

Placement of

C. lebetis

in the papers above was based only on characters of theadult stage. When characters from all life stages are considered, in particular thepupa and larva, it is readily apparent that

C. lebetis

is a taxon distinct from

C. tricinc-tus

.

Cricotopus (Isocladius) lebetis

Sublette

Cricotopus lebetis

Sublette, 1964: 118 (description of adult male and fe-male).

Cricotopus tricinctus

(Meigen, 1818) partim: Boesel 1983:81(synonymy; inkey); Oliver et al. 1990: 24 (synonymy); and other North American au-thors.

Male imago (n = 10, unless otherwise noted)

Color: In life, pale green with blackish-brown markings; these colors fade to palebrown/stramineous with dark brown to brownish markings in alcohol preserved ma-terial. In alcohol preserved material, brown to dark brown on antennae, head, tho-racic vittae (vittae sometimes joined posteriorly by diffuse brown area), scutellum,postnotum, median anepisternum, almost all to ventral

⅔

of preepisternum, ventralhalf of anterior anepisternum II and approximate ventral half of epimeron. Wingsclear with light brown veins; halteres pale. Legs (Fig. 1) with fore and hind coxaelight, mid coxa brown; all trochanters light; fore femur light brown basally, muchdarker in apical

⅓

to

½

; mid and hind femora basally light with brown apical

⅓

to

¼

;tibiae with brown basal and apical bands, fore tibia slightly darker in middle thanmid and hind tibiae; fore tarsi brown, mid and hind tarsi light brown to stramineous.Abdomen (Fig. 2) with T I and IV stramineous; T II with posterior

½

brown; T III withposterior

4

/

5

brown; T V mostly brown, with paler anterior and posterior margins; T VIwith brown band across middle; T VII mostly stramineous, often with brown mark-ings posterolaterally, sometimes almost completely dark; T VIII mostly brown, withnarrow posterior stramineous band; T IX mostly brown; gonocoxites and gonostylistramineous.

Length. Body (excluding head): 2.35-2.88, 2.55 mm (n = 5); thorax 0.70-0.85, 0.74mm (n = 5); abdomen 1.65-2.05, 1.86 mm (n = 8).

Head. Temporal setae 6-8, 7; clypeal setae 4-8, 6; cibarial sensillae 2-9, 6. Lengthsof palpomeres 2-5 (n = 8): 30-45, 37; 50-63, 56; 53-73, 63; 88-107, 97. AR 0.77-0.93, 0.83.

Thorax. Setae: lateral antepronotal (n = 8) 0-2, 1; acrostichal (n = 6) 10-13, 12; dor-socentral (n = 7) 8-14, 10; prealar (n = 9) 3-6, 4; scutellar (n = 9) 6-7, 6.

Wing. Length (n = 7) 1.08-1.30, 1.16 mm; width (n = 7) 310-380, 342. VR (n = 7)1.14-1.23, 1.18. Costal extension (n = 6) 18-50, 31. Setae: brachiolum 1; squama (n =9) 4-8, 6; R

1

(n = 8) 2-4, 3.Legs. Lengths of tibial spurs: fore 30-40, 34; mid 12-17, 14 (n = 9); 15-18, 17; hind

14-20, 18; 35-44, 38. Sensilla chaetica: mid 8-15, 12 (n = 9); hind 18-29, 23. Hind tibialcomb with 8-10, 9 setae (n = 8), setal length 27-43, 35. Pulvilli developed, about

½

length of claw. Lengths and proportions of legs:

174


83(2) June, 2000

Abdomen

(n = 9). T III with 2-4, 3 median setae and 2-4, 3 lateral setae; T IV with2-5, 3 median setae and 3-5, 4 lateral setae.

Hypopygium (Fig. 3). T IX with 5-10, 7 setae; laterosternite IX with 3-5, 4 setae.Transverse sternapodeme width 83-155, 100 (n = 9); phallapodeme length 63-78, 70(n = 9). Virga absent. Superior volsella well developed; inferior volsella (Figs. 4-6) lin-guiform to triangular. Gonocoxite length 163-205, 179; gonostylus length 80-98, 86,with well developed crista dorsalis (Fig. 7); GC/GS 1.99-2.13, 2.07. Megaseta length14-16, 15 (n = 5).

Female imago (n = 5, unless otherwise noted)

Color. Mostly as in male, except thoracic vittae sometimes joined by diffuse browncoloration, so that dorsum of thorax appears mostly brown (in fluid preserved speci-mens). Legs light brown, with weaker vittate pattern than in male (pale areas not aspale as in male). Abdominal tergites similar to male, except T IX and gonocoxite IXbrown.

Length. Body (excluding head) 2.23-2.65, 2.51 (n = 3); thorax 0.68-0.77, 0.74; abdo-men 1.55-1.90, 1.78 (n = 3).

Head. Temporal setae 2-7, 5; clypeal setae 7-9, 8; cibarial sensillae 3-10, 6. Lengthsof palpomeres 2-5 (n = 4): 30-40, 34; 43-53, 49; 48-55, 52; 88-103, 95. Antenna with 5flagellomeres; AR 0.40-0.55, 0.47.

Thorax

.

Setae: lateral antepronotal 1-2, 2; acrostichal 9-12, 11; humeral 2-3, 2;dorsocentral 7-10, 9; prealar 3-4, 3; scutellar 5-8, 7.

Wing. Length (n = 4) 1.08-1.20, 1.00 mm; width (n = 4) 380-420, 395. VR (n = 4)1.18-1.22, 1.20. Costal extension (n = 3) 40-60, 50. Setae: brachiolum 1; squama 4-11,7; R (n = 4) 4-5, 4; R

1

(n = 4) 0-3, 2; R

4+5

(n = 4) 1-3, 2.Legs. Lengths of tibial spurs: fore 21-25, 23; mid 10-13, 12; 12-15, 14; hind 11-15,

13; 33-38, 36. Sensilla chaetica: mid 18-21, 20; hind 22-34, 27. Hind tibial comb with9-12, 10 setae, setal length 35-37, 36 (n = 3). Pulvilli developed, about

½

length of claw.Lengths and proportions of legs (n = 4):

P1 P2 P3fe 440-600, 510 475-605, 535 430-640, 535ti 560-735, 627 480-650, 551 510-730, 604ta1 260-375, 304 190-275, 226 260-375, 305 (n = 9)ta2 140-210, 164 100-160, 124 125-185, 151 (n = 9)ta3 110-150, 126 85-115, 96 120-160, 134 (n = 9)ta4 80-95, 83 55-70, 63 65-85, 73 (n = 9)ta5 65-80, 73 60-70, 65 60-80, 68 (n = 9)LR 0.46-0.51, 0.48 0.39-0.42, 0.41 0.48-0.55, 0.50BV 3.09-3.42, 3.24 3.53-3.96, 3.78 3.15-3.49, 3.34 (n = 9)SV 3.50-3.94, 3.75 4.47-5.07, 4.82 3.50-3.89, 3.74

P1 P2 P3fe 370-430, 406 420-490, 458 420-490, 461ti 465-520, 500 430-480, 468 490-530, 520ta1 210-245, 226 175-200, 190 245-280, 268ta2 95-120, 106 80-90, 86 100-125, 113ta3 60-90, 76 60-75, 65 90-110, 101ta4 45-60, 50 40-50, 44 45-50, 49ta5 50-60, 55 55 50-60, 58LR 0.44-0.47, 0.45 0.40-0.42, 0.41 0.50-0.53, 0.51BV 3.82-4.18, 3.95 4.33-4.67, 4.46 3.75-4.02, 3.91SV 3.88-4.11, 4.01 4.85-4.89, 4.87 3.57-3.71, 3.67



Abdomen

(n = 3). T III with 2-3, 2 median setae and 2 lateral setae; T IV with 2 me-dian setae and 2-3, 2 lateral setae.

Genitalia

(Figs.

8-10). Notum 105-143, 124 long (measured to bifurcation); seminalcapsule diameter 50-60, 57 (n = 3), cercus length 85-103, 97 (n = 4) (measured fromventral aspect). Spermathecal ducts with at least one loop. Coxosternapodeme as inFig. 10. T IX with 3-5, 4 setae; gonocoxite IX with 8-12, 11 setae.

Pupa (n = 10, unless otherwise noted)

Color. Exuviae pale yellow with narrow, light brown bands at posterior of T II (overhooklet row) and T III; T IV entirely pale; T V with posterior 2/3 light brown; T VI withmedian light brown area; lateral margins of T VI-VIII and anal lobes light brown.

Length. Total 2.50-3.18, 2.79 mm (n = 8); cephalothorax 0.75-0.95, 0.81 mm (n = 7);abdomen 1,70-2.30, 1.99 mm (n = 9).

Cephalothorax. Frontal setae 60-88, 73 (n = 6) long, 2 wide; dorsal median an-tepronotal seta 58-73, 64 (n = 6) long; ventral median antepronotal seta 83-155, 96 (n= 8) long; lateral antepronotal seta 25-40, 33 (n = 8) long. Median suture area smooth.Thoracic horn (Fig. 11) fusiform with sparsely scattered minute spinules; 50-83, 67long; 15-20, 17 (n = 9) maximum width. Precorneal setae lengths: PC

1

88-105, 101 (n= 9); PC

2

65-93, 77 (n = 8); PC

3

63-100, 82 (n = 7). Dorsocentral setae lengths: DC

1

33-48, 39; DC

2

25-60, 36 (n = 9); DC

3

30-40, 36 (n = 8); DC

4

35-48, 39 (n = 9); DC

1

stouterthan DC

2

. Wing sheath without bacatiform papillae or nasiform tubercles.Abdomen (Fig. 12). T II with 49-68, 56 hooklets arranged in double row. Pedes spu-

rii B weakly developed on T II; pedes spurii A present on S IV-VI. Tergite I with oneanterolateral seta, T II-VIII with 3 lateral setae (2 dorsal and one ventral). Dorsalshagreen on T I sparse, scattered minute spinules in weak, longitudinal lateral bands;T II with scattered weak spinules over most of surface; T III with fine spinules overmost of surface; T IV-VI with larger spinules over most of surface, with coarserspinules at center of tergites; T VII-VIII with anterior band of fine spinules; anal discwith small anteromedian area of fine spinules. Conjunctiva III-IV, IV-V and V-VI withspinules. Ventral shagreen consists of small posterolateral groups of minute spinuleson S I; S II-V with weak longitudinal bands of minute spinules; S VI-VII with smallanterolateral groups of minute spinules. Anal lobes with 3 macrosetae; anal lobelength 200-218, 208 (n = 7). Lengths of anal lobe macrosetae (n = 9): seta 1 (anterior-most seta) 78-88, 82; seta 2 (middle seta) 73-95, 85; seta 3 80-102, 91. The anal loberatio (ALR) varies depending on which macroseta is measured and compared to theanal lobe length, thus ALR1 (using anteriormost seta) 0.39-0.43, 0.40; ALR2 0.38-0.45, 0.42; ALR3 0.40-0.47, 0.43 (all ALR n = 7).

Fourth instar larva (n = 11, unless otherwise noted)

Color. In life, the body is green with blue bands on the second and third thoracicsegment; the blue color is bleached on alcohol preserved specimens but the thorax re-mains darker than the remainder of the body in such material. Head capsule pale yel-low-brown, premandibles light brown, mentum and apical

⅓

to

½

of mandible darkbrown to black. Claws of parapods translucent to pale brown.

Head. Postmentum length 170-205, 186 (n = 9). Labrum (Fig. 13) with simple S I.Total antennal length 61-73, 64 (Fig. 14). Length of antennal segments 1-5: 31-43, 37;13-18, 16; 9-13, 11; 3-4, 4; 3-4, 4; 2. Ring organ 8-10, 9 (n = 5) from base of basal seg-ment; sensory pits slightly above to around same level as ring organ. Lauterborn or-gans extend to apex of antennal segment 3. AR 1.11-1.60, 1.34. Premandible (Fig. 15)apically bifid; length 69-80, 74. Mandible (Fig. 16) length 120 = 137, 127; with 3 inner

176


83(2) June, 2000

Figs. 1-10. Cricotopus lebetis adult structures. 1. Male fore, mid and hind legs; 2.Male abdomen; 3. Hypopygium; 4-5. Inferior volsella variation in Florida material; 6.Inferior volsella, Louisiana specimen; 7. Variation of gonostylus due to angle of obser-vation; 8. Female genitalia, ventral; 9. Female genitalia, lateral; 10. Female coxoster-napodeme.



teeth; apical tooth length 13-16, 14 (n = 5); width of inner teeth 23-27, 25 (n = 5). Outermargin of mandible mostly smooth; inner margin of mandible without spines; man-dibular margin at base of seta subdentalis without minute teeth. Seta internapresent, usually with 6 branches. Mentum (Fig. 17) with 13 teeth; second lateral toothsmall and fused to first. Maxilla as in Fig. 18.

Body. Small claws of anterior parapods (Fig. 19) with apical tooth much largerthan inner teeth. Abdominal segments I-VII with long setal tufts (Fig. 20); setal tuftswith about 25-50 setae, longest setae about 385 long; tuft on VII with smaller andfewer (about 11-20) setae. Anal tubules elongate-ovoid.

D

ISCUSSION

The color pattern of adults can be variable in many species of

Cricotopus

;

C. lebetis

is no exception. In males, tergite VII is apparently most susceptible to color variation;it may be almost totally unmarked with brown or, as in the majority of the Florida ma-terial examined, marked with brown in the posterolateral corners. Sublette (pers.comm.) has seen material of

C. lebetis

from Baton Rouge, Louisiana, in which T VII isalmost completely infuscate.

In general, the type series is darker than the Florida material examined. Sublette(1964) stated that the female pronotum (= antepronotum) was infuscate; in the Flor-ida material examined the antepronotum is stramineous, similar to other unmarkedbody areas.

Adults of

C. lebetis

are most likely to be confused with

C. tricinctus

, as originallynoted by Sublette (1964). In males, the inferior volsella of

C. tricinctus

is generallynarrower and more triangular than that of

C. lebetis

. There is variation in the shapeof this lobe; figure 6 is from a Louisiana paratype and is similar to, but still shorterand broader than, the volsella of some European

C. tricinctus

(see Hirvenoja 1973:Fig. 189(3) and 189(5)).

Females of

C. lebetis

are also similar to

C. tricinctus

; lower counts of sensilla cha-etica on the mid and hind basitarsi (means of 22 and 27) will separate the Florida

C.lebetis

females examined from those of

C. tricinctus

(means of 48 for both legs), follow-ing the data from Hirvenoja (1973). It must be noted that the descriptions above (forall life stages of

C. lebetis

, with the exception of the two males from Louisiana) are forindividuals from one population in Florida. These individuals may be smaller thanpopulations of this species from other areas; setal counts and other measurementsmay also show a wider range once more material is examined.

Pupae of the two species are also similar, but the thoracic horn of

C. lebetis

is fusi-form compared to the elongate digitiform horn of

C. tricinctus

(see Hirvenoja 1973:Fig. 190(2)).

Larvae of

C. lebetis

are similar to other members of the

C. sylvestris

group, but dif-fer in bearing setal tufts on abdominal segments I-VII; in

C. tricinctus

larvae, only ab-dominal segments I-VI possess setal tufts. All larvae of

C. lebetis

examined had simpleS I; the S I of related species are bifid (but note that the bifurcation of some of theseother species may be unequal, with one branch much larger than the other; see Hir-venoja 1973: Fig. 191(3)).

An unresolved problem is that of the origin of North American

Cricotopus lebetis.Is this a species that was introduced with hydrilla, or is it native to North Americaand has seized upon introduced hydrilla as a suitable host plant? Is C. lebetis a facul-tative miner of hydrilla or does it attack other plants? Hirvenoja (1973) noted that Eu-ropean C. tricinctus larvae mine in the leaves of Potamogeton, and that other aquaticplant species may also be attacked.


Figs. 11-20. Cricotopus lebetis pupal structures; 13-20, larval structures. 11. Vari-ation in thoracic horn; 12. Abdomen, dorsal; 13. Labrum; 14. Antenna; 15. Premandi-ble; 16. Mandible; 17. Mentum; 18. Maxilla; 19. Small claws of anterior parapod; 20.Lateral setae of abdominal segment II.

Epler et al.: Redescription of Cricotopus lebetis 179

As seen above, C. tricinctus and C. lebetis have been confused in North America;Sublette (pers. comm.) has seen true C. tricinctus specimens from the north centralU.S. and compared them with Palaearctic material from the British Museum (NaturalHistory) and from Hirvenoja’s collection; thus both species occur in the Nearctic. TheCrystal River specimens constitute the first record of C. lebetis from Florida. The pos-sibility exists that C. lebetis has been misidentified as C. tricinctus or other species inother countries where hydrilla occurs (i.e., Japan). Cricotopus nitens (Kieffer, 1921)and C. taiwanus Tokunaga, 1940, both described from Taiwan, have a similar colorpattern; if not distinct species, either may be a senior synonym. The best way to solvethis riddle would be to rear all life stages of hydrilla-associated Cricotopus throughoutthe range of the plant.

ACKNOWLEDGMENTS

We are extremely grateful to Dr. J. E. Sublette, Tucson, AZ, for his assistance inidentifying C. lebetis, reviewing an early draft of this paper and the loan of specimens.We also thank B. A. Caldwell for providing a pre-submission review of this paper andDr. C. L. de la Rosa for providing the Spanish abstract. The senior author wishes to es-pecially thank Dr. Barry Merrill and Judy Merrill (Merrill Consultants, Dallas, TX)for providing laboratory and computer equipment. This is Florida Agricultural Exper-iment Station Journal Series No. R-07159.

REFERENCES CITED

BECK, W. M. JR., AND E. C. BECK. 1966. The Chironomidae of Florida: A problem ininternational taxonomy. Gewässer und Abwässer 41/42: 129-135.

BOESEL, M. W. 1983. A review of the genus Cricotopus in Ohio, with a key to adults ofspecies of the northeastern United States (Diptera, Chironomidae). Ohio J. Sci.83: 74-90.

CUDA, J. P., B. R. COON, J. L. GILMORE, AND T. D. CENTER. 1999. Preliminary reporton the biology of a hydrilla stem tip mining midge (Diptera: Chironomidae).Aquatics 21(4): 15-18.

EPLER, J. H. 1988. Biosystematics of the genus Dicrotendipes Kieffer, 1913 (Diptera:Chironomidae: Chironominae) of the world. Mem. American Entomol. Soc. 36:1-214.

HIRVENOJA, M. 1973. Revision der Gattung Cricotopus van der Wulp und ihrer Ver-wandten (Diptera, Chironomidae). Ann. Zool. Fennici 10: 1-363.

JOHANNSEN, O. A., AND H. K. TOWNES. 1952. Tendipedidae (Chironomidae). Guide tothe Insects of Connecticut. Part VI. The Diptera or true flies of Connecticut.Fifth fasicle: Midges and Gnats. St. Geol. Nat. Hist. Surv. Bull. 80: 3-26.

KIEFFER, J. J. 1921. Chironomides des Philippines et de Formose. Philippine J. Sci. 18:557-593.

OLIVER, D. R., AND M. E. DILLON. 1989. 2. The adult males of Chironomidae (Diptera)of the Holarctic region—Key to subfamilies. Entomol. Scandinavica Suppl. 34:11-15.

OLIVER, D. R., M. E. DILLON, AND P. S. CRANSTON. 1990. A catalog of Nearctic Chi-ronomidae. Research Branch Agriculture Canada Pub. 1857/B. 89 pp.

SÆTHER, O. A. 1980. Glossary of chironomid morphology terminology (Diptera: Chi-ronomidae). Entomol. Scandinavica Suppl. 14: 1-51.

SCHMITZ, D. C., B. V. NELSON, L. E. NALL, AND J. D. SCHARDT. 1991. Exotic plants inFlorida: a historical perspective and review of the present aquatic plant regu-lation program. pp. 303-326 in Center, T. D., R. F. Doren, R. L. Hofstetter, R. L.Meyers, and L. D. Whiteaker (eds.), Proceedings, Symposium on Exotic Plant


Pests, 2-4 November 1988, Miami, Florida. U.S. Dept. of the Interior, NationalPark Service, Washington, D.C.

SUBLETTE, J. E. 1964. Chironomidae (Diptera) of Louisiana. I. Systematics and imma-ture stages of some lentic chironomids of west-central Louisiana. Tulane Stud.Zool. 11: 109-150.

SUBLETTE, J. E., L. E. STEVENS, AND J. P. SHANNON. 1998. Chironomidae (Diptera) ofthe Colorado River, Grand Canyon, Arizona, USA, I: Systematics and ecology.Great Basin Naturalist 58: 97-146.

TOKUNAGA, M. 1940. Chironomoidea from Japan (Diptera), XII. New or little-known

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

Ceratopogonidae and Chironomidae. Philippine J. Sci. 72: 255-311 + 4 plates.

BUZZING BEES (HYMENOPTERA: APIDAE, HALICTIDAE)ON SOLANUM (SOLANACEAE): FLORAL CHOICE AND

HANDLING TIME TRACK POLLEN AVAILABILITY

TODD E. SHELLY1, ETHEL VILLALOBOS2 AND STUDENTS OF THE FALL 1997 OTS-USAP2,3

1Present address: USDA-APHIS, P.O. Box 1040, Waimanalo, HI 96795and Hawaiian Evolutionary Biology Program, University of Hawaii

Honolulu, HI 96822

2Organization for Tropical Studies-Undergraduate Semester Abroad ProgramDuke University, 410 Swift Avenue, Durham, NC 27705

3Students (in alphabetical order): Lisa Bell, Aisha Burden, Mark Fox,Ilmi Granoff, Nihara Gunawardene, Melisa Holman, Allison Hornor,

Jane MacLeod, Julia Michalek, Casuarina McKinney-Richards,Adam Ruff, Aaron Smith, Darcy Thomas, and Olivia Watson

ABSTRACT

Flower selection and pollen-collecting effort were monitored for 3 species of beesthat sonicate flowers of Solanum wendlandii Hook. for pollen in southern Costa Rica.Between 0700-0900 hours, Bombus pullatus (Fkln.), Euglossa erythrochlora Moure,and Pseudaugochloropsis graminea (Fabricius) foraged more frequently at new flow-ers (that had opened the day of observation) than old ones (that had opened at least1 day before observation). Between 0900-1100 hours, however, this preference was nolonger evident, and all 3 species visited new and old flowers with similar frequency. E.erythrochlora and P. graminea spent more time harvesting pollen during 1) initial(first or second) visits to new flowers than initial visits to old flowers and 2) initial vis-its to new flowers than final (seventh or later) visits to new flowers. Similar, althoughnot statistically significant, trends were evident for B. pullatus as well. An experi-ment using pollinator exclusion bags revealed that the reduced foraging effort at in-dividual flowers was resource-dependent and was not simply a time-dependentphenomenon.

Key Words: Apidae, buzz pollination, Costa Rica, foraging behavior, Halictidae,Solanum

180


83(2) June, 2000

Pests, 2-4 November 1988, Miami, Florida. U.S. Dept. of the Interior, NationalPark Service, Washington, D.C.

S

UBLETTE

, J. E. 1964. Chironomidae (Diptera) of Louisiana. I. Systematics and imma-ture stages of some lentic chironomids of west-central Louisiana. Tulane Stud.Zool. 11: 109-150.

S

UBLETTE

, J. E., L. E. S

TEVENS

,

AND

J. P. S

HANNON

. 1998. Chironomidae (Diptera) ofthe Colorado River, Grand Canyon, Arizona, USA, I: Systematics and ecology.Great Basin Naturalist 58: 97-146.

T

OKUNAGA

, M. 1940. Chironomoidea from Japan (Diptera), XII. New or little-known

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

Ceratopogonidae and Chironomidae. Philippine J. Sci. 72: 255-311 + 4 plates.

BUZZING BEES (HYMENOPTERA: APIDAE, HALICTIDAE)ON

SOLANUM

(SOLANACEAE): FLORAL CHOICE AND HANDLING TIME TRACK POLLEN AVAILABILITY

T

ODD

E. S

HELLY

1

, E

THEL

V

ILLALOBOS

2

AND

S

TUDENTS

OF

THE

F

ALL

1997 OTS-USAP

2,3

1

Present address: USDA-APHIS, P.O. Box 1040, Waimanalo, HI 96795and Hawaiian Evolutionary Biology Program, University of Hawaii

Honolulu, HI 96822

2

Organization for Tropical Studies-Undergraduate Semester Abroad ProgramDuke University, 410 Swift Avenue, Durham, NC 27705

3

Students (in alphabetical order): Lisa Bell, Aisha Burden, Mark Fox,Ilmi Granoff, Nihara Gunawardene, Melisa Holman, Allison Hornor,

Jane MacLeod, Julia Michalek, Casuarina McKinney-Richards,Adam Ruff, Aaron Smith, Darcy Thomas, and Olivia Watson

A

BSTRACT

Flower selection and pollen-collecting effort were monitored for 3 species of beesthat sonicate flowers of

Solanum wendlandii

Hook. for pollen in southern Costa Rica.Between 0700-0900 hours,

Bombus pullatus

(Fkln.),

Euglossa erythrochlora

Moure,and

Pseudaugochloropsis graminea

(Fabricius) foraged more frequently at new flow-ers (that had opened the day of observation) than old ones (that had opened at least1 day before observation). Between 0900-1100 hours, however, this preference was nolonger evident, and all 3 species visited new and old flowers with similar frequency.

E

.

erythrochlora

and

P

.

graminea

spent more time harvesting pollen during 1) initial(first or second) visits to new flowers than initial visits to old flowers and 2) initial vis-its to new flowers than final (seventh or later) visits to new flowers. Similar, althoughnot statistically significant, trends were evident for

B

.

pullatus

as well. An experi-ment using pollinator exclusion bags revealed that the reduced foraging effort at in-dividual flowers was resource-dependent and was not simply a time-dependentphenomenon.

Key Words: Apidae, buzz pollination, Costa Rica, foraging behavior, Halictidae,

Solanum

Shelly et al.: Buzzing Bees on

Solanum 181

R

ESUMEN

Se monitoreó la selección de flores y el esfuerzo en recolectar polen de tres especiesde abejas que frecuentan flores de

Solanum

wendlandii

Hook. en al Sur de Costa Rica.Entre las 0700 y 0900 h,

Bombus

pullatus

(Fkln.),

Euglossa

erythrochlora

Moure y

Pseudaugochloropsis

graminea

(Fabricius) forrajearon con mayor frecuencia floresnuevas (flores que abrieron el mismo día de hacerse la observación) que flores viejas(flores que abrieron por lo menos un día antes de hacerse la observación). Sin em-bargo, esta preferencia no se observó entre las 0900 y 1100 h, ya que las tres especiesvisitaron flores nuevas y viejas con la misma frecuencia.

E.

erythrochlora

y

P.

grami-nea

emplearon más tiempo cosechando polen en visitas iniciales (primera o segundavisitas) a flores nuevas que en visitas iniciales a flores viejas. Además, emplearon mástiempo en visitas iniciales a flores nuevas que en visitas finales (séptima visita en ade-lante) a flores nuevas. Se observó una tendencia similar (aunque no estadísticamentesignificativa) en

B. pullatus

. Un experimento empleando bolsas para excluír poliniza-dores demostró que la reducción en el esfuerzo por forrajear flores individuales estuvo

determinada por la disponibilidad de alimento y no por el horario.

The manner in which nectar rewards influence flower selection and floral handlingtimes has been well studied for a number of bee species (e.g.,

Cresswell 1990, Giurfaand Nunez 1992, Dukas and Real 1993). Early studies of nectar-collecting by bees(e.g., Waddington and Heinrich 1979, Pyke 1979) lent strong empirical support to thedevelopment of optimal foraging theory (Krebs and McCleery 1984). In contrast,fewer studies have investigated the foraging choices of bees harvesting pollen, andthese have yielded mixed results regarding the ability of bees to assess pollen rewardsfrom individual flowers. For example, Haynes and Mesler (1984) observed bumblebeesforaging on inflorescences of a lupine species and found bees did not discriminate be-tween old, pollen-poor flowers and younger flowers that contained greater pollen re-wards. Based on patterns of turning frequency and directionality, Hodges and Miller(1981) similarly concluded that bumblebees were not adjusting their foraging move-ments in response to pollen availability.

On the other hand, several studies have demonstrated that bees do modify theirforaging behavior in response to either anticipated or actual pollen rewards for in-dividual floral visits. In the first demonstration of “distant assessment”, Pellmyr(1988) showed that bumblebees assessed pollen rewards prior to alighting, based onage-dependent changes in floral shape, and rejected old flowers in favor of younger,pollen-rich flowers. He also showed that bumblebees adjusted their handling timewith pollen availability and spent more time on younger flowers than older ones.Likewise, Gori (1989) experimentally removed pollen and found that bumblebees re-sponded by visiting fewer flowers per inflorescence. Buchmann and Cane (1989) andHarder (1990) also reported a positive relation between pollen availability and han-dling time for individual floral visits, indicating immediate assessment of pollen re-turns.

The present study examined whether bees foraging on

Solanum wendlandii

Hook.selectively visited younger (pollen-rich) flowers over older flowers and also spent moretime foraging on younger than older flowers. In addition, a pollinator exclusion exper-iment was conducted to assess whether, among young flowers, floral handling timevaried between “virgin” vs. previously visited flowers.

182


83(2) June, 2000

M

ATERIALS

AND

M

ETHODS

Flowers of

S

.

wendlandii

have purple petals that fade with age (flowers probablylast no more than 3 days). Five, large, tubular anthers are present with distal sectionspurple and basal parts yellow. Nectaries are absent, and flowers offer only pollen,which is released through 2 minute apical pores per anther (Michener 1962). Bees areable to gather the pollen efficiently only via sonication or buzzing of the anthers. Vis-iting bees grasp the anther cone and rapidly contract their indirect flight muscles(thus producing an audible sound or buzz), which transfers vibrations to the anthersand expels pollen onto the bee. Bees then groom and transfer the pollen to specialstructures (or corbiculae) on their hind legs for storage.

The study was conducted at the Las Cruces Biological Station of the Organizationfor Tropical Studies in southwestern Costa Rica. The patch of

S

.

wendlandii

observedwas growing on a stone wall in the station clearing immediately adjacent to mixed pri-mary and secondary pre-montane forest (elevation 1,100 m). Data were gathered from18 September to 4 October, 1997, an interval falling toward the end of a 9-month rainyseason. Observations were restricted to sunny days with air temperatures ranging be-tween 20–23

°

C.Visits by 3 species of buzz-pollinating bees were recorded continuously for individ-

ually tagged flowers of

S

.

wendlandii

between 0700-1100 hours over 4 days. Based onpreliminary observations, the peak period of floral visitation occurred between 0800-1000 hours. Buzz-pollinators were never seen at the flowers prior to 0715 hours, andthus our data most likely describe complete sequences of bee visitation to the focalflowers. The 3 principal buzz-pollinating species included (in order of increasing bodysize) a halictid,

Pseudaugochloropsis graminea

(Fabricius), a euglossine,

Euglossaerythrochlora

Moure, and a bumblebee,

Bombus pullatus

(Fkln.). Several other

Eu-glossa

spp. were observed sonicating the flowers, but these were infrequent visitors.On a given morning, 1-8 observers recorded the time of day, bee species, and dura-

tion of pollen collecting (to the nearest s) for individual foraging visits to 2-5 pairs offlowers, each pair consisting of a “new” (i.e., newly opened the same day as the obser-vation) and an “old” (i.e., open for at least 1 day prior to the observation) flower. Op-erationally, the duration of pollen collection (here termed handling time) was equatedwith audible buzzing of the anthers. Flower age was determined by tagging stems 1day before making observations with a small piece of green tape. Tags were placed be-low fully developed buds (set to open the following morning and be “new” flowers) andthe closest (already) open flower. Paired new and old flowers were less than 30 cmapart in all cases.

We compared handling times of 1) “initial” visits (first or second visit observed overall bee species) to new vs. old flowers, 2) initial vs. “final” visits (seventh or later visitobserved over all bee species) to new flowers, and 3) final visits to new flowers vs. ini-tial visits to old flowers. Note that the terms “initial” and “final” refer to the sequentialorder of visits compiled over all bee species

for particular flowers

and not to inter-flo-ral visitation sequences

for particular bees

. Counting visits independently of speciesidentity provided only a rough index of pollen depletion, since possible interspecificdifferences in pollen removal were not documented. Note also that initial visits to oldflowers refer, not to their first or second visits in absolute terms, but to the first or sec-ond visits recorded during our observations. Being at least 1 day old, old flowers hadmost likely been visited multiple times on the day(s) prior to our observations (a validassumption given that 49 of the 52 new flowers we observed received 2 or more visitsby sonicating bees).

As shown below, for 2 of the species there was a significant reduction in the timespent gathering pollen at new flowers through the morning, i.e., between early (0700-


Solanum 183

0900 hours) and late (0900-1100 hours) observation periods. To determine whetherthis decrease was time-dependent (foraging rule: if late morning, spend less time pernew flower) vs. resource-dependent (foraging rule: if pollen supply depleted, spendless time per new flower), we placed fine mesh bags on a total of 24 flower buds 1 daybefore opening. Buds were enclosed completely with nylon mesh screening secured tothe stem with a wire clasp. Bags were sufficiently large that the exposed anthers ofnewly opened flowers were well below the screening, out of reach of potential pollina-tors. For each bagged flower, we tagged (but did not bag) a nearby (within 30 cm) fullydeveloped bud. The following morning we removed the bags at 0900 hours and contin-uously recorded visits for the next 2 h; unbagged, new flowers were observed contin-uously from 0700-1100 hours. On a given day, an observer monitored visitscontinuously to 8 pairs of new flowers (i.e.,

1 bagged, 1 unbagged). This experimentwas completed over 3 mornings.

Upon completing behavioral observations, we collected flowers to estimate pollensupplies for (1) new, unvisited flowers (buds were bagged 1 day before opening andbagged flowers were collected the following day, (2) new, visited flowers (buds weretagged but not bagged 1 day before opening and flowers were collected the followingmorning), and (3) old, visited flowers (already open flowers were tagged and collectedthe following day. All flowers were collected at 1100 hours, and anthers were removedwith a forceps, placed in a drying oven (55

°

C) for 24 h, and weighed to the nearest0.001 g using a Mettler AE260 Analytical Balance.

Variation in floral handling times within and among bee species and antherweights among flowers was first analyzed using one-way ANOVA to detect significantinter-group variation overall and then the Tukey test to identify significant differ-ences between specific groups (log

10

transformed values were used in both tests to con-trol for the association between mean and variance; Zar 1996). However, data on floralvisitation were not normally distributed (even after log

10

transformation), and conse-quently for this parameter within-species variation was first analyzed using theKruskal-Wallis test (ANOVA by ranks) and then the non-parametric Dunn’s test(Daniels 1990). In all cases, there was direct correspondence between the 2 types oftests: when ANOVA (or the Kruskal-Wallis test) detected (or, conversely failed to de-tect) significant variation overall, the Tukey test (or Dunn’s test) also identified (or,conversely, failed to identify) specific, significant inter-group differences. Conse-quently, only the results of the Tukey tests (or Dunn’s tests) are presented. For pair-wise comparisons in the flower bagging experiment, we used the Mann-Whitney test,a nonparametric analogue of the Student’s t test (Zar 1996).

R

ESULTS

The 3 species displayed the same temporal pattern of abundance (Fig. 1). Few flo-ral visits occurred before 0730 hours. Activity peaked between 0730-0830 hours andthen declined steadily until the end of observations at 1100 hours. Despite frequentchecks, no bees were seen at the flower patch in the afternoon or early evening. For all3 species, approximately

⅔

of all floral visits occurred prior to 0900 hours.The species also exhibited the same pattern of preference for floral age (Table 1).

Individuals of all species preferred new over old flowers in the early morning butshowed no such preference later in the morning. The early morning preference fornew flowers was quite pronounced: new flowers were, on average, visited 3-4 timesmore often than old flowers. Reflecting the decline in overall activity, visitation ratesto new flowers decreased significantly between early and late morning for all species.Visitation rates declined with time for old flowers as well, though this difference wasnot significant for any bee species.

184


83(2) June, 2000

During initial visits,

P

.

graminea

and

E

.

erythrochlora

spent significantly moretime collecting pollen from new flowers than old ones (Table 2). Also, when visitingnew flowers, these same species spent significantly more time collecting pollen duringinitial visits than final ones. These same trends were apparent for

B

.

pullatus

as well,although they were not statistically significant (Table 2). Handling times for initialvisits to new flowers were similar to initial visits to old flowers in all 3 species (Table 2).

Regarding interspecific comparisons,

P

.

graminea

spent significantly more timecollecting pollen during initial visits to new flowers than

E

.

erythrochlora

or

B

.

pulla-tus

(Table 2). Handling times did not vary among species for final visits to new flowersor initial visits to old flowers.

The incidence of initial and final visits to new flowers was not independent of thetime of day, and the majority (51/69 = 74% over all species) of final visits to new flow-ers occurred after 0900 hours. Thus, as noted above, the decrease in buzzing durationsbetween initial and final visits to new flowers noted for

P

.

graminea

and

E

.

erythro-chlora

may have reflected a time-dependent, rather than a resource-dependent, shiftin foraging behavior. However, in the flower bagging experiment mean handling timeswere greater for the initial visits to “virgin” new flowers than for the concurrent, finalvisits to unbagged, new flowers for all 3 species -

P

.

graminea

: 26 s vs. 11 s, respec-tively (n

1

= 11, n

2

= 8);

E

.

erythrochlora

: 15 s vs. 6 s, respectively (n

1

= 6, n

2

= 6);

B

.

pul-latus

: 18 s vs. 8 s, respectively (n

1

= 17, n

2

= 17); P < 0.05 in all cases, Mann-Whitneytest). Pollen-collection times for initial visits to previously bagged, new flowers weresimilar to those recorded for initial visits to unmanipulated, new flowers in the earlymorning for all 3 species (P > 0.05 in all cases; Mann-Whitney test). Despite these

Fig. 1. Total numbers of visits recorded over all focal flowers (N = 52 pairs of newand old flowers) in relation to time of day.


Solanum 185

findings, bees visited virgin and previously available new flowers with similar fre-quency: over all 3 species, the mean numbers of visits recorded between 0900-1100hours were 1.85 (

±

1.2) and 2.0 (

±

1.3) for virgin and unbagged new flowers, respec-tively (P > 0.05; Mann Whitney test).

Anther (mg dry) weights were significantly greater for new, unvisited flowers (x–

±

1 SD = 149

±

14 mg) than either new, visited (131

±

12 mg) or old, visited (125

±

9mg) flowers (P < 0.05 in both cases; Tukey test using transformed [log

10

x] data).Among visited flowers, anthers from new flowers weighed more than those from oldflowers, although this difference was not statistically significant (P > 0.05; Tukey testusing transformed [log x

10

] data).

D

ISCUSSION

Foraging choices frequently involve “non-energetic” benefits (Rasheed and Harder1997). That is, although their collection requires energy expenditure, the resources donot provide energy directly to the forager but serve other functions. For bees, pollenharvesting yields non-energetic benefits: pollen serves primarily as a protein sourcefor developing larvae, while nectar is the chief energy source for flight and other ac-

T

ABLE

1

.

V

ISITATION

RATES

TO

NEW

AND

OLD

FLOWERS

DURING

EARLY

(0700-0900

HOURS

)

AND

LATE

(0900-1100

HOURS

)

MORNING

.

VALUES

GIVEN

REPRESENT

MEAN

NUM-BER

(

±

1 SD)

OF

VISITS

PER

FLOWER PER 2 H SAMPLING PERIOD; N = 52 FLOWERSIN ALL CASES. VALUES WITHIN A ROW FOLLOWED BY THE SAME LETTER ARE NOTSIGNIFICANTLY DIFFERENT FOLLOWING THE DUNN’S TEST (P = 0.05).

Early Late

New Old New Old

P. graminea 1.4a (1.4) 0.3b (0.6) 0.6b (1.0) 0.25b (0.6)

E. erythrochlora 0.8a (0.9) 0.3b (0.4) 0.4b (0.6) 0.2b (0.5)

B. pullatus 1.8a (1.5) 0.5b (0.5) 0.9b (1.0) 0.5b (0.5)

TABLE 2. HANDLING TIMES AT NEW AND OLD FLOWERS DURING INITIAL (FIRST OR SEC-OND) AND FINAL (SEVENTH OR GREATER) VISITS TO INDIVIDUAL FLOWERS.MEAN VALUES TO NEAREST S (±1 SD, N) ARE PROVIDED. VALUES WITHIN AROW FOLLOWED BY SAME LOWERCASE LETTER ARE NOT SIGNIFICANTLY DIF-FERENT; VALUES WITHIN A COLUMN FOLLOWED BY THE SAME UPPERCASE LET-TER ARE NOT SIGNIFICANTLY DIFFERENT FOLLOWING THE TUKEY TEST (USINGTRANSFORMED [LOG10 X] DATA; P = 0.05).

New Old

Initial Final Initial

P. graminea 37aA (24, 34) 15bA (10, 24) 10bA (6, 19)

E. erythrochlora 18aB (11, 19) 8bA (4, 14) 6bA (4, 14)

B. pullatus 14aB (13, 45) 9aA (6, 31) 8aA (6, 29)


tivities (Heinrich 1979). In addition to the fact that nectar is an easily measured andmanipulated resource, the research emphasis on nectar foraging by bees deriveslargely from the working assumption that energy is an appropriate fitness “currency”for foraging animals. Nonetheless, energy-mediated constraints on bee activity mightbe expected to generate a common pattern of foraging behavior regardless of the typeof resource collected (i.e., energy- or non-energy-based). In fact, Rasheed and Harder(1997) found that pollen-gathering bumblebees foraged in a manner qualitativelysimilar to that reported for nectar-gatherers, i.e., in both cases, bees maximized for-aging efficiency or benefit-to-metabolic cost ratio.

The present study provides additional evidence that pollen-foraging bees modifytheir behavior in response to anticipated and actual pollen returns from individualflowers. All 3 species studied preferred new over old flowers in the early morning. Aspetal color changed greatly with age, it seems likely that bees used reflectance pat-terns (in the visual or ultraviolet spectra) as long-distance cues of pollen supplies. Bylate morning, however, the bees did not differentiate between new and old flowers,and, as the measurements of anther weight suggest, this shift reflected an increasedsimilarity in the pollen abundance of new and old flowers. Thus, the bees presumablycould distinguish between new and old flowers late in the morning but, owing to re-duced pollen levels in the new flowers, “ignored” this distinction. Additional data onapproach and rejection probabilities for flowers of different ages are required to con-firm color-based discrimination in the early morning.

In contrast, bees were apparently unable to make long-range assessment of pollenavailability among new flowers. In the bagging experiment, bees visited virgin andunbagged new flowers with equal frequency. This finding was not unexpected, sincethe pollen is concealed within minute pores on the anther (Michener 1962). Potentialcues, such as pollen odor (Buchmann 1983) or “bruise marks” left on the anthers byprevious visitors (J. Cane, pers. comm.), were presumably either absent or weak. Con-sistent with other studies, therefore, our data suggest that long-distance assessmentof nectar (Neff and Simpson 1990) or pollen (Pellmyr 1988) rewards may depend ex-clusively on “gross” features of floral morphology, such as overall shape or color.

Upon alighting, bees clearly adjusted their harvesting effort to match pollen avail-ability. In the bagging experiment, handling times for all 3 species were significantlylonger for virgin, new flowers than for unbagged, new flowers. Observations for P.graminea and E. erythrochlora showed that initial visits to new flowers were signifi-cantly longer than either initial visits to old flowers or final visits to new flowers.Other studies (Pellmyr 1988, Buchmann and Cane 1989, Harder 1990) report thissame trend, supporting the general observation that, as bee visitation continues in aflower patch, individual foragers encounter diminishing pollen returns per flower andtherefore spend less time at individual flowers.

In sum, our data indicate that pollen-collecting bees are sensitive to varying re-source levels within individual flowers and respond by selecting and intensively han-dling more rewarding flowers. Thus, while the nature of the rewards differ, pollen-andnectar-foraging bees appear similar in attempting to maximize the rate of resourcecollection. Instead of maximizing the rate of fuel (nectar) intake, however, pollen-col-lecting bees forage in a manner that increases the rate of pollen delivery to developinglarvae.

ACKNOWLEDGMENT

We thank Luis Diego Gomez for weather data and for identifying the plant, RaulRojas for much logistical support, Jim Ackerman for supplying a key for euglossines,

Shelly et al.: Buzzing Bees on Solanum 187

the staff of the Instituto Nacional de Biodiversidad for assistance in identifying thebees, and Emma and Miranda Shelly for assistance with data collection. We alsothank Jim Cane for supplying references and encouragement and Jack Neff for help-ful comments on an earlier draft.

REFERENCES CITED

BUCHMANN, S. L. 1983. Buzz pollination in angiosperms, pp. 73-113 in C. E. Jones andR. J. Little (eds.). Handbook of experimental pollination biology. Van NostrandReinhold, New York.

BUCHMANN, S. L., AND J. H. CANE. 1989. Bees assess pollen returns while sonicatingSolanum flowers. Oecologia (Berl) 81: 289-294.

CRESSWELL, J. E. 1990. How and why do nectar-foraging bumblebees initiate move-ments between inflorescences of wild bergamot Monarda fistulosa (Lamiaceae).Oecologia (Berl) 82: 450-460.

DANIELS, W. W. 1990. Applied nonparametric statistics. PWS-KENT Publishing, Boston.DUKAS, R., AND L. A. REAL. 1993. Effects of recent experience on foraging decisions by

bumble bees. Oecologia (Berl) 94: 244-246.GIURFA, M., AND J. NUNEZ. 1992. Foraging by honeybees on Carduus acanthoides:

pattern and efficiency. Ecol. Entomol. 17: 326-330.GORI, D. F. 1989 Floral color change in Lupinus argenteus (Fabaceae): why should

plants advertise the location of unrewarding flowers to pollinators? Evolution43: 870-881.

HARDER, L. D. 1990. Behavioral responses by bumble bees to variation in pollen avail-ability. Oecologia (Berl) 85: 41-47.

HAYNES, J., AND M. MESLER. 1984. Pollen foraging by bumblebees: foraging patternsand efficiency on Lupinus polyphyllus. Oecologia (Berl) 61: 249-253.

HEINRICH, B. 1979. Bumblebee economics. Harvard University Press, Cambridge, MA.HODGES, C. M., AND R. B. MILLER. 1981. Pollinator flight directionality and the as-

sessment of pollen returns. Oecologia (Berl) 50: 376-379.KREBS, J. R., AND R. H. MCCLEERY. 1984. Optimization in behavioural ecology, pp. 91-

121 in J. R. Krebs and N. B. Davies (eds.). Behavioural ecology: an evolutionaryapproach. Blackwell Scientific Publications, London.

MICHENER, C. D. 1962. An interesting method of pollen collecting by bees from flowerswith tubular anthers. Rev. Biol. Trop. 10: 167-175.

NEFF, J. L., AND B. B. SIMPSON. 1990. The roles of phenology and reward structure inthe pollination biology of wild sunflower (Helianthus annuus L., Asteraceae).Israel. J. Bot. 39: 197-216.

PELLMYR, O. 1988. Bumble bees (Hymenoptera: Apidae) assess pollen availability inAnemonopsis macrophylla (Ranunculaceae) through floral shape. Ann. Ento-mol. Soc. America 81: 792-797.

PYKE, G. H. 1979. Optimal foraging in bumblebees: rules of movement between flow-ers within inflorescences. Anim. Behav. 27: 1167-1181.

RASHEED, S. A., AND L. D. HARDER. 1997. Foraging currencies for non-energetic re-sources: pollen collection by bumblebees. Anim. Behav. 54: 911-926.

WADDINGTON, K. D., AND B. HEINRICH. 1979. The foraging movements of bumblebeeson vertical “inflorescences”: an experimental analysis. J. Comp. Physiol. 134:113-117.

ZAR, J. H. 1996. Biostatistical analysis. Prentice-Hall, Upper Saddle River, New Jersey.

188


83(2) June, 2000

FIRST RECORD OF RICE THRIPS (THYSANOPTERA: THRIPIDAE) IN TRINIDAD, WEST INDIES

G

RAHAM

W

HITE

Caroni Research StationWaterloo Road, Carapichaima, Trinidad & Tobago, West Indies

Rice Thrips,

Stenchaetothrips biformis

Bagnall (Thysanoptera: Thripidae) wasfirst observed in Trinidad at the rice department of Caroni (1975) Ltd.

1

, on July 291997.

The specimens were identified by A. K. Walker of the British Museum of NaturalHistory. The Identification was facilitated by CARINET, the Caribbean loop of BionetInternational.

The thrips were observed on seedling rice (

Oryza sativa

L grown under irrigatedconditions. At the time of discovery, the rice plants were showing silver streaks typicalof the damage caused by


.Prior to this observation, monitoring of other seedling pests

Hydrellia

sp. (Ephy-dridae: Diptera) and

Sogatodes orizicola

(Muir) (Delphacidae: Homoptera) had beenin progress so it is probable that the thrips were discovered soon after their arrival inthe area.

The field in which the thrips were collected was subsequently sprayed with insec-ticide for control of

Hydrellia

and the fate of the thrips is unknown. Since the discov-ery, two rice crops have been planted. No more thrips have been found duringmonitoring exercises.

The pest complex at the Caroni Rice Department at present includes

Hydrellia

sp.,

Tagosodes

spp.,

Diatraea saccharalis

(F.) (Pyralidae: Lepidoptera),

Rupela albinella

(Cram.) (Pyralidae: Lepidoptera), and

Oebalus poecilus

(Dall.) (Pentatomidae: Het-eroptera).


may be controlled by flooding. Should the species be-come economically important their presence may compromise the current practice ofdraining field for control of

Hydrellia

sp.This record follows the recent arrival of


in Guyana inJuly-August 1994 (Munroe 1995), and Venezuela, near Calabozo, January-February1995 (Cermeli et al. 1995). Calabozo is roughly 700 km west-southwest of Trinidad.

S

UMMARY

Specimens of Rice Thrips,


Bagnall (Thysanoptera:Thripidae) was collected on seedling rice in Trinidad, West Indies. This constitutes thefirst record of rice thrips in Trinidad.

R

EFERENCES

C

ITED

C

ERMELI

, M., E. G

ARCÍA

,

AND

M. G

AMBOA

. 1995:


(Bagnall)(Thysanoptera: Thripidae) nueva plaga del arroz (Oryza sativa L.) en Venezu-ela. Boletín de Entomología Venezolana N.S. 10(2):209-210.

M

UNROE

, L. 1995. A new pest in Guyana’s rice fields.

Caraphin News No. 11

pp. 1-2.

1

Caroni (1975) Ltd. Is a state owned agroprocessing company which is the largest producer ofrice in Trinidad.

Scientific Notes

189

AN AQUATIC BARRIER TRAP FOR MONITORING ADULTRICE WATER WEEVILS (COLEOPTERA: CURCULIONIDAE)

R

AYMOND

L. H

IX

1

, D

ONN

T. J

OHNSON

1

AND

J

OHN

L. B

ERNHARDT

2

1

University of Arkansas, Dept. of Entomology321 Agri Building, Fayetteville AR 72701

2

University of Arkansas, Rice Research and Extension CenterPO Box 351, Stuttgart AR 72160

The rice water weevil,

Lissorhoptrus oryzophilus

Kuschel (Coleoptera: Curculion-idae), is a pest of rice in the southeastern U.S. and California. Although adult weevilsfeed on rice leaves causing longitudinal scars, larvae feed on the roots causing eco-nomic injury by reducing yield. Carbofuran (Furadan®, FMC Corp., Philadelphia, PA)has been used to manage rice water weevil larvae since the late 1960’s. However, car-bofuran is no longer legal for use in rice. The new alternative insecticides, lambda-cy-halothrin (Karate®, Zeneca Inc., Wilmington, DE) and diflubenzuron (Dimilin®,Uniroyal Chemical Co., Middlebury, CT) require application within 10 d after perma-nent flood (Bernhardt 1997). Lambda-cyhalothrin is targeted against adult weevils,and diflubenzuron is targeted against eggs. The two scouting methods used to deter-mine the need for carbofuran application against larvae were the leaf scar methodand larval core sample (Tugwell & Stephen 1981, Morgan et al. 1989). Inspection forleaf scarring from adult feeding is inadequate, and the inspection of rice roots for lar-vae is too late in determining the need of an insecticide application with the newer in-secticides. Therefore, a new scouting method is urgently needed.

A double-ended barrier trap was developed based on weevil swimming behaviorand tested on F

1

adult weevils. In a preliminary test on 22 July 1998, 16 double-endedbarrier traps were placed next to rice plants in a small late rice plantation (Hix et al.1999). Barrier trap catch means of adult weevils were 73.9 (

±

9.4 SE) per trap on 23July 1998 and 54.4 (

±

6.4 SE) per trap on 24 July 1998. The rice root core sample meanfor this bay was 72.9 (

±

7.0 SE) larvae per core on 1 August 1998. The Arkansas eco-nomic threshold is 10 larvae per core.

In this note, we describe how to assemble this trap from the materials depicted inFig. 1a. When appropriate, English measurements are listed in parentheses.

Aluminum screen or aluminum flashing—35 cm

´

11 cm2 Boll weevil traps with collecting cups—Technical Precision Plastics,

Mebane, NC4 Fishing bobbers—5 cm (2 in)2 PVC pipes—10.5 cm

´

2.1 cm (0.5 in schedule 40)8 Plastic cable ties—10.2cmEpoxy20 gauge stainless steel wireDowel rod if aluminum screen is used - 35 cm

´

0.5 cm (

3

/

16

in)

Trap construction begins by marking the aluminum screen or flashing 6.0 cm fromthe corners on the long sides and 3.0 cm from the corners on the short sides. The cor-ners are then trimmed off by cutting between these marks providing a barrier withcorrect taper for insertion into the boll weevil traps. If screen is used for the barrier,a dowel should be wired to the screen at 3.0 cm from the upper edge of the barrier withstainless steel wire (Fig. 1a). This imparts rigidity to the trap allowing it to float prop-

190


83(2) June, 2000

erly. If aluminum flashing is used for the barrier, 2 small holes should be drilled oneach of the 4 tapered edges. The barrier is inserted into each boll weevil trap usingplastic tabs (marked by arrows in Fig. 1b) on the boll weevil traps for proper barrierorientation and trap strength. The barrier is then wired to the boll weevil traps withstainless steel wire. The float system for the trap is made by epoxying the bottom ofthe fishing bobbers into each end of 2 PVC pipes. The top holes of the bobbers aresealed with epoxy to make them water tight. Hot glue could be used for the float as-

Fig. 1. (a) Boll weevil tops with collecting cups, 5 cm (2 in) fishing bobbers, PVCpipe, 10.2 cm plastic cable ties, screen (cut to specifications), and dowel rods with ar-row indicating points to which they are attached to screen; (b) tabs (denoted by ar-rows) in boll weevil traps may be used for proper screen alignment and trap strength.

Scientific Notes

191

sembly but is inferior to epoxy for durability in the field. A plastic cable tie is attachedto each of the 4 braces on the upper side of the trap assembly. Plastic cable ties arethen looped around PVC pipe of the floats and through each cable tie previously at-tached to the braces of the boll weevil traps (Fig. 2a). Traps are positioned in the fieldvia flags placed through the space in one of the trap ends (Fig. 2). The trap should be

Fig. 2. (a) Cable ties are looped around the PVC (denoted by double arrows) pipeof the float assembly and cable ties previously attached to the trap braces on the up-per side of the trap assembly, the single arrow indicates space through which a flagmaybe placed to hold the trap in position; (b) placement and appearance of properlyplaced trap in the field.

192


83(2) June, 2000

suspended beneath the float assemblies allowing about 1.5 cm of the barrier to emergefrom the water.

Traps should be checked every 24 h or 36 h for weevils. Weevils can survive beingsubmerged for about 96 h, but frequent inspection of the traps will minimize a buildupof debris and decomposition of other insects caught in these traps. This trap designmight be useful in collecting other species of aquatic weevils and small to mediumsized aquatic insects.

S

UMMARY

The construction of rice water weevil barrier traps is described. These traps areconstructed by attaching a barrier between boll weevil traps and suspending them inthe water by flotation devices. Floating barrier traps were designed to monitor adultrice water weevils from the first day of permanent flood until ten days post flood.

A

CKNOWLEDGMENT

This project was funded in part by the University of Arkansas Experiment Stationand the Arkansas Rice Promotion Board.

R

EFERENCES

C

ITED

B

ERNHARDT

, J. L. 1997. Control of the rice water weevil with Karate. Arthropod Man-agement Tests. 22: 289.

H

IX

, R. L., D. T. J

OHNSON

, J. L. B

ERNHARDT

, J. D. M

ATTICE

,

AND

B. L. L

EWIS

. 1999.Trapping adult rice water weevils with floating cone and barrier traps,

in

TheB. R. Wells Rice Research Studies 1998. Arkansas Agri. Exp. Stat. Res. Series468: 135-141.

M

ORGAN

, D. R., N. P. T

UGWELL

,

AND

J. L. B

ERNHARDT

. 1989. Early rice field drainagefor control of rice water weevil (Coleoptera: Curculionidae) and evaluation of anaction threshold based upon leaf-feeding scars of adults. J. Econ. Entomol. 82:1757-1759.

T

UGWELL

, N. P.,

AND

F. M. S

TEPHEN

. 1981. Rice water weevil

Lissorhoptrus oryzophi-lus

seasonal abundance, economic levels, and sequential sampling plans. Ar-kansas Agri. Exp. Stat. Bul. 849.

Scientific Notes

193

APPLICATION OF ALARM PHEROMONE TO TARGETS BY SOUTHERN YELLOWJACKETS (HYMENOPTERA: VESPIDAE)

H

AL

C. R

EED

1

AND

P

ETER

J. L

ANDOLT

USDA, ARS, 5230 Konnowac Pass Rd., Wapato, WA 98951, USA

1

Current address: Department of Biology, Oral Roberts University, Tulsa, OK 74171

Alarm pheromones have been demonstrated for a number of species of social Vesp-idae including several hornets and yellowjackets (Vespines) (Landolt et al. 1997).Maschwitz (1964a, b) first demonstrated alarm pheromone responses in the yellow-jackets

Vespula vulgaris

L. and

V. germanica

(Fab.) in response to crushed wasps andbody parts. Pheromone-mediated alarm has since been observed in other vespines:

Dolichovespula saxonica

(Fab.) (Maschwitz 1984), the

southern yellowjacket

V. squa-mosa

(Drury) (Landolt & Heath 1987, Landolt et al. 1999), the eastern yellowjacket

V. maculifrons

(Buysson) (Landolt et al. 1995),

Provespa anomala

Saussure(Maschwitz & Hanel 1988), and

Vespa

crabro

L. (Veith et al. 1984). 2-Methyl-3-butene-2-ol was identified as a component of the alarm pheromone of

V. crabro

(Veithet al. 1984), and N-3- methylbutylacetamide was isolated and identified as an alarmpheromone of the southern and eastern yellowjackets (Heath & Landolt 1988,Landolt et al. 1995).

The source of alarm pheromones in social wasps generally is the venom, althoughthe head is implicated as an additional source of alarm pheromone for

V. vulgaris

(Al-diss 1983) and

V. squamosa

(Landolt et al. 1999). Alarm behavior in

V. germanica

and

V. vulgaris

occurred in response to the squashed sting apparatus, sting sac, and sol-vent extract of the sting sac (Maschwitz 1964b) and in

D. saxonica

as a response tocrushed venom glands (Maschwitz 1984). Veith et al. (1984) stimulated alarm in

V.crabro

with squashed venom sacs or venom. Landolt & Heath (1987) isolated analarm pheromone of

V. squamosa

in solvent extracts of the venom sac and glands. Al-diss (1983) observed alarm in

V. vulgaris

in response to crushed conspecific heads, andLandolt et al. (1999) stimulated alarm and attack in the southern yellowjacket witha solvent extract of conspecific heads. Alarm pheromones known in several species of

Polistes

also originate in the venom (reviewed by Landolt et al. 1997).Despite repeated demonstrations of alarm responses of social wasps to conspecific

body parts and extracts of body parts, the alarm signalling process itself remains un-known. We do not know how wasps release alarm pheromone. It is hypothesized thatalarm pheromone in venom is released when wasps spray venom or is deposited whenwasps sting (Aldiss 1983, Maschwitz 1964b, Greene et al. 1976). An alarm pheromoneoriginating in the head of workers may be released at the mouthparts and applied orevaporated from the mandibles (Landolt et al. 1999).

We report here experimental evidence that an alarm pheromone is deposited on asubstrate or target when attacked by southern yellowjackets. We also demonstratedpersistence of that alarm pheromone activity that is uncharacteristic of alarm phero-mones generally. Alarm pheromones in social insects typically are quite volatile andshort-lasting, an advantage in permitting normal colony activities to resume once athreat has passed (Matthews & Matthews 1978).

Preliminary observations that led to this study indicated possible contaminationof protective clothing and equipment following attacks by southern yellowjackets.This included seemingly unprovoked responses by yellowjackets to investigators oneor more days following other experiments and a residual odor on material and objects

194


83(2) June, 2000

that had been attacked by workers. We conducted experiments to determine if attack-ing wasps leave a material that elicits alarm and attack in other workers.

Observations and experiments with southern yellowjackets were made in AlachuaCounty, Florida. All testing was done with vigorous underground colonies. The bioassayfor these tests involved a cork (3.7 cm

´

3.7 cm) connected to a wooden dowel (3 m longby 1.2 cm diam.) with 2 interlocking eye hooks screwed into the dowel and cork. Theeye hooks permitted movement of the cork on the dowel that made it easier to detectwasp contact with the target. Three colored push pins (red, blue, and green) werestuck into the bottom of the cork. This target (cork with pins) was waved from side toside about 0.3 m in front of a colony entrance to induce attack from guard workerspresent in the nest entrance. During these presentations, workers generally attackedthe cork as well as the hooks and push pins. During attacks workers made stingthrusts and also appeared to bite the target, with their mandibles open and contact-ing the cork, hooks, or pins.

An experiment was conducted to determine if freshly attacked targets elicit alarm,as evidence of the deposition of alarm pheromone by wasps onto targets during earlierattacks. Corks were first presented at nest entrances until hit by wasps, with 6-10wasps contacting the cork. This treated cork was placed in a glass jar in an ice chestand transported to the laboratory and placed in a freezer. This procedure was re-peated to accumulate 5 treated corks. A new dowel and cork was used for each repli-cate of this procedure. A treated cork was subsequently exposed to ambient fieldconditions for 3 min and was then presented to a second test colony. The cork wasmoved slowly to

⅓

m upwind of the colony where it remained for the assay duration.Alarm behavior and hits to the cork were noted for 2 min, with the use of a tape re-corder. As a control, an unexposed cork and dowel were presented in the same mannerbefore each assay of a treated cork. Five treated corks were each tested 4 - 5 h apartover several days. Numbers of hits and landings on the five treated corks ranged from1 to 76 (mean

±

SE = 33.4

±

30.4), significantly greater than the no hits or landingsthat occurred on the five control corks (p = 0.036 by Student’s t test).

A second experiment was conducted to determine if the alarm activity of materialapplied by yellowjackets to the cork remained active after exposure at ambient tem-peratures. The bioassay protocol was the same as above except that each treated corkwas aired in the laboratory for 15 h before field tests. The five treated corks weretested over 4 different days, with 2 tested 2.5 h apart on one day. Again, the five con-trol corks elicited no response, with no evidence of alarm and no contacts of yellow-jackets with the corks. The 15 h old corks however, elicited alarm and attack to thecorks. Numbers of hits and landings ranged from 4 to 83 per assay (mean

±

SE = 35

±

33.9), significantly greater than the response to the control (t = 2.33, p = 0.04).These two experiments demonstrate that alarmed

V. squamosa

apply chemicals totargets that they attack and that such attacked targets may subsequently remain ac-tive in eliciting alarm and attack responses from southern yellowjackets. At this time,the source of the pheromone applied is not known. During stinging attacks on thecorks, hooks, and pins, venom could have been applied to those surfaces. Also, waspsattacking the targets were seen to bite on the cork, hooks, and pins, with the possibil-ity that other alarm pheromones from gnathal or cephalic glands which open to themouthparts (Landolt & Akre 1979) could be applied to mark the targets. The southernyellowjacket is known to possess alarm pheromone both in the venom (Landolt &Heath 1987, Heath & Landolt 1988) and in the head (Landolt et al. 1999).

The possible adaptive significance of such a long lasting alarm signal is apparentwhen considering the functions of sting autotomy in social insects, the lack of stingautotomy in yellowjackets, and the nature of predators of social wasps and bees. Sting

Scientific Notes

195

autotomy is the loss of the sting and venom sac after a stinging episode, such as occursin the honey bee,

Apis mellifera

L. (Free 1987) and in the social wasp

Polybioides raph-igastra

(Saussure) (Sledge et al. 1999). In both of these species, sting autotomy prob-ably permits the prolonged release of alarm pheromone from the sting apparatusfollowing stinging, marking the intruder and focusing subsequent attacks (Free 1987,Sledge et al. 1999). A similar strategy has been suggested for species of

Apis

(Pickettet al. 1982, Schmidt et al. 1997), including

A. mellifera and

Apis cerana

(Fab.), basedon the large quantities of eicosenol in the sting apparatus. It is hypothesized that thiscompound serves as a carrier to prolong the release of more volatile pheromone com-pounds and to mark an intruder to focus the defending bees. Although the southernyellowjacket does not exhibit sting autotomy, the deposition of a long lasting alarmpheromone during stinging attacks on vertebrate predators may similarly serve tochemically mark the animal. This would focus attacks on the intruder and also alertthe colony if and when this predator approached the nest again.

Several vertebrate predators, such as skunks and raccoons, commonly prey on yel-lowjacket colonies (Akre & Reed 1984). A possible strategy of a vertebrate predator isexemplified by the honey buzzard, a successful nest predator of European yellowjack-ets (Cobb 1979, cited in Akre & Reed 1984). This bird was persistent in its attacks onexcavated subterranean

Vespula

nests over a period of days. The buzzard was drivenaway from nests repeatedly by wasps, but later reapproached the nest to continue theattack. Such an “attack, retreat, reattack” scenario is a likely strategy for other ver-tebrate nest predators. Thus, a long lasting alarm pheromone that marks a predatormay be highly advantageous to yellowjackets. However, caution must be exercised ininterpreting how these results of alarm pheromone persistence from a wooden corkmay relate to alarm pheromone on vertebrate skin, fur, or feathers. Additional exper-imentation with leather or feather targets could help address this question, as wouldchemical analysis of odors emitted by attacked objects.

S

UMMARY

Alarmed southern yellowjacket workers attacking corks placed near colony en-trances applied an alarm pheromone that stimulated alarm and attack behavior inanother colony 3 min or 15 h after pheromone deposition. Observations of wasps at-tacking corks indicated deposition or application of alarm pheromone could be madeboth from the sting and from the mandibles. This long lasting material may serve tomark an attacking vertebrate predator so that it is quickly detected and attackedagain upon its return to a wasp colony.

R

EFERENCES

C

ITED

A

KRE

, R. D.

AND

H. C. R

EED

. 1984. Vespine defense, 3, pp. 55-94,

in

H. R. Hermann,(ed.), Defensive mechanisms in social insects. Praeger, New York, 259 pp.

A

LDISS

, J. B. J. F. 1983. Chemical communication in British social wasps (Hy-menoptera: Vespidae). Ph.D. Dissertation, Univ. Southampton, U.K. 252 pp.

C

OBB

, F. K. 1979. Honey buzzard at wasps’ nest. Brit. Birds 72: 59-64.F

REE

, J. B. 1987. Pheromones of social bees. Cornell Univ. Press, Ithaca, New York.218 pp.

G

REENE

, A., R. D. A

KRE

,

AND

P. J. L

ANDOLT

. 1976. The aerial yellowjacket,

Dolicho-vespula

arenaria

(Fab.): Nesting biology, reproductive production, and behavior(Hymenoptera: Vespidae). Melanderia 26: 1-34.

H

EATH

, R. R.,

AND

P. J. L

ANDOLT

. 1988. The isolation, identification, and synthesis ofthe alarm pheromone of

Vespula squamosa

(Drury) (Hymenoptera: Vespidae)and associated behavior. Experientia 44: 82-83.

196


83(2) June, 2000

L

ANDOLT

, P. J.,

AND

R. D. A

KRE

. 1979. Occurrence and location of exocrine glands insome social Vespidae (Hymenoptera). Ann. Entomol. Soc. America 72: 141-148.

L

ANDOLT

, P. J.,

AND

R. R. H

EATH

. 1987. Alarm pheromone behavior of

Vespula squa-mosa

(Hymenoptera: Vespidae). Florida Entomol. 70: 222-225.L

ANDOLT

, P. J., R. R. H

EATH

, H. C. R

EED

,

AND

K. M

ANNING

. 1995. Pheromonal medi-ation of alarm in the eastern yellowjacket (Hymenoptera: Vespidae). FloridaEntomol. 78: 101-108.

L

ANDOLT

, P. J., R. L. J

EANNE

,

AND

H. C. R

EED

. 1997. Chemical communication in so-cial wasps, pp. 216-235,

in

R. K. Vandermeer, M. Breed, M. Winston, and K. Es-pelie (eds.), Pheromone communication in social insects: ants, wasps, bees, andtermites. Westview Press, Boulder, CO.

L

ANDOLT

, P. J., H. C. R

EED

,

AND

R. R. H

EATH

. 1999. An alarm pheromone from theheads of worker southern yellowjackets,

Vespula squamosa

(Drury) (Hy-menoptera: Vespidae). Florida Entomol. 82: 356-359.

M

ASCHWITZ

, U. W. 1964a. Alam substance and alarm behavior in social Hymenoptera.Nature 204: 324-327.

M

ASCHWITZ

, U. W. 1964b. Gefahrenalarmstoffe und Gefahrenalarmierung bei so-zialen Hymenopteren. Z. Verg. Physiol. 47: 596-655.

M

ASCHWITZ

, U. W. 1984. Alarm pheromone in the long cheeked wasp

Dolichovespulasaxonica

(Hymenoptera: Vespidae). Deutsch Entomol. 31: 33-34.M

ASCHWITZ

, U. W.,

AND

H. H

ANEL

. 1988. Biology of the southeast Asian nocturnal wasp,

Provespa

anomala

(Hymenoptera: Vespidae). Entomol. Generalis 14: 47-52.M

ATTHEWS

, R. W.,

AND

J. R. M

ATTHEWS

. 1978. Insect behavior. John Wiley and Sons,NY, 507 pp.

P

ICKETT

, J. A., I. H. W

ILLIAMS

,

AND

A. P. M

ARTIN

. 1982. (Z)-11-Eicosen-1-ol, an im-portant new pheromonal component from the sting of the honeybee

Apis mel-lifera

L. (Hymenoptera: Apidae). J. Chem. Ecol. 8: 163-175.S

CHMIDT

, J. O., E. D. M

ORGAN

, N. J. O

LDHAM

, R. R. D

O

. N

ASCIMENTO

,

AND

F. R. D

ANI

.1997. (Z)-11-Eicosen-1-ol, a major component of

Apis cerana

venom. J. Chem.Ecol. 23: 1929-1939.

S

LEDGE

, M. F., F. R. D

ANI

, A. F

ORTUNATO

, U. M

ASCHWITZ

, S. R. C

LARKE

, E. F

RANCES-CATA

, R. H

ASIM

, E. D. M

ORGAN

, G. R. J

ONES

,

AND

S. T

URILAZZI

. 1999. Venom in-duces alarm behavior in the social wasp

Polybioides raphigastra

(Hymenoptera: Vespidae): an investigation of alarm behavior, venom volatilesand sting autotomy. Physiol. Entomol. 24: 234-239.

V

EITH

, J., N. K

OENIGER

,

AND

U. W. M

ASCHWITZ

. 1984. 2-methyl-3-butene-2-ol, a ma-jor component of the alarm pheromone of the hornet,

Vespa crabro

. Naturwis-senschaften 71: 328-329.

Scientific Notes

197

LABORATORY TRANSMISSION OF

XANTHOMONAS CAMPESTRIS

PV.

RAPHANI

BY A TARDIGRADE (PARACHELA = MACROBIOTIDAE)

T

HOMAS

G. B

ENOIT

, J

ENNIFER

L

OCKE

, J

AMES

R. M

ARKS

AND

C

LARK

W. B

EASLEY

Department of Biology, McMurry University, Abilene, Texas 79697

Tardigrades are animals currently classified in the phylum Tardigrada and arephylogenetically most closely related to the arthropods (Aguinaldo et al. 1997). Theylive in a variety of microhabitats, being active when a film of moisture is present andbecoming dormant when dry. Their small size (mean length 300-350 m) allows manyof the species to live relatively undisturbed on the surface of a variety of plants andin the soil. The anterior end of the complete digestive system is equipped with sharpstylets and a muscular pharynx which allows a piercing-sucking method of feeding.Some species have a large diameter buccal tube which permits ingestion of solid par-ticles up to the size of fungal spores. Bacteria have often been observed on the surfaceof tardigrades and are ingested by a number of tardigrade genera (Kinchin 1994). Ithas been assumed that these bacteria are food, although symbiosis has been sug-gested (Kinchin 1989, 1993).

During a previous study of lichen-dwelling species of tardigrades, we discoveredthat up to half of the individual tardigrades examined harbored members of the phy-topathogenic bacteria

Xanthomonas

and

Pseudomonas

(Krantz et al. 1999). Evidencedeveloped in that study also suggested that these bacteria dwell inside the tardigrade,perhaps having originated from detritus or food. Furthermore, the bacteria appear tobe shed as the animal moves about. Tardigrades harboring phytopathogenic bacteriamight transport those bacteria to suitable plant hosts where they could then causedisease. Experimental evidence reported here strengthens this interpretation byshowing that in the laboratory tardigrades can acquire and transfer

Xanthomonascampestris

pv.

raphani

Lincoln. Infections can be produced in healthy radish plantsfrom these transferred bacteria.

Xanthomonas campestris

pv.

raphani

ATCC 49079 was obtained from The Ameri-can Type Culture Collection, Rockville, MD. Individual tardigardes,

Macrobiotushufelandi

Schultze

(Parachela: Macrobiotidae), were collected from samples of lichengrowing on trees along the shore of a fresh water lake. The samples were soaked inwater for 3.5 h during which the dormant forms of the tardigrades became active. Thetardigrades were concentrated by filtering the water containing them through 90 or120

m

m mesh screens. They were immediately collected from the screens, rinsed 3times in tap water that had been passed through a sterilizing nylon membrane filter,and then used in transmission experiments.

Xanthomonas campestris

pv.

raphani

was grown in TGCP broth (per liter: yeastextract, 2.5 gm; glucose, 20 gm; peptone, 2.5 gm; NaCl, 1 gm; K

2

HPO

4

, 1 gm; MgSO

4

@7H

2

O, 0.5 gm; CaCO

3

, 10 gm) or on nutrient agar (Difco, Sparks, MD) at 30

°

C. Radish,

Raphanus sativus

L. (‘Cherry Blossom’ seed BWI Bulk Seeds, Inc., U.S.A.), werewashed with 70% (v:v) isopropanol before planting in sterile 30

´

290 mm tubes filledto

⅓

of their volume with moist, sterile potting soil and fitted with sterile spongeplugs. The tubes were incubated at 30

°

C in growth chambers on a continuous 24 hlight cycle.

Tardigrades were inoculated with

X. campestris

pv.

raphani

by adding bacteriafrom a 24 h-old culture to a concentration of approximately 1

´

10

6

cells / ml of waterin which some lichen samples were soaked for 3.5 hours. Confirmation of inoculationwas obtained by previously described methods (Krantz

et al.

1999) as follows. Tardi-

198


83(2) June, 2000

grades were rinsed 3 times in 50

m

l droplets of sterile tap water by using a small wire(Irwin) loop to capture the tardigrades and transfer them from droplet to droplet.They then were placed on the surface of a nutrient agar plate, in the center of a circle(2 cm in diameter) which was marked on the undersurface of the petri dish. The plateswere incubated for 5 days at 30

°

C during which the tardigrades freely moved acrossthe surface of the agar. Microorganisms shed by the tardigrades developed into mac-roscopic colonies during this period. Smooth, round, lemon-yellow colonies typical of

X

.

campestris

pv.

raphani

which arose completely outside of the circle were used to in-oculate leaves of radish seedlings by first dispersing a single colony in 5 ml of sterilewater and subsequently placing 50

m

L of this suspension on the leaf of a radish plant.Isolates which produced leaf spot within 7 days were scored as being

X. campestris

pv.

raphani

originating from inoculation of the tardigrade during soaking of the lichens.Uninoculated tardigrades were tested this way also to determine if

X. campestris

pv.

raphani

was associated naturally with the tardigrade population.To determine if

M

.

hufelandi

could be inoculated with

X. campestris

pv.

raphani

from the lesions on infected radish leaves, single infected leaves were removed asep-tically from separate plants and placed topside-up on separate nutrient agar plates.A single rinsed tardigrade suspended in 50

m

l of sterile water was deposited with a mi-cropipet onto the leaf spot lesion of each leaf. The plates were incubated for 48 h at30

°

C and scored for the presence of

X. campestris

pv.

raphani

colonies on the agaraway from the leaf. The presence of these colonies would indicate that the bacteriahad been transferred off the leaf by the tardigrade. As controls, infected leaves with-out tardigrades and uninfected leaves with tardigrades also were tested.

To test whether tardigrades harboring

X. campestris

pv.

raphani

could facilitatean infection on healthy radish plants, 25 individual radish seedlings were inoculatedeither with a single inoculated or uninoculated tardigrade prepared as describedabove. The plants were incubated as described above for up to two weeks and werescored for the appearance of bacterial leaf spot on the leaf where the tardigrade hadbeen placed.

Every attempt to inoculate

M. hufelandi

with

X. campestris

pv.

raphani

was suc-cessful. In the 5 replicate experiments in which tardigrades were suspended withthese bacteria in water, and subsequently rinsed and placed on nutrient agar plates,17 to 23 suspected colonies of

X. campestris

pv.

raphani

arose per plate. All of thesecolonies were used successfully to produce leaf spot disease in healthy radish plants,confirming that they represented the pathovar used to inoculate the tardigrades andnot naturally-occurring microbiota. Tardigrades which were not inoculated with

X.campestris

pv.

raphani

did not produce colonies capable of causing disease in radishplants. Plates inoculated with the last drops of sterile water used to rinse the tardi-grades produced no colonies.

In all 5 replicates in which uninoculated tardigrades were introduced onto dis-eased radish leaves

lying on the surface of nutrient agar plates, and subsequentlywere allowed to move freely across the surfaces of the plates, 15 to 24 colonies of thepathogen arose per plate, indicating that tardigrades can acquire and move

X.campestris

pv.

raphani

from diseased leaves. Plates in similar experiments using un-infected, non-diseased leaves produced no colonies of

X. campestris

pv.

raphani

nordid plates containing only diseased leaves without tardigrades.

In all 10 replicates in which inoculated tardigrades were introduced onto the leavesof healthy radish plants, the radish plants displayed symptoms of leaf spot diseasewithin 2 weeks. The symptoms were identical to those observed when a washed pure sus-pension of

X. campestris

pv.

raphani

only was used to inoculate radish leaves. Inoculatedtardigrades therefore were able to shed viable

X. campestris

pv.

raphani

which then

Scientific Notes

199

caused an infection on the leaves. Leaves treated with uninoculated tardigrades or cell-free spent TCGP broth from

X. campestris

pv.

raphani

cultures did not become diseased.Tardigrades were easily inoculated with

X. campestris

pv.

raphani

. No failures toinoculate were observed in the experiments. Although it is uncertain where the inoc-ulated bacterial cells resided, either in or on the animal, they appeared to associatequickly and tightly with tardigrades. The xanthan exopolysaccharide produced by

X.campestris

pv

. raphani

may have mediated attachment of the bacteria to the tardi-grade exoskeleton. Or, the tardigrades may have ingested the bacteria while changingfrom the dormant to active form during the soaking period. Because the tardigradeswere not washed free of the attached

X campestris

pv

. raphani

it is possible that a spe-cialized relationship exists between them. Even when tardigrades were placed on thelesions of infected leaves, the pathogen associated with the tardigrades at least wellenough to be transported off of the leaves and across the surface of nutrient agarplates. Perhaps in nature tardigrades can transport

Xanthomonas

spp. through theenvironment, and perhaps even from plant-to-plant, since tardigrades are mobile andtheir small size also would allow them to become airborne.

Xanthomonas

spp. havebeen reported to travel through the environment and possibly from plant-to-plant bya wide variety of means including insects and small airborne particles (Bashan 1985).

The experiments in which inoculated tardigrades were placed on uninfectedplants showed that tardigrades can deposit

Xanthomonas

on a susceptible host, whichthen can become infected. It is unknown whether the pathogen was injected into theleaves during feeding by the tardigrades or merely deposited on the surface duringother activities, or some combination of the two. However, every plant that received asingle, infected tardigrade developed bacterial leaf spot, indicating that the bacteriaremained viable and infectious during their association with the tardigrades and thatthe animals effectively transported the pathogens. The results as a whole suggest thatsome cases of

X. campestris

infection in nature may be produced by bacterial cellsshed from tardigrades.

S

UMMARY

Tardigrades were inoculated with

X. campestris

pv.

raphani

by soaking in a sus-pension of the bacteria or by contact with leaf spot lesions. Bacteria shed from thesetardigrades caused leaf spot disease in radish plants.

R

EFERENCES

C

ITED

A

GUINALDO

, A

NNA

M

ARIE

A., J

AMES

M. T

UBERVILLE

, L

AWERNCE

S. L

INFORD

, M

ARIA

C. R

IVERA

, J

AMES

R. G

AREY

, R

UDOLF

A. R

AFF

,

AND

J

AMES

A. L

AKE

. 1997. Evi-dence for a clade of nematodes, arthropods and other moulting animals. Nature387: 489-493.

B

ASHAN

, Y. 1985. Field dispersal of

Pseudomonas syringae

pv.

tomato

,

Xanthomonascampestris

pv.

vesicatoria

, and

Alternaria macrospora

by animals, people,birds, insects, mites, agricultural tools, aircraft, soil particles, and watersources. Canadian J. Bot. 64: 276-281.

K

INCHIN

, I. M. 1989.

Hypsibius anomalus

Ramazzotti (Tardigrada) from gutter sedi-ment. Microscopy 36: 240-244.

K

INCHIN

, I. M. 1993. An observation on the body cavity cells of

Ramazzottius

(Hyps-ibiidae, Eutardigrada). Quekett J. Microscopy 37: 52-55.

K

INCHIN

, I. M. 1994. The biology of tardigrades. Portland Press, London. 186 pp.K

RANTZ

, S. L., T. G. B

ENOIT

,

AND

C. W. B

EASLEY

. 1999. Phytopathogenic bacteria as-sociated with Tardigrada. Zool. Anz. 238: pp. 259-260.

200


83(2) June, 2000

FIRST RECORD OF

GONATOCERUS TRIGUTTATUS

(HYMENOPTERA: MYMARIDAE) FROM EGGS OF

HOMALODISCA COAGULATA

(HOMOPTERA: CICADELLIDAE) WITH NOTES ON THE DISTRIBUTION OF THE HOST

S

ERGUEI

V. T

RIAPITSYN

1

AND

P

HIL

A. P

HILLIPS

2

1

Department of Entomology, University of CaliforniaRiverside, California 92521, USA

2

University of California Cooperative Extension, Ventura CountyVentura, California 93003, USA

The glassy-winged sharpshooter,

Homalodisca coagulata

(Say), is primarily a south-eastern US species, but due to an apparent accidental introduction it has recently be-come a problem pest of crops and ornamentals in southern California (Blua et al. 1999);its spread into central and northern California is quite likely. Triapitsyn et al. (1998) re-ported results of the 1997 survey of

H. coagulata

egg parasitoids in Florida and Louisi-ana, where it is native, and also in southern California. In Monticello, Florida, its mainegg parasitoids were the mymarid wasp

Gonatocerus ashmeadi

Girault throughout theseason, and the trichogrammatid wasp

Zagella

sp. during July and August. This tri-chogrammatid was later found (S. V. Triapitsyn, unpublished data, and J. D. Pinto, per-sonal communication) to be different from the closely related species

Zagella floridae

Viggiani, described from a female and a male collected from jasmine in Fort Lauderdale,Florida (Viggiani 1985), thus representing a new, undescribed species.

Despite the fact that parasitization of

H. coagulata

eggs by

G. ashmeadi

may attimes reach 80% in Florida and Louisiana, and up to 100% in California (Phillips1998, Triapitsyn et al. 1998), it has become clear that the large numbers of

H. coagu-lata

in southern California are due in part to poor natural control of the first, earlyspring, generation of the sharpshooter.

In an effort to find a more effective natural enemy of the spring brood, we visitedTexas (Weslaco), and the state of Tamaulipas, Mexico, in late April 1999. Both stateshave never been surveyed for

H. coagulata

natural enemies, yet this leafhopper isknown to occur there (Young 1958, Turner & Pollard 1959). Young (1958, 1968) re-ported this species from Mexico, without providing further details about its distribu-tion. We examined a specimen of this species from Llera de Canales, which is in thetropical part of Tamaulipas. It is the most southern record of

H. coagulata

known inNorth America, based on the published locality data and according to our assessmentof

H. coagulata

distribution (Fig. 1). This leafhopper is abundant from eastern Texasto southern Georgia and northern Florida, but its populations in central Texas (westof Brown and Kerr Counties) and in central Florida decrease considerably (Turner &Pollard 1959). It appears to be rare also in northeastern Mexico, according to our ownobservations and the study of the leafhopper collection of the Universidad Autónomade Tamaulipas in Ciudad Victoria.

Homalodisca insolita

(Walker) and several

Oncometopia

species are the prevalentsharpshooters in southern Florida, Mexico, and central America (Young 1968), although

H. insolita

not long ago invaded the traditional areas of

H. coagulata

distribution(Turner & Pollard 1959). The sharpshooters

Oncometopia clarior

(Walker) and

O.

sp.near

nigricans

(Walker) occur on citrus in both Nuevo León and Tamaulipas, Mexico (see“Material Examined”). In Weslaco, Texas, we could not find any

H. coagulata

on citrus,but we saw last year’s egg masses, characteristic of this sharpshooter, with parasitoidexit holes on leaves of a mescal bean tree,

Sophora secundiflora

(Ortega) de Candolle.

Scientific Notes

201

In parks and citrus groves of central Tamaulipas, we were not able to find anyadult or nymph stages of

H. coagulata

despite the fact that we saw many last year’segg masses characteristic of the glassy-winged sharpshooter, all of them with evi-dence of parasitization. Only on one occasion did we collect

H. coagulata

adults andnymphs and also found its parasitized eggs both on citrus and peach, grown in theshade of larger trees in a private garden near Valle Hermoso, in Río Bravo del NorteValley (northern Tamaulipas).

In the laboratory of the Universidad Autónoma de Tamaulipas, the mymaridwasps

Gonatocerus triguttatus

Girault (taxonomic determination by S. V. Triapitsyn)emerged from

H. coagulata

egg masses collected in Valle Hermoso, thus providing uswith the first known host record for

G. triguttatus

. Interestingly, the type series of thisspecies was reared from an egg-mass of a leafhopper on orange in Trinidad (Girault1916).


was redescribed and illustrated by Huber (1988), whoalso indicated its distribution in Texas (Cameron, Hidalgo, and Val Verde Counties)and its presence in several states in Mexico, without, however, giving details of theMexican material; these are provided below under “Material Examined”. This speciesis probably widely distributed throughout Central America and parts of South Amer-ica (first author has seen unidentified specimens from Brazil, Costa Rica, Guatemala,Guyana, and Mexico that may be conspecific with

G. triguttatus

).We would expect that

G. triguttatus

would also attack other sharpshooter speciesin the genera

Homalodisca

and

Oncometopia

. Our observations in the citrus groves inTamaulipas indicate that the early spring populations of the sharpshooters

H. coagu-lata

,

O. clarior

, and

O.

sp. near

nigricans

are under good natural control by egg para-sitoids.

We believe that, subject to obtaining proper permits, prompt introduction of

G.triguttatus

into southern California from southeastern Texas or northeastern Mexicois warranted. Finding live

G. triguttatus

beyond

H. coagulata’s

range would be verydifficult because its host associations there are unknown. Other obvious candidates

Fig. 1. Homalodisca coagulata distribution map in North America.

202


83(2) June, 2000

for importation into California are

Gonatocerus fasciatus

Girault from Louisiana and

Zagella

sp. from Florida (Triapitsyn et al. 1998). If established, these species may en-hance the overall natural control of

H. coagulata

in southern California.Material Examined.


: MEXICO: Baja California Sur: LasBarracas (ca. 30 km E. of Santiago), 18-IV-1984, P. DeBach, 11 females; 10 km N. ofLa Paz, 28-X-1983, J. D. Pinto, 2 females. Nuevo León: El Carmen, 10-VII-1983, A.González H., 2 females; Hacienda El Canada, 12-VII-1983, A. González H., G. Gordh,M. A. Rodríguez P., 2 females; Linares, 23-X-1961, H. Suarez, 1 female (on citrus); SanJuan, 14-VII-1983, A. González H., F. Reyes V., 2 females [CNCI, Canadian NationalCollection of Insects, and UCRC, University of California at Riverside, det. J. Huber].Tamaulipas, nr. Valle Hermoso, 22-IV-1999, S. Triapitsyn & P. Phillips: 6 females (ex.

H. coagulata

eggs on peach); 18 females, 3 males (ex.

H. coagulata

eggs on citrus).


: MEXICO, Tamaulipas: Llera de Canales, 20-VIII-1994, D.Covarrubias, 1 female (on lemon); nr. Valle Hermoso, 22-IV-1999, S. Triapitsyn & P.Phillips, 1 female, 2 males.

Oncometopia clarior

: MEXICO, Tamaulipas: Ciudad Vic-toria, 28-VI-1997, Hernandez & Villegas, 1 female; Llera de Canales, 21-VI-1997,Mtz., Monrreal & Teran, 1 male (on orange).

Oncometopia

sp. nr.

nigricans

: MEXICO:Nuevo León, Montemorelos, 24-IV-1999, S. Triapitsyn & P. Phillips, 1 male (on citrus).Tamaulipas, nr. Santander Jiménez, 22-IV-1999, S. Triapitsyn & P. Phillips, 1 male(on

Hibiscus

sp.) [unless stated otherwise, all above specimens, including vouchers,deposited in the collection of Universidad Autónoma de Tamaulipas, Ciudad Victoria,Mexico].

We thank Svetlana N. Myartseva for her help with parasitoid rearing, EnriqueRuíz Cancino and Vladimir A. Trjapitzin for letting us use the facilities, arranging theloan of specimens, and providing various assistance (all Universidad Autónoma deTamaulipas, Ciudad Victoria, Mexico). Gordon Gordh (USDA-ARS, Weslaco, Texas) isacknowledged for facilitating our survey efforts in Texas. We are indebted to RaymondJ. Gill (California Department of Food and Agriculture, Sacramento) for the sharp-shooter identifications. Robert W. Brooks (Snow Collections, Natural History Mu-seum, University of Kansas, Lawrence) provided data about the geographicaldistribution of

H. coagulata

in Texas and John T. Huber (CNCI) pointed at additionalrecords of

G. triguttatus

in Mexico.

S

UMMARY

A survey of egg parasitoids of the glassy-winged sharpshooter,

Homalodisca coag-ulata

(Say), was conducted in Tamaulipas, Mexico and Texas, USA in April 1999. Themymarid


Girault was reared from egg masses of this leafhop-per on citrus and peach in northern Tamaulipas. This discovery is the first known hostrecord of

G. triguttatus

; its other, probable, host associations

are indicated. This wasp,whose geographical distribution also includes the eastern Rio Grande basin in Texasand the states of Baja California Sur and Nuevo León, Mexico, is identified as a po-tential biological control agent for introduction into California against

H. coagulata

.

R

EFERENCES

C

ITED

B

LUA

, M. J., P. A. P

HILLIPS

,

AND

R. A. R

EDAK

. 1999. A new sharpshooter threatensboth crops and ornamentals. California Agric. 53(2): 22-25.

G

IRAULT

, A. A. 1916. New miscellaneous chalcidoid Hymenoptera with notes on de-scribed species. Ann. Entomol. Soc. America 9: 291-308.

H

UBER

, J. T. 1988. The species groups of

Gonatocerus

Nees in North America with arevision of the

sulphuripes

and

ater

groups (Hymenoptera: Mymaridae). Mem.Entomol. Soc. Canada 141: 1-109.

Scientific Notes

203

P

HILLIPS

, P. A. 1998. The glassy-winged sharpshooter: a potential threat to Californiacitrus. Citrograph 83(12): 10-12.

T

RIAPITSYN

, S. V., R. F. M

IZELL

, III, J. L. B

OSSART

,

AND

C. E. C

ARLTON

. 1998. Egg par-asitoids of


(Homoptera: Cicadellidae). Florida Ento-mol. 8 (2): 241-243.

T

URNER

, W. F.,

AND

H. N. P

OLLARD

. 1959. Life histories and behavior of five insectvectors of phony peach disease. Tech. Bull. United States Dep. Agric. 1188, 28pp.

V

IGGIANI

, G. 1985. A new species of

Zagella

(Hym. Trichogrammatidae) from Florida.Boll. Lab. Entomol. Agr. “Filippo Silvestri”, Portici 42: 15-17.

Y

OUNG

, D. A. 1958. A synopsis of the species of

Homalodisca

in the United States.Bull. Brooklyn Entomol. Soc. 53(1): 7-13.

Y

OUNG

, D. A. 1968. Taxonomic study of the Cicadellinae (Homoptera, Cicadellidae).

♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦♦

Part 1. Proconiini. United States Nat. Mus. Tech. Bull. 261, 287 pp.

SUITABILITY OF THREE LEGUMES FOR DEVELOPMENT OF

TETRANYCHUS OGMOPHALLOS

(ACARI: TETRANYCHIDAE)

O. B

ONATO

1, 2

, P. L. S

ANTAROSA

2

, G. R

IBEIRO

2

AND

F. L

UCCHINI

2

1

IRD (ex ORSTOM), Entomologie Agricole, B.P. 5045, 34032 Montpellier, France

2

Embrapa—Meio Ambiente, C.P. 69, 13820-000 Jaguariúna/SP, Brazil

Tetranychus ogmophallos

Ferreira & Flechtmann is a new tetranychid found onpinto peanut,

Arachis pintoi

(Krap. & Greg.) (Fabaceae), in an experimental field ofthe Centro Nacional de Pesquisa Agropecuária do Cerrado

(EMBRAPA) in Planal-tina—DF, Brazil (Ferreira & Flechtmann 1997). Pinto peanut is a Brazilian legumemainly used as a nitrogen fixator in pasture fields or as green manure in tropical cropssuch as coffee, banana, and oil-palm (Grof 1984, De la Cruz et al. 1994, Suarez-Vasquez et al. 1992). Because of its agronomic characteristics pinto peanut

is cur-rently exported from Brazil to other countries of Latin America and to Australia(Asakawa & Ramirez 1989, Vilarreal & Chavez 1991, Cook et al. 1990). Thus, throughexports of pinto peanut

T. ogmophallos

could be accidentally introduced into new ar-eas, and is a quarantine issue.

The objective of this work was to compare the suitability of three legumes of eco-nomic importance, common bean,

Phaseolus vulgaris

L., peanut,

A. hypogeae

L., andsoybean,

Glycine max

Merrill, as host plants of

T. ogmophallos.

Studies were con-ducted under laboratory conditions at 26

±

0.5

°

C, 75

±

10% RH and a photoperiod of13:11 (L:D). Experiments were carried out using the progeny of about 50 females fromthe quarantine laboratory of CENARGEN, EMBRAPA, Brasília. Mites were reared onbean leaves

in the same environmental conditions as above. To study survival rate,developmental time, and sex-ratio, 4-5 females were placed on the lower surface of aleaf disk (4 cm

2

) maintained on water-soaked cotton. After 1 h, females and excesseggs were eliminated to obtain one egg per disk; 70 disks were kept for each treat-ment. The disks were monitored three times a day (at 7 a.m., 1 p.m., and 7 p.m.) untiladult emergence. For oviposition study, one female teliochrysalis (last pre-imago in-star) and two males were placed on a leaf disk and the males were removed 48 h after

204


83(2) June, 2000

the female had emerged. The number of eggs laid per female was monitored daily. Thedisks were changed every 4 days. Thirty females were followed per legume. Demo-graphic parameters (net reproductive rate, R

0

, generation time, G, and intrinsic rateof natural increase, r

m

) were determined using a program developed by Hulting et al.(1990). Biological parameters (survivorship, developmental time, fecundity, and lon-gevity) were compared between treatments using a one-way ANOVA (LEAS 1989). IfANOVA revealed significant differences, means were compared using the Scheffémethod. R

0

and r

m

were compared using the SNK test (Hulting et al. 1990).Development from egg to adult occurred on all three plant species tested. Develop-

mental time (Table 1) recorded on peanut was significantly different than those re-corded on common bean and on soybean (F = 108.16; df = 2-144; P < 0.0001). On eachplant species, 85

±

5% of eggs reached adult stage, with no significant differences be-tween treatments. The sex-ratio was 80% female for all 3 legumes.

Longevity in days for adult females differed significantly among host plant(Table 1) (F = 21.7; df = 2-79; p < 0.05). The highest total fecundity was obtained oncommon bean (F = 24.9; df = 2-79; p < 0.05), whereas no significant difference wasfound between peanut and soybean.

The R

0

calculated on common bean was significantly higher than R

0

obtained onsoybean (q = 3.6, df = 3, 19; p < 0.05) and on peanut (q = 4.1, df = 3, 19; p < 0.05). Nosignificant difference was found between the latter two legumes (q = 0.5, df = 3, 19; p> 0.05) (Table 1).

The r

m

values differed significantly between host plants. The highest r

m

was ob-tained on common bean (q = 8 with soybean and 7.3 with peanut, df = 3, 19; p < 0.05)and the lowest on peanut

(q = 4.1 with soybean, df = 3, 19; p < 0.05) (Table 1)

.

Results of the present study indicate that

T. ogmophallos

is not only able to developon common bean, soybean, and peanut, but displayed high rates of increase whenreared on these three plants. Values of the biological and demographic parameterswere in the range observed for other

Tetranychus

spp. under similar environmentalconditions (Gutierrez 1976, Carey & Bradley 1982, Tsai et al. 1989, Rai et al. 1995).

Because tetranychids are polyphagous and

T. ogmophallos

showed high rates of in-crease when reared on these three different plants, it seems reasonable to speculatethat

T. ogmophallos

also could develop on a wide range of hosts. Thus, further inves-tigations are needed in order to precisely characterize this range.

T

ABLE

1. B

IOLOGICAL

AND

DEMOGRAPHIC

PARAMETERS

OF

T

ETRANYCHUS

OGMOPHAL-LOS

ON

3

PLANT

SPECIES

AT

26

°

C, 75% R

H

,

AND

13:11

L

:

D

.

ParametersArachis

hypogeae Glycine maxPhaseolusvulgaris

Developmental time

1

(egg to adult)14.2

±

0.2 11.9

±

0.8 11.7

±

0.4

Total fecundity 60.0

±

2.9 63.9

±

5.7 104.3

±

7.8Longevity of females

1

16.5

±

0.8 15.4

±

1.2 25.3

±

1.1Mean generation time

1

(G) 20.1 18.9 20.7Net reproductive rate (R

0

) 41.05

±

2.0 45.7

±

4.1 75.9

±

4.9Intrinsic rate of increase (r

m

) 0.190

±

0.003 0.215

±

0.001 0.232

±

0.004

Means are followed by

±

SE.

1

Days.

Scientific Notes

205

S

UMMARY

The suitability of common bean,

Phaseolus vulgaris

, soybean,

Glycine max,

andpeanut,

Arachis hypogeae

, as food substrates for the mite

Tetranychus ogmophallos

was evaluated. The mite performed better on common bean (r

m

= 0.232) although it de-veloped and reproduced well on soybean (r

m

= 0.215) and peanut

(r

m

= 0.190).

R

EFERENCES

C

ITED

A

SAKAWA

, N. M.,

AND

R. R

AMIREZ

. 1989. Inoculation and planting methodology for

Arachis pintoi

. Pasturas Tropicales 11: 24-26.C

AREY

, J. R.,

AND

J. W. B

RADLEY

. 1982. Developmental rates, vital schedules, sex-ra-tios and life tables for

Tetranychus urticae

,

T. turkestani

and

T. pacificus

(Ac-arina: Tetranychidae) on cotton. Acarologia 23: 333-345.

C

OOK

, B. G., R. J. W

ILLIAMS

,

AND

G. P. M. W

ILSON

. 1990.

Arachis pintoi

Krap. et Greg.nom. nud. (pinto peanut) cv. Amarillo. Australian J. Exp. Agric. 30: 445-446.

D

E

LA

C

RUZ

, R., S. S

UAREZ

,

AND

J. E. F

ERGUSON

. 1994. The contribution of

Arachispintoi

as a ground cover in some farming systems of Tropical America, pp. 102-108.

In

P. C. Kerridge, and B. Hardy [eds.] Biology and agronomy of forage

Ara-chis

. Centro Intern. Agric. Trop. Pub. 240, Cali, Colombia.F

ERREIRA

, D. N. M.,

AND

C. H. W. F

LECHTMANN

. 1997. Two new phytophagous mites(Acari: Tetranychidae, Eriophyidae) from

Arachis pintoi

from Brazil. Syst.Appl. Acarol. 2: 181-188.

G

ROF

, B. 1984. Forage attributes of the perennial groundnut Arachis pintoi in a trop-ical savanna environment in Colombia.

In

Proc. International Grassland Con-gress, 15., Kyoto, 1985. Centro Intern. Agric. Trop. Cali, Colombia, p. 168-170.

G

UTIERREZ

, J. 1976. Etude biologique et écologique de

Tetranychus neocaledonicus

André (Acarien: Tetranychidae). Cahiers Orstom. Paris.H

ULTING

, F. L., D. B. O

RR

,

AND

J. J. O

BRYCKI

. 1990. A computer program for calcula-tion and statistical comparison of intrinsic rates of increase and associated lifetable parameters. Florida Entomol. 73: 601-612.

LEAS. Logiciels d’Enseignement et d’Analyses Statistiques. 1989. Chaire de Mathé-matiques. Unité de Biométrie de l’Ecole Nationale Supérieure Agronomique deMontpellier France.

R

AI

, A. B., A. S. S

EJALIA

, C. B. P

ATEL

,

AND

S. K

UMAR

. 1995. The rate of natural in-crease of red spider mite when reared on okra. Gujarat Agricul. Univ. J. 21:130-136.

S

UAREZ-VASQUEZ, S., M. WOOD, AND S. NORTCLIFF. 1992. Crecimiento y fijacíon de ni-trógeno por Arachis pintoi establecido con Brachiara decumbens. Cenicafe 43:14-21.

TSAI, S. M., K. S. KUNG, AND C. I. SHIH. 1989. The effect of temperature on life historyand population parameters of Kanzawa spider mite, Tetranychus kanzawaiKishida (Acarina: Tetranychidae) on tea. Plant Protect. Bull. Taiwan 31: 119-130.

VILARREAL, M., AND O. CHAVEZ. 1991. Adaptación y producción de gramíneas y legu-minosas forrajeras en San Carlos, Costa Rica. Pasturas Tropicales 13: 31-38.

206


83(2) June, 2000

BOOK REVIEW

M

ATTHEWS

, M. 1999. Heliothine Moths of Australia: A Guide to Pest Bollwormsand Related Noctuid Groups. CSIRO Publishing, Collingwood, Vict., Australia. x +320 p. (23 color pl.) (17

´

25 cm), plus CD-ROM. ISBN 0-643-06305-6. Hardback. $90.

Although written from a project to identify the bollworms and related pest speciesof this important economic group of moths, Matthews has provided a careful taxo-nomic revision of the group that all students of the Noctuidae will find useful. Thebook treats 38 species from Australia, with 18 new synonymies and 8 new species, in5 genera. The book has excellent illustrations, including 23 color plates of all adultsand known larvae, plus 460 other figures (halftone photographs or SEM micrographs)illustrating genitalia and other morphological characters of adults and immatures.The CD-ROM included with the book includes a complete listing of all label data fromthe over 14,800 specimens examined for the study and taxonomic lists. The illustra-tions are all clear and finely printed. The color plates are very sharp and true to thecolor tone of the moths and larvae. The book is finely printed and well bound.

Following an introduction on the methodology used, there are chapters on the eco-nomic importance of Heliothinae in Australia, heliothine biology, systematics, mor-phology, phylogeny, and identification keys to genera and species. Thereafter, theauthor presents the species in the format of a traditional taxonomic revision, with di-agnoses and descriptions of all taxa. Each species is discussed in detail, with new de-scriptions, whether of known or new species, followed by notes on bionomics. Eachspecies has a range map included for Australia and Tasmania. After the taxonomicsection, there follows a checklist of all species for Australia, and then a chapter givingdetailed nomenclatural notes for all scientific names, including genera and all syn-onyms. Prior to the monochrome figures of genitalia, there is a short section withnotes on the slides and specimens used for the figures.

The color plates include 2 plates showing some Australian habitats of heliothines,followed by 2 useful color plates showing adults greatly enlarged and overprintedwith feature names for maculation and wing venation terms, plus details of the legs.There are 8 color plates showing the Australian species from museum specimens(about life size). There then is one color plate with enlarged views of the wing featuresthat allow identification of two closely related species:

Helicoverpa armigera

and

H.punctigera

. Finally, there are 10 color plates illustrating adults in nature, larvae andpupae of many of the species.

The author gives a detailed treatment for each species for Australia, but ratherless detailed discussion of the genera. This is partly due to his earlier work on theworld genera of Heliothinae (1991.

Classification of the Heliothinae

), where he al-ready went into detail as to the generic limits for the subfamily. He gives further evi-dence, particularly getting into molecular data, of the complexity of the heliothinegenera of Australia. Since Hardwick split

Helicoverpa

from the well-known genus

He-liothis

in his 1965 monograph on North American heliothines, there has been contin-ued argument from specialists as to whether

Helicoverpa

should be a subgenus or afull genus. Part of these varying opinions were based no doubt on the similarities ofthe species to be found in North America. Matthews, in covering all the many Austra-lian species, clearly shows in this new study that the variation of the group is muchmore complex outside of North America, thus further supporting Hardwick’s morepreliminary work in splitting

Heliothis

. Thus, the Australian fauna in particular dem-onstrates that the old concept of

Heliothis

is too broad to include so many differentspecies groups. Although many of the Australian species have characters that are

Book Review

207

very similar,

Helicoverpa

in particular is distinct enough to be a genus on its own, andthe other Australian groups vary so much from

Heliothis

that one concludes with Mat-thews that they are best treated as separate genera: thus,

Adisura, Heliothis, Helio-cheilus, Australothis

, and

Helicoverpa

.Economic entomologists using this book will be able to accurately identify all Aus-

tralian heliothine adults and larvae: most species are distinct enough that the colorplates of the adults will suffice for identification, and only a few may require genitalicdissection for species confirmation. The widespread Old World pest,

Helicoverpa ar-migera

, occurs in Australia, thus the book is of use in other regions as well, particularlysince it is so carefully prepared and presented. The book is particularly important forthose involved in checking ports of entry for exotic pests, since anything from Australiacan be checked using this book as an identification guide for this group of moths.

Being virtually the finest and best illustrated revision of this group of moths forany region of the world, all researchers on Noctuidae will need this book on their ref-erence shelf, and likewise for economic entomologists.

J. B. HeppnerFlorida State Collectionof Arthropods, FDACS, DPIP. O. Box 147100Gainesville, Florida 32614

morales-ramos et al.: two populations of c. grandis 137...

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