Systematic studies of New World Encarsia species and a survey of the parasitoids of Bemisia tabaci in Florida, the Carri...

MISSING IMAGE

Material Information

Title:
Systematic studies of New World Encarsia species and a survey of the parasitoids of Bemisia tabaci in Florida, the Carribbean and Latin America
Physical Description:
ix 284 leaves : ill. ; 29 cm.
Language:
English
Creator:
Evans, Gregory Allyn, 1956-
Publication Date:

Subjects

Subjects / Keywords:
Entomology and Nematology thesis Ph. D
Dissertations, Academic -- Entomology and Nematology -- UF
Genre:
bibliography   ( marcgt )
non-fiction   ( marcgt )

Notes

Thesis:
Thesis (Ph. D.)--University of Florida, 1993.
Bibliography:
Includes bibliographical references (leaves 265-283).
Statement of Responsibility:
by Gregory Allyn Evans.
General Note:
Typescript.
General Note:
Vita.

Record Information

Source Institution:
University of Florida
Rights Management:
All applicable rights reserved by the source institution and holding location.
Resource Identifier:
aleph - 001957683
oclc - 31323148
notis - AKD4296
sobekcm - AA00003657_00001
System ID:
AA00003657:00001

Full Text







SYSTEMATIC STUDIES OF NEW WORLD ENCARSIA SPECIES
AND A SURVEY OF THE PARASITOIDS OF BEMISIA TABACI
IN FLORIDA, THE CARIBBEAN AND LATIN AMERICA










By

GREGORY ALLYN EVANS


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


1993











ACKNOWLEDGEMENTS


I thank God for the opportunity to have been a part of this cooperative effort

to bring about the control of the sweet potato whitefly and that I could discover,

once again, the grandeur of His creation through these very minute organisms. I

thank my wife Blanca, and my children for their support throughout the study.

There are several people who made major contributions to the success and

completion of this document; I am thankful and indebted to Dr. Fred D. Bennett,

who served as the chairman of my committee through most of the duration of the

study and was a major collector of the parasitoids reared from Florida and other

countries, to the chairman of my committee Dr. David J. Schuster, and to my

committee members, Dr. Harvey L. Cromroy, Dr. Virenda Gupta, Dr. Lance

Osborne and Dr. Jonathan Reiskind; I thank them for their guidance, support and

assistance, and for their review of this manuscript.

Parasitoid identifications were made by: L. Massner, M. Hayat, A. Polaszek,

M. Rose, G. Viggiani and J. Woolley. A. Hamon identified the majority of the

aleyrodid and scale insect hosts. C. Arnaud identified many of the host plants in

the study. Collections of parasitoids were made by: B. Alvarado, T. Bellows, F. D.

Bennett, H. J. Browning, R. Caballero, R. D. Cave, Y. Chen, J. H. Frank, K. A.

Hoelmer, R. C. Lambe, G. Leibee, L. Nong, R. Nguyen, L. S. Osborne, J. Pena,






T. Perring, D. J. Schuster, P. Stansly, F. Stonaker, S. Toppertzhoffen and E.

Vasquez. Specimens were processed and mounted on slides by Y. Chen, F.

Stonaker and E. Vazquez.

Brett Linbo-terharr assisted in the statistical analysis of the whitefly and parasitoid

population studies.

The encouragement and support of the co-investigators D. J. Schuster and

J. F. Price, are greatly appreciated. Financial support for the investigation was

provided under the CSRS Special Grant 89-34135-4581 'Biological Factors

Affecting the Abundance of the Sweetpotato Whitefly in the Caribbean Including

Florida'.


iii











TABLE OF CONTENTS


ACKNOWLEDGEMENTS ....................................

ABSTRACT .............................................


CHAPTER


1. INTRODUCTION .............................

2. BIOLOGY AND POPULATION STUDIES ...........

2.1. Whiteflies
2.1.1 Geographic distribution ..............
2.1.2. Systematics and evolutionary relationships
2.1.3. Life Stages ......................
2.1.4. Sex ratio ........................
2.1.5. Developmental rate ................
2.1.6. Fecundity ........................
2.1.7. Longevity ........................
2.1.8. M ortality .........................
2.1.9. Host plant specificity, suitability and
preference .......................
2.1.10. M igration ........................
2.1.11. Population dynamics ...............


2.2. Parasitoi(
2.2.1.
2.2.2.


Is
Life cycle and development ......
Longevity ...................


2.2.3. Host feeding


2.2.4.
2.2.5.
2.2.6.


Host specificity, suitability and preference
Adelphoparasitism ................
Population dynamics ...............


3. HISTORICAL REVIEW AND HIGHER CLASSIFICATION ...

3.1. Introduction ...................................


. .


11111






3.2. New World Encarsia studies .......................
3.3. Generic concepts ................................

4. MATERIALS AND METHODS ............................


4.1.
4.2.
4.3.


Natural enemy survey .............................
Systematic studies ...............................
Morphology and explanation of terms and measurements ...


5. KEYS TO GENERA, SPECIES GROUPS AND SPECIES ........


5.1.
5.2.
5.3.


Introduction ....................................
Taxonomic key to genera ..........................
Biological characteristics of the genera ................
5.3.1. Family Platygasteridae Genus Amitus ............
5.3.2. Family Eulophidae Genus Euderomphale .........
5.3.3. Family Encyrtidae Genus Metaphycus ...........
5.3.4. Family Signiphoridae Genus Sianiphora ..........


5.3.5. Family Aphelinidae ........................
5.3.5.1. Genus Azotus ..................
5.3.5.2. Genus Cales ...................
5.3.5.3. Genus Dirphys ..................
5.3.5.4. Genus Encarsiella ...............
5.3.5.5. Genus Eretmocerus ..............
5.4. Key to species and species group ..................

6. ENCARSIA SPECIES AND SPECIES GROUPS .............


55
55
59
59
59
60
60


.60
.. 60
61
61
61
.62
.62


6.1. Species groups ..................
6.2. Species groups and species diagnosis an
6.2.1. Cubensis Group .............
6.2.2. Formosa Group .............
6.2.3. Parvella Group ..............
6.2.4. Strenua Group ..............
6.2.4.1. Strenua subgroup ..
6.2.4.2. Quercicola subgroup
6.2.4.3. Bella subgroup .....
6.2.5. Opulenta Group .............
6.2.6. Inaron Group ...............
6.2.7. Lutea Group ................
6.2.8. Lahorensis Group ............
6.2.9. Porteri Group ...............
6.2.10. Plaumanni Group ............
6.2.11. Aurantii Group ..............


d discussion .....


101
105


. .106
..112
..129
. 138
. 138
. 145
. 152
. 152
..160
. 164
. 165
. 168
. 172
. 173





6.2.12. Citrina Group .............................. 175
6.2.13. Inquirenda Group ......................... 176
6.2.14. Perflava Group ............................. 176
6.2.15. Elegans Group ........................... 176
6.2.16. Tricolor Group ........................... 176
6.2.17. Singularis Group ........................... 177
6.3. Encarsiella alboscutellaris .... ... .... ............. 177
6.4. Summary ........................... ... ...... 179

7. CLADISTICAL ANALYSIS ............................... 180

7.1. Introduction ................ ................. 180
7.2. Methods .................. ........ ........... 181
7.2.1. Analysis .......... ...................... 181
7.2.2. Terminal taxa ............... ............. 181
7.2.3. Characters and character matrix ............... 183

8. SURVEY OF NATURAL ENEMIES ........................ 197
8.1. Introduction .................................... 197
8.2. Methods ...................................... 198
8.3. Results ................. .......... ......... 199
8.3.1. SPW F Parasitoids .......................... 199
8.3.2. Alternate hosts of SPWF parasitoids ............. 204
8.3.3. Population dynamics of SPWF and parasitoids ...... 205
8.3.3.1.Gainesville .......................... 206
8.3.3.1.1. SPWF population .... ..... 206
8.3.3.1.2. Parasitoid population dynamics ... 208
8.3.3.1.3. Parasitoid complex ............ 209
8.3.3.1.4. Parasitoid sex ratio ............ 210
8.3.3.1.5. Host-plant relationships ........ 211
8.3.3.1.5.1. SPWF seasonality ....... 211
8.3.3.1.5.2. Parasitoid seasonality ..... 212
8.3.3.1.5.3. Parasitoid complex ....... 212
8.3.3.1.6. Host plant examples ........... 215
8.3.3.1.6.1. Brassica oleracea ...... 215
8.3.3.1.6.2. Chamaesvce hyssopifolia ..216
8.3.4. Other Florida sites ........................... 217
8.3.4.1. Bradenton .......................217
8.3.4.2. Homestead .... .... ....... ......218
8.3.4.3. Immokalee .......................219
8.3.5. Discussion. ..... ..... ..... .... ............ 219






9. DISCUSSION ...................................... 241
9.1. The role of systematics in biological control .............. 241
9.2. Biological control of the SPWF ....................... 244
9.2.1. Parasitoid requirements .................... ..245
9.2.1.1. Affinity to parasitize the SPWF ............ 245
9.2.1.2. Synchronization of life cycles ............. 245
9.2.1.3. Asexual versus sexual parasitoids ......... 246
9.2.1.4. Parasitoid fecundity and developmental rate .. 249
9.2.2. Parasitoid utilization ..............................250
9.2.2.1. Current status ............................. 250
9.2.2.2. Parasitoid conservation ...................... 251
9.2.2.3. Augmentation and manipulation ................ 252
9.2.2.3. Integrated pest management .................. 253

APPENDICES .......................................... 256

A. HOST-PARASITOID LIST ..........................256

B. HOST PLANT-ALEYRODID LIST ..................... 260

C. MEASUREMENTS AND STATISTICS FOR NEW SPECIES 264

REFERENCE LIST ....................................... 265

BIOGRAPHICAL SKETCH .................................. 284


vii










Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy

SYSTEMATIC STUDIES OF NEW WORLD ENCARSIA SPECIES
AND A SURVEY OF THE PARASITOIDS OF BEMISIA TABACI
IN FLORIDA, THE CARIBBEAN AND LATIN AMERICA

By

GREGORY ALLYN EVANS

DECEMBER 1993


Chairperson: Dr. David J. Schuster
Major Department: Entomology and Nematology

Fifty-four species of the genus Encarsia, including the description of three

new species, Encarsia lanceolata spec. nov., E. pseudocitrella, and E. polaszeki

spec. nov.; 13 new distribution records; and 35 new host records are reported from

the New World. The synonomy of E. hispida De Santis with E. meritoria Gahan is

re-established, E. protransvena Viggiani is placed in synonomy with E. armata

(Silvestri) and Encarsia alboscutellaris De Santis is transferred to the genus

Encarsiella. Seventeen worldwide Encarsia species groups are recognized, 12

occur in the New World. Two new species groups, the Plaumanni and Porteri

Groups, and 3 new subgroups of the Strenua Group, Strenua, Quercicola and

Bella, are proposed.


viii





Taxonomic keys and figures are given to the genera of parasitoids that

attack aleyrodids, worldwide Encarsia species groups, and Encarsia species that

parasitize aleyrodids in the New World. Each species group is diagnosed and

individual Encarsia species that parasitize whiteflies in the New World are

diagnosed and redescribed.

Results of a cladistical analysis of 15 Encarsia species, 2 Encarsiella

species and Pteroptrix chinensis based on 40 morphological and 1 biological

characters indicated a close relationship between the genus Encarsia and the

genus Pteroptrix, and supported the transfer of Encarsia alboscutellaris De Santis

to the genus Encarsiella.

Parasitoids were reared from the sweet potato whitefly (SPWF), Bemisia

tabaci, and 37 other whitefly species in a survey conducted in Florida, the

Caribbean and Latin America between 1988 and 1992. Nineteen and 12 parasitoid

species were reared from the SPWF in the New World and Florida, respectively,

including 3 new Encarsia species. Seasonal and geographic distribution and host

records for each parasitoid species, and the parasitoid complex of the SPWF on

various host plants and locations, are given.










CHAPTER 1


INTRODUCTION



The following study was initiated in response to the need to gather

information on the identity and biology of native and exotic parasitoids of Bemisia

tabaci (Gennadius). B. tabaci, also known as the tobacco, cotton or sweet potato

whitefly, referred to from here on as SPWF, which has for many years been a

serious pest in the Caribbean and Central American countries. Although it was

recorded from Florida as early as 1894 (Quaintance, 1900), it was seldom a

serious pest in the United States until outbreaks occurred in 1981 in the irrigated

desert regions of the southwestern United States, causing an estimated $100

million in agricultural losses (Duffus & Flock, 1982). Damages caused by the

SPWF include reduction of crop yields by direct feeding injury and loss of host

plant vigor; product contamination by honeydew and consequent production of

sooty mold fungus; induction of growth disorders; and transmission of at least 19

different viruses (Osborne, 1992). In 1986, the Florida greenhouse foliage industry

experienced severe population outbreaks of the SPWF on poinsettia plants (Price

1987) that resulted in statewide losses in excess of $2 million (Osborne, 1992).






2

Squash silverleaf and tomato irregular ripening were observed as widespread

outbreaks of SPWF occurred on Florida field vegetable crops in 1987 and 1988

(Maynard & Cantliffe, 1989). These new disorders were attributed to the

association of SPWF on these crops (Schuster et al. 1990; Yokomi et al. 1990).

Losses due to irregular ripening, gemini virus, and increased control costs on

tomatoes alone in Florida were estimated at $125.3 million (Schuster, 1992).

Ornamental growers in Florida were forced to abandon some crops and increase

pesticide applications on many others; tomato growers sprayed as often as twice

a week for SPWF. Peanut yields were reduced in some fields to only one-third of

that of previous years due to heavy SPWF infestations (Leidner, 1991). Annual

losses to all Florida agriculture were conservatively placed at $141 million

(Schuster, 1992). Large populations of SPWF were reported on cotton grown in

the Texas Rio Grande Valley in 1991 causing an estimated loss of $80 million

(Faust, 1992). Epidemic outbreaks were reported on cotton, melons, lettuce and

other crops in the Imperial Valley, California and in Arizona (Gill, 1992). Large

populations of SPWF occurred on the sesame crop in Venezuela and on tomatoes

in the Dominican Republic and Honduras and other countries resulting in heavy

crop losses (F.D. Bennett, personal communication). The total losses caused by

the SPWF to U.S. agriculture in 1991 was estimated at least $500,000,000

(Perring et al. 1993).

The rapid rise of the SPWF to key pest status has been attributed to the

widespread use of pesticides and changes in crop culture in the early 1970's. This






3
led to the SPWF's accelerating development of physiological resistance to

pesticides and the disruption of its natural enemy fauna (Byrne et al. 1990).

Resistance of the SPWF to various pesticides has been documented by Prabhaker

et al. (1985) and others.

Perring et al. (1992) suggested that the sudden change in the pest status

of the SPWF was due to the invasion of a new, but morphologically similar species

to which the common name, "the silverleaf whitefly" was given. Populations of the

silverleaff whitefly" have a wider host range, varying ability to transmit viruses, and

different biochemical markers, and are reproductively incompatible with laboratory

colonies of the historically "indigenous" (pre-1986), cotton-derived SPWF

populations.

In 1987, Florida biological control and plant protection specialists, virologists

and pathologists, growers and grower groups, together with scientists from

Arizona, California and Texas, initiated a multi-faceted Bemisia research program.

The program focused on developing alternative control and management measures

to suppress SPWF populations to acceptable economic levels. Conservation and

introduction of biological control agents were key elements of the overall integrated

pest management strategy. Parasitoids have played an important role in the control

of several whitefly pest species (Table 1.1). A statewide survey of the natural

enemies of the SPWF was initiated in 1988 to gather information on the parasitoid

complex of the SPWF in Florida and the humid neotropics, and to identify

promising natural enemy species from the neotropics (and other areas of the







Table 1.1. Parasitoids used for the biological control of whitefly pests species.

Whitefly species Parasitoid species Reference

Aleurocanthus woglumi Amitus hesperidum Flanders 1969.
Encarsia opulenta
E. merceti
E. clypealis
E. smith
Eretmocerus serious

Aleurothrixus floccosus Cales noacki Rose and Woolley, 1984

Dialeurodes citri Encarsia lahorensis Nguyen and Sailer, 1979

Parabemisia myricae Encarsia transvena Rose et al. 1981

Eretmocerus debachi Rose and Rosen, 1992

Siphonius phillvreae Encarsia partenopea Gould et al. 1992

Tetraleurodes sp. Cales noacki Rose and Woolley, 1984

Trialeurodes vaporariorum Encarsia formosa Osborne, 1981


world) for eventual introduction into Florida and the southern United States. The

objectives of the survey were to identify the species of parasitoids commonly found

parasitizing SPWF; measure the ratio between emergent SPWF and parasitoids

as a comparative indicator of parasitism; identify important SPWF wild host plant

reservoirs; identify changes in the parasitoid complex composition attributable to

seasonal, geographic and host plant variation; determine if SPWF and its

parasitoids can overwinter in northern Florida. In addition, parasitoids were reared

from several other whitefly species in order to determine the host range for each

of the parasitoid species reared from SPWF (Bennett et al. 1990).


]






5
Previous surveys of parasitoids of SPWF have been carried out in Pakistan

by Dr. A. I. Mohyuddin of the International (formerly Commonwealth) Institute of

Biological Control (IIBC), and in Africa and the Middle East by Prof. D. Gerling

(University of Tel Aviv, Israel). Results of these have been largely published

(Lopez-Avila 1986; Gerling, 1985; Rivnay and Gerling, 1987) but contain some

errors. Thanks to the accumulation of a large amount of reliably reared material

specimens from SPWF by F. D. Bennett, as well as the opportunity to examine the

rich material from the IIBC survey, many of these errors were corrected by

Polaszek et al. (1992).

The present study is divided in two parts. The first part is a taxonomic

review of the species of Encarsia described from the New World along with the

description of three New World species. Also included are biological information,

references and a key for the following genera of primary and secondary whitefly

parasitoids: Azotus, Cales, Dirphys, Eretmocerus, Encarsiella (Aphelinidae); Amitus

(Platygasteridae); Euderomphale (Eulophidae); Metaphycus (Encyrtidae); and

Signiphora (Signiphoridae). The key that is presented for the Encarsia species of

the New World includes the species that were described from areas outside the

survey area but whose distribution includes the New World. Host, distribution and

biological information for these species are also presented.

The second part of this study examines the population dynamics of the

SPWF and its parasitoid complex on different host plants and in different localities

in Florida. In addition, data are presented on the distribution and relative

abundance of parasitoids reared from SPWF in other countries.






6

Several obstacles had to be overcome, particularly during the earlier phases

of the survey. Taxonomically, the genus is not very well understood, especially in

the New World. Systematic knowledge of the genus in the United States has

remained dormant for more than 50 years, with the exception of work done on the

biology and utilization of a limited number of Encarsia species. Most of these

species were introduced for the biological control of specific whitefly and scale

pests. Initially, it was necessary to seek the assistance of Dr. Andrew Polaszek

(Natural History Museum, London), one of the few experts on the taxonomy of this

genus in the world. The genus Encarsia is considered to be a taxonomically

difficult genus for several reasons. The minute size of the organisms, which rarely

exceed one millimeter in length, usually necessitates that they be cleared of their

body content and mounted on microscope slides in order to observe the characters

that are used to distinguish them from other species. Only a small portion of the

New World Encarsia species are probably known at present. Of the more than 170

described species worldwide (Hayat 1989), only thirty-six species (currently valid)

have been described from the New World (Table 1.2). Fifteen exotic species have

been reported in the New World. Most of these species are of Oriental origin that

were introduced as biological control agents for specific whitefly and scale insect

pests (Table 1.3). This survey adds 3 new species and 3 new distribution records

for exotic species to increase the number of New World species to fifty-four.

Distribution and host range information for the New World Encarsia species is very







Table 1.2. Encarsia species described from the New World.


Species Author Year Country Host


alboscutellaris
americana
aurantii
basicincta
bella
berlesei
brasiliensis
brunnea
catherinae
ciliata
citrella
citrina
coquilletti
cubensis
desantisi
ectophaga
elongata
formosa
gallardoi
guadaloupae
haitiensis
lopezi
luteola
lycopersici
meritoria
nigricephala
peltata
pergandiella
perniciosi
peruviana
plaumanni
porter
portoricensis
protransvena
quaintancei
quercicola
townsendi
variegata


De Santis
DeB.& Rose
Howard
Gahan
Gahan
Howard
Hempel
Howard
Dozier
Gahan
Howard
Howard
Howard
Gahan
Viggiani
Silvestri
Dozier
Gahan
Marelli
Viggiani
Dozier
Blanchard
Howard
De Santis
Gahan
Dozier
Cockerell
Howard
Tower
Rust
Viggiani
Mercet
Howard
Viggiani
Howard
Howard
Howard
Howard


1979
1981
1894
1927
1927
1906
1904
1908
1933
1927
1908
1891
1908
1931
1981
1938
1937
1924
1933
1985
1932
1940
1895
1957
1927
1937
1911
1907
1913
1913
1987
1927
1907
1985
1907
1908
1907
1908


Argentina
Mexico
USA:CA
Puerto Rico
USA: MI
USA: DC
Brazil?
Puerto Rico
Haiti
Puerto Rico
USA: FL
USA: CA
USA: CA
Cuba
Argentina?
Argentina
USA: LA
USA: ID
Argentina
Guadeloupe
Haiti
Argentina
USA: DC
Argentina
USA: FL
Puerto Rico
USA: CO
USA: DC
USA: MA
Peru
Brazil
Chile
Puerto Rico
USA: FL
USA: DC
USA: CA
Mexico
USA: FL


?Aleyrodidae
Aleyrodidae
Diaspididae
Aleyrodidae
Diaspididae
Diaspididae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Diaspididae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Diaspididae
Diaspididae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Diaspididae
Diaspididae
?Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Aleyrodidae







Table 1.3. Extralimital Encarsia species occurring in the New World.



Species Author Year Country Host


armata
clypealis
diaspidicola
divergens
fasciata
inaron
lahorensis
lounsburyi
lutea
merceti
opulenta
sankarani
smith
strenua
transvena


Silvestri
Silvestri
Silvestri
Silvestri
Malenotti
Walker
Howard
Paoli & Berl.
Masi
Silvestri
Silvestri
Hayat
Silvestri
Silvestri
Timberlake


1927
1928
1909
1926
1917
1839
1911
1916
1909
1926
1928
1989
1926
1927
1926


Vietnam
Vietnam
South Africa
Singapore
Italy
U.K.
Pakisan
Italy
Italy
Singapore
China
India
China
Macao
USA: Ha


Aleyrodidae
Aleyrodidae
Diaspididae
Aleyrodidae
Diaspididae
Aleyrodidae
Aleyrodidae
Diaspididae
Aleyrodidae
Aleyrodidae
Aleyrodidae
Diaspididae
Aleyrodidae
Aleyrodidae
Aleyrodidae


limited. Such information could provide clues for species determination and

utilization.

Eleven of the 16 species that were described from the United States were

collected in California, Florida, or Washington, D. C. Only 2 species have been

described from Mexico; 8 of the 10 South American species were described from

Argentina or Brazil; and 5 of the 9 Caribbean species were described from Puerto


Rico.






9
Many of the species descriptions lack sufficient detail to allow the species

to be distinguished from similar species. Species identification often necessitates

that specimens be compared with those of the type series or at least reference

specimens identified by an expert on the group. Most species are only known from

the type specimen or type series. This precludes the ability to determine what

intraspecific variation exists in the species. Often it is difficult or impossible to

observe the finer morphological details that can be useful in distinguishing closely

related species in specimens from the type series. Many specimens are nearly 100

years old and were not properly cleared before mounting them in Canada balsam.

Some specimens may show artifacts resulting from the mounting process or the

mounting medium used. Antennal segments may be longer or wider and colors

less pronounced in preserved specimens than in fresh ones.

The presence of cryptic species in the family Aphelinidae is well known and

is a major concern in the identification of Encarsia species. Slight differences in

morphological characters may represent intraspecific variation or may be indicative

of a morphologically similar but distinct species. DNA analyses or detailed

morphological examination of a long series of a species representing different

habitats and environmental conditions may give some insight to the species

identity. However, for biparental species, the question is better addressed by

conducting reproductive crosses of isolated individuals as was done by Rao and

DeBach (1969) for species in the genus Aphytis (Aphelinidae).






10
Morphological characters of male Encarsia are sometimes useful in species

or species group determination. However, in several species, males are not known.

Males do not exist in the uniparental species that reproduce parthenogenetically.

Obviously, it is impossible to use reproductive isolation as a means for

discriminating cryptic species in uniparental species. In other species, males

probably occur but have not been collected. Production of males has been induced

in some thelytokous species by the ingestion of antibiotics or exposure to higher

temperatures. Theoretically, these treatments kill or inhibit the endosymbiotic

bacteria found in the reproductive tract of the female that prevent the development

of the male gametes (Zehori-Fein et al. 1992).

Males of many Encarsia species develop as hyperparasites on females of

their own species or on a different species present in the host insect species.

Therefore, positive association of males and females of the species reared from

the same sample may be dubious. Considering the importance of Encarsia species

in the biological control of whiteflies and the increasing number of species currently

being introduced against the SPWF, it was imperative that the earlier described

species be studied not only to establish their correct identities, but also to avoid

the unnecessary creation of invalid synonyms.










CHAPTER 2


BIOLOGY AND POPULATION STUDIES




2.1. Whiteflies

2.1.1. Geographic Distribution

There are about 1,200 described species of whiteflies (Mound & Halsey,

1978). The majority of these species are tropical in origin, where about 724

species have been described versus about 420 from temperate regions. Strong et

al. (1984) found that the number of species of whiteflies in Europe was correlated

with latitude, ranging from 3 species in northern Sweden to 32 species in southern

Spain.



2.1.2. Systematics and Evolutionary Relationships

The family Aleyrodidae is most closely related to the Psyllidae, which, along

with the Coccoidea and Aphidoidea, forms the Sternorrhyncha in the order

Homoptera. There are 126 genera of aleyrodids within the two subfamilies that

have been designated. The Aleurodicinae, comprised of 17 genera and about 100

mainly Neotropical species, is considered to be the more primitive of the two






12

subfamilies; the Aleyrodinae, to which all other species are assigned, is

cosmopolitan in distribution. The phylogenetic relationships between the genera of

the family are not well understood; there is even some doubt as to validity and

evolutionary significance of some of the characters that have been used to

separate the two subfamilies.



2.1.3. Life Stages

Whiteflies have six life stages: the egg, the crawler (1st nymphal instar or

stadium), two sessile nymphal instars (2nd and 3rd instar), the "pupa" (4th instar),

and the adult or imago. The terms nymphal" and "larval" have been used

interchangeably in the literature to denote the immature forms. The fourth instar

is not a true pupa because feeding occurs during the first part of this stage and

transformation into the adult takes place in the first part without a "pupal" moult.

The term "nymph" will be used to denote the first three immature stages, and the

term "pupa" will be utilized to denote the last immature stage.



2.1.4. Sex Ratio

Gameel (1978) reported the sex ratio of 1:1 for the SPWF on cotton and

Van Lenteren and Noldus (1990) have reported similar sex ratios for Trialeurodes

vaporariorum. The sex ratio of the SPWF is sufficiently variable to preclude

generalization (Butler et al. 1986; Gerling et al. 1986). Horowitz and Gerling (1992)

found that the sex ratio of the SPWF ranged from a female-biased condition early





13
in the season to male biased at summer's end on cotton in Israel. Their study

showed that young females laid proportionately more female-producing eggs than

older females. Like most whiteflies, the SPWF is arrhenotokous, females can

regulate the sex of their progeny as long as they have a sufficient sperm supply.

Mated females can lay both haploid (male) or diploid (female) eggs, whereas

unmated females lay only haploid (male) eggs.



2.1.5. Developmental Rate

Developmental rate, fecundity, longevity and mortality of whiteflies may vary

with temperature, relative humidity, rainfall and host plant (Table 2.1). Favorable

conditions for SPWF development include high temperature, low humidity, periods

of no or low rainfall and mild winters (Jagdev & Butler, 1985). Lower and upper

threshold temperatures for development are 10 and 320C, respectively. Although

the SPWF may develop faster (17 days) at 300 C, it appears that 270C (25-50

days) is a more optimal temperature for overall whitefly population development;

longevity and fecundity of the SPWF decrease at the higher temperatures (Ozgur

et al. 1986). Hendi et al. (1987) reported the following developmental rates (days)

for SPWF instars on tomatoes (300C and 5% RH): preoviposition (1-2), 1st instar

(2-4), 2nd instar, (2-4), 3rd instar (2-4), and pupa (6). Zalom and Natwick (1987)

reported that the SPWF had seven to nine generations per year in the

southwestern United States. They determined the number of degree days the

SPWF required to complete its development in field cages (degree-days) as eggs







Table 2.1 Developmental time (days) of Bemisia tabaci reared at different
temperatures on various host plants.


Temperature (OC) Host Dev. Reference
time


16.8
21.3
25

14.9
30.0

27
30

30.0

24.1 RH=70%
25.3 RH=69.8%


cotton
cotton
cotton

cotton
cotton

cotton
cotton

tomato

bean
bean


68
36
24.2

65.1
16.6

25-50
17

21

28.3
25.3


Coikesen et al. 1987



Butler et al. 1983


Ozgur et al. 1986


Hendi et al. 1987.

Eichelkraut 1986


(27), nymphs (268.5) and pupae (73); the average generation time was achieved

at 369.5 degree-days.

Developmental rate may differ for different host plants (Table 2.1) or

varieties. (Boica & Vendramin 1986). Overall population development may depend

on the characteristics of the leaf surfaces of the host plant. Pubescent leaf and

closed- canopy cotton varieties were reported to be much more heavily infested

than glabrous, smaller-leaf, open-canopy varieties (Ozgur et al. 1986). Moderate

pubescence seems preferable as it provides a physical barrier to natural enemies

and a favorable microclimate.






2.1.6. Fecundity

Fecundity is highly variable and may be influenced by plant host species,

cultivar, physiological state of the host plant, temperature and humidity. Ogzur et

al. (1986) reported that fecundity of the SPWF varied from 80 to more than 300

eggs per female on cotton in the Sudan. Butler et al. (1983) found that the average

number of eggs per female SPWF on cotton was 81 and 72 at 26.7 and 32.20C,

respectively. Van Lenteren and Noldus (1990) reported that T. vaporariorum

fecundity was constant from 18 to 27C but decreased at lower and higher

temperatures. They also reported higher fecundity rates forT. vaporiorarum reared

on eggplant versus that for those reared on cucumber, tomato or sweet pepper.

The ovipositional rate is age-dependent, gradually increasing during the first days,

reaching a plateau shortly before the female dies.



2.1.7. Longevity

Longevity of adult whiteflies varies on different host plants and at different

temperature and relative humidity regimes but may also vary on the same host

plant. SPWF adults lived 10 to 15 days in the field during summer temperatures

(around 280C) and 20 to 60 days in winter (around 15C) (Gerling, Horowitz and

Baumgartner, 1986). Adult lifespan was 8 to 43 and 17 to 27 days, for SPWF

males and females, respectively.







2.1.8. Mortality

Both abiotic and biotic factors affect SPWF mortality (Figure 2.1). Abiotic

factors include temperature, relative humidity, wind, and rain; biotic factors include

parasitoids, predators, pathogens, and host nutrition. Unfavorable levels of

temperature, humidity and rainfall are the most important mortality factors in

crawler and first instar SPWF nymphs. Parasitization is the most important

mortality factor during the advanced nymphal stages (Horowitz et al. 1984). The

degree of mortality to immature stages may vary on different host plants, and even

on the same host plant. Lenteren and Noldus (1990) found that pre-adult mortality

of T. vaporiorarum was much higher on sweet pepper than on tomato, cucumber

or eggplant. Hendi et al. (1985) observed high natural mortality (64.4 to 97.5%) of

SPWF on tomatoes in Egypt.



2.1.9. Host Plant Specificity, Suitability and Preference

There have been over 500 hosts recorded for the SPWF from 74 families

of crop and weed species. Van Lenteren and Noldus (1990) pointed out the

distinction that needs to be made between a preferred host species and a suitable

host species. Whiteflies often develop faster, have lower immature mortality and

higher fecundity rates on a preferred host or variety than on a suitable host. Host

plant suitability can be affected by the addition of nitrogen fertilizers (Onillon et al.

1986).







2.1.10. Migration

SPWF adults demonstrate two distinct flight patterns. Short-distance flight

occurs under the plant canopy. Newly emerged adults leave the lower leaves from

which they emerged and move to the upper leaves to feed and oviposit. Long-

distance flight, sometimes called the dispersal phase, occurs when adults take off

from their host plant, get caught in an air current and drift passively, sometimes

for several miles. Temperature and light thresholds affect the number of SPWF

adults taking flight (Bellows et al. 1988). SPWF populations are considered to be

continuously mobile in areas where a variety of host plants are consistently

available. Intensification of agriculture and off season production of new suitable

host plants of the SPWF provide bridges between cotton seasons and may lead

to the current increased pest status (Coudriet et al. 1985).



2.1.11. Population Dynamics

Many biotic and abiotic factors affect the population dynamics of the SPWF.

Temperature typically affects fecundity, mortality and developmental rates; rainfall

and relative humidity are other important abiotic factors. Nutritional content,

development and physical characteristics of the host, availability of alternate hosts,

natural enemies (predators, parasitoids and fungi) and man's activities through the

use of cultural, chemical and other control practices also play important roles in the

overall population dynamics of the whitefly.







2.2. Parasitoids

The genus Encarsia belong to the Aphelinidae, a small-sized family of the

chalcidoid Hymenoptera. There are about 175 described species of the genus

Encarsia (Hayat 1989). The majority of Encarsia species are parasitoids of

whiteflies, but several are parasitoids of diaspine scales and a few have been

reared from other hosts, such as from Lepidoptera eggs. The genus is more highly

represented in the Palearctic and Oriental region where more species have been

described. However, the disparity in the number of species in the Orient versus

other regions of the world may be due, in part, to the greater effort that has been

made, primarily by Viggiani and Hayat, in documenting the fauna of that region,

versus little effort that has made in the other regions.



2.2.1. Life Cycle and Development

Some Encarsia species are uniparental, but the majority are biparental.

Mated females can lay both fertilized diploid eggs that become females, and

unfertilized haploid eggs that become males. Female eggs are usually laid in the

haemocoel of aleyrodid and diaspine scale insect hosts and develop into primary

endoparasitoids. In most of the biparental species that have been studied, the

male eggs are laid on the outside of the body of the larval female primary

parasitoid. The males develop as hyperparasites on females (or males) of their

same species (adelophoparasitoid) or on a different parasitoid species in the host

insect. Females of E. smith may develop either as primary parasitoids of the citrus






19
blackfly or as hyperparasitoids of E. opulenta (Ru & Sailer 1987). The production

of males and females may be affected by the availability of mates and suitable

hosts, and the action of symbiotic bacteria in the reproductive tract of the females

that inhibit the production of male gametes (Godfray & Hunter, 1993). Also, Wilson

and Woolcock found that sex was determined in Ooencyrtus submetallicus

(Encyrtidae) by the temperature to which the females were exposed. Females are

synovigenic. Eggs are produced in two ovaries, each composed of three ovarioles.

The eggs develop and mature continuously throughout the female's lifespan but

most are laid during the first few days after emergence. Fecundity in Encarsia

species may be highly variable, depending upon the parasitoid species,

temperature, relative humidity and host nutrition. Fecundity data for several species

of Encarsia are given in Table 2.2.




Table 2.2. Fecundity of six Encarsia species and Eretmocerus mundus.


Species TempoC eggs/? Reference


E. formosa 17 165.6 Vet and van Lenteren, 1981
E. formosa 25 442.2 Arakawa, 1982
E. inquirenda 22-26 20.6 Gerson, 1968
E. lahorensis 26 20-50 Viggiani, 1984
E. luteola 25 63 Gerling et al. 1987
E. luteola 22-30 57.8 Gerling et al. 1987
E. meritoria 17 85.4 Vet and van Lenteren, 1981
E. pergandiella 26.6 47 Gerling, 1985
E. pergandiella ? 48.5 Viggiani, 1984
E. pergandiella 17 124.9 Vet and van Lenteren, 1981







20

Encarsia species pass through three larval instars. The first and second

instars have no functional spiracles. The third instar has open, functional spiracles

and voids its meconia before moving on to the prepupal and pupal phases. The

position of the meconia in the host body can differ for different species. For


Table 2.3. Developmental time (days) of
Eretmocerus mundus.


nine Encarsia species and


Species 0C Egg Larva Pupa Total References


E. desert 9 25

E. formosa 9 24

E. inquirenda 9 20
E. inquirenda 9 28

E. lahorensis 9 24
E. lahorensis 24
E. lahorensis 9 25
E. lahorensis e 25

E. lutea 9 25
E. lutea d 25
E. lutea 9
E. lutea 9 15
E. lutea d 15
E. lutea d 28

E. meritoria 9 12
E. meritoria 9 28

E. pergandiella 9 24
E. pergandiella d 24

E. quaintancei

E. transvena

Eretmocerus mundus 9 -
Eretmocerus mundus 9 25


8-17
9-12

8
7

16.1
25.6
8.8




7
9


10.5
- 10.5


- 11-13

15

62.3
26.3

24.5
12-15
7-8 20-28
7-9 19-24

6 13-18
5 14-15
15
6.2 22.3
14.8 40.4
5.7 14.5

75.0
11.0

5 15
4-5 14

- 10-25

12.9

16.6
10 20.5


Gerling et al. 1987

Gerling, 1983

Gerson, 1968
Gerson, 1968

Ru & Sailer, 1979
Ru & Sailer, 1979
Viggiani, 1978
Viggiani, 1978

Gerling & Foltyn, 1987
Gerling & Foltyn, 1987
Vet and Lenteren, 1980
Abdel et al. 1987
Abdel et al. 1987
Abdel et al. 1987

Avilla et al. 1991
Avilla et al. 1991

Gerling, 1966
Gerling, 1966

Dysart, 1966

Kapadia & Puri, 1990

Kapadia & Puri, 1990
Foltyn & Gerling, 1985





21
species in which the males develop as hyperparasites, the first instar male larvae

develop inside the hymenopterous host and the second instar exits the host and

develops to maturity as an ectoparasitoid. The developmental rate of males may

be equal to or shorter than that of the female. Females may all emerge within a

few days, or have their emergence spread out over one to two weeks. This

assures insemination of females even in small, isolated host populations (Gerling

et al. 1987). Hunter (1989) found that hyperparasitic male E. perqandiella

developed faster on 9-day-old (pupal) E. perqandiella females than on 7-day-old

females. Data suggested that single release colonization is likely to be limited by

a delay of 7 to 9 days between the oviposition of female eggs and suitability of

these females for oviposition of male eggs. As males take an additional 15 to 16

days to develop, the total time elapsed between the time when colonizing females

lay eggs and the first mating of F1 female progeny is 22 to 25 days.

The duration of the developmental period is linked to climatic factors and

to the condition of the host. The rate of parasitoid development depends primarily

on temperature and is usually shorter for males (Table 2.3). Female E.

pergandiella (24C) develop in 15 days and males in 13 to 14 days. (Gerling,

1966). Avilla et al. (1991) reported that E. meritoria females completed their life

cycle in 11 days at 280C and in 75 days at 12C. Flanders (1960) found that at

constant temperature (800F), E. perniciosi completed its life cycle in about 30 days

if eggs were deposited in the first-instar host, 18 days if they were deposited

immediately before the host scale was half grown, and 26 days if they were

deposited after the host was half grown. Development may be highly variable for






22
different Encarsia species. E. transvena developed in an average of 12.9) days

(Kapadia & Puri, 1990), while most other species complete their development in

15 and 25 days (Table 2.3).


2.2.2. Longevity

Adult longevity is dependent primarily upon reproductive physiology,

available food, temperature and humidity. Greatest longevity was recorded for E.

formosa (99 days at 15-160C) and E. desert luteolala) (84 days) (Gerling, 1992).

Most whitefly species are tropical or subtropical in origin and do not have a winter

diapause, but their development slows and their populations decrease during the

cooler or drier months. Those that overwinter do so in preimaginal stages. In

Israel, E. lutea overwinters on SPWF nymphs on Lantana and other weed species.

(Gerling, 1984). E. lahorensis females oviposit in diapausing stages of Dialeurodes

citri; the egg hatches but the first instar larva does not develop into the second

instar until its host has resumed activity and has become partially inflated (Viggiani

1984). Summer survival may vary greatly since it involves intensive host-

associated activities such as searching and ovipositing. Gerling (1990) considers

the "effective longevity", that time during which the females lay most of their eggs,

to be a more meaningful parameter for understanding reproductive strategies than

are total longevity. Synovigenic whitefly parasitoids have been found to complete

their effective longevity in less than 20 days at temperatures above 200C.







2.2.3. Host Feeding

Host feeding by parasitoids on whitefly immatures has been reported forE.

formosa (Gerling, 1966), E. pergandiella (Gerling 1966b), E. opulenta (Dowell et

al. 1981), E. desert, E. transvena and E. lahorensis (Gerling, 1990) and probably

is a common occurrence. The female thrusts her ovipositor through the nymph's

body and laps up the exudate. Host feeding has been found to increase longevity

and promote oogenensis (fecundity) in Encarsia females and may be an important

mortality factor, especially in younger whitefly instars. Arakawa (1982) reported that

each female E. formosa is able to kill an average of 423.7 and 101.3 host by

parasitization and host feeding, respectively, for a total of 534 hosts attacked.

DeBach (1943) reported up to 56% host mortality caused by feeding of

Metaphycus helveolus (Compere) on black scale, Saissetia oleae (Bern.). The

number of scales killed by host-feeding, which was largely when the scales were

rather small for parasitization, was much greater than the number killed by

parasitization. The relative importance of host-feeding was greater during the early

phases of an infestation, whereas that of parasitization is higher later, as the

scales increased in size. The value of parasite predatism is affected by the relative

abundance of the host and parasite. Higher levels of host-feeding occur at lower

host's density and with an increase in proportion of host below the size suitable for

parasitism (Flanders 1953). Bartlett (1964) found that Microterys flavus (Howard)

mutilated Coccus hesperidum L. scales when host-feeding was initiated but the

host did not bleed freely. Host-feeding tendencies developed in individuals






Pages
Missing
or
Unavailable






25

which later hyperparasitize Eretmocerus individuals inside SPWF nymphs (Gerling,

1983). E. tricolor females were reported to preferentially hyperparasitize female E.

inaron rather than their own species. E. opulenta preferentially hyperparasitizes

whiteflies containing Amitus females (Dowell et al. 1981). Females are able to

distinguish between hosts parasitized by conspecifics but not those parasitized by

other species. Some Encarsia species may develop as tertiary parasitoids of males

of their own species or different species, creating a very complex ecosystem

(Figure 2.2).



2.2.5. Adelphoparasitism

Various authors have speculated on the evolutionary significance of

adelphoparasitism in the Aphelinidae. Hassell et al. (1983) showed that

theoretically adelphoparasitism confers stability on the host-parasitoid dynamics.

Walter (1988) argued that male heteronomous aphelinids have host relationships

that differ from those of their conspecific females. In an ancestral species,

deposition of female eggs became uncoupled, in terms of behavior, at oviposition,

from the deposition of male eggs. During the evolution of heteronomous host

relationships, males and females would have been subject to different selection

pressures. As a result, males must have become adapted to their hosts

independently of females becoming adapted to theirs. Godfray et al. (1992) stated

that in general the observed sex ratio will be strongly correlated with the relative

abundance of parasitized and non-parasitized hosts. If the hosts are rare, search





26
time will limit reproductive success, and the female will lay a male or female egg

when she finds a host. Lifetime fitness is maximized by parasitizing all suitable

hosts that are encountered.



2.2.6. Population Dynamics

Several studies have examined the dynamics of the SPWF parasitoid

complex in different regions of the world. Puri et al. (1990) reported that E.

transvena and Eretmocerus mundus were the primary parasitoids of SPWF on

cotton in India. Maximum parasitization (54.6%) was reported during the more

humid months of the year. E. lutea and Eretmocerus mundus are the major SPWF

parasitoid species in the Sudan. Izaguirre (1993) found that E. pergandiella made

up 51 to 87% and E. nigricephala 5 to 38 % of the SPWF parasitoid fauna on

beans in Honduras. Interestingly, Eretmocerus only represented a maximum of 4%

of the total parasitoids collected from various climatic areas during the two-year

study. Gerling (1966) reported on the bionomics of the parasitoid complex of

Trialeurodes abutiloneus and B. tabaci on cotton in California.
















































9 I EMIGRATION




A s CULTURAL PRACTICES
PARASITISM crop rotation
M rate of planting
cultivation
Scrop residue removal

PESTICIDE APPLICATION


PATEOCENS


HYPER-
PARASITISH

SOIL FACTORS
fertility
structure
water
biotic factors


















Figure 2.1. Factors affecting the development and mortality of the SPWF.





























Phytophagous Primary Secondary Tertiary
host parasitism parasitism parasitism

Encarsia <-- Encarsia
pergandiella rpergandiella e

Encarsia Encarsia Encarsia
__...----7'nigricephala nigricephala d pergandiella d


E. strenua
males unknown


Figure 2.2. Intra- and interspecific relationships of parasitoids of the SPWF.










CHAPTER 3


HISTORICAL REVIEW AND HIGHER CLASSIFICATION



3.1. Introduction

The genus Encarsia belongs to the Aphelinidae, a small-sized family of the

chalcidoid Hymenoptera, containing forty-nine valid genera (Hayat 1985) and

approximately 860 species. Morphologically, the Aphelinidae show affinity to both

the Eulophidae and the Encyrtidae, and have in the past been placed as a

subfamily of each one of them in the past (Ashmead, 1904; Peck, 1951; Burks,

1967; Riek,1970). Most aphelinid genera resemble the Eulophidae in the structure

of the mesonotum (parapsidal grooves complete, axillae advanced into the base

of the parapsides) and in the small number of antenna segments. Other genera,

especially Coccobius, Marietta and Eutrichosomella, resemble the Encyrtidae in the

shape of the mesopleura, in the large saltatorial mid-tibial spur, and in the

presence of a well-developed strigil on the foreleg. Peck et al. (1964) regarded the

family as a transitional taxon between the Eulophidae and the Trichogrammatidae.

The group was given family rank by Foerster (1856), who established the

family Myinoidae, based on the genus Mvina Nees, a junior synonym of Aphelinus





30
Dalman. The family is also closely allied to the Signiphoridae (Woolley, 1988).

Several schemes have been suggested for the subfamily and tribal classifications

of the family. Early workers (Howard, 1907; Ashmead, 1904) divided the then

subfamily Aphelininae (Encyrtidae) into two tribes the Aphelinidini with 5-

segmented tarsi and the Pteroptricini with 4-segmented tarsi. De Santis (1946)

offered the first modern day classification recognizing three subfamilies:

Aphelininae, Coccophaginae and Calesinae. Ferriere (1964) added a fourth

subfamily, the Eriaporinae. Yasnosh's (1976) classification of the family into seven

subfamilies was not accepted by other workers (DeBach and Rosen, 1976). Hayat

(1985) proposed four subfamilies and eight tribes. Viggiani and Battaglia (1984)

defined five aphelinid subfamilies based on male genitalia characters, placing the

genus Eretmocerus as the sole member of the subfamily Eretmocerinae. They did

not recognize the subfamily Eriaporinae and placed the genus Eriaphvtis,

traditionally placed in the Eriaporinae, in the subfamily Aphelininae. The first

"modern" key to the world genera of Aphelinidae was published by De Santis

(1946). Ferriere (1965) and Nikolskaja and Jasnosh (1966) published keys to the

genera based on females only. Hayat (1983;1985) provided the most recent

comprehensive keys to the aphelinid genera.



3.2. New World Studies in the Genus Encarsia

Major contributors to the knowledge of the North American and Caribbean

Encarsia fauna include L. O. Howard (1894-1912), who described thirteen species:






31
United States (10), Mexico (1), and Puerto Rico (2); A. B. Gahan (1927-1931), 6

species: U.S.A. (3), Puerto Rico (2) and Cuba (1); and Dozier (1932-1937), 4

species: U.S.A. (1), Haiti (2) and Puerto Rico (1). Other early contributors include

Tower (1913) and Cockerell (1911). Modern-day contributors include DeBach and

Rose (1981); Viggiani (1989), who described one species from the U. S. and

provided figures and additional notes on many of the previously described New

World species; and Polaszek et al. (1992).

De Santis is the major contributor to the knowledge of the South American

fauna of Encarsia. He described four species of Encarsia (two currently valid),

redescribed in more detail several species, and has made major contributions to

the overall study of the Aphelinidae and other chalcidoid families from the area.

Several authors have described single species from South America (Blanchard,

1940; Hempel, 1904; Marelli, 1933; Mercet, 1927; Silvestri, 1936; Viggiani, 1987).



3.3. Generic Concepts

Encarsia Foerster 1878:65.

Type-species: Encarsia tricolor Foerster by original designation.



Synonyms

Aleurodiphilus DeBach and Rose 1981:659. Type-species: Aleurodiphilus

americanus DeBach and Rose, by original designation. Synonomy by Hayat

1983:85.





32
Aspidiotiphaqus Howard 1894a:229. Type-species: Coccophagus citrinus Craw, by

original designation. Synonomy by Viggiani & Mazzone 1979:44.

Doloresia Mercet 1912:294. Type-species: Doloresia filicornis Mercet by original

designation. Synonomy by Mercet 1930a:191.

Mimatomus Cockerell 1911:464. Type-species: Mimatomus peltatus Cockerell, by

monotypy. Synonomy with Prospaltella by Girault 1917:114.

Paraspidiotiphacus Alam 1956:359. Type-species: Aspidiotiphagus flavus

Compere, by original designation (as subgenus of AspidiotiphaCqus).

Prospalta Howard 1894b:6. Type species: Coccophacus aurantii Howard,

designated by the ICZN under its Plenary Powers, Opinion 845

(preoccupied by Prospalta Walker, 1857, in Lepidoptera).

Prospaltella Ashmead 1904:126. Replacement name for Prospalta Howard nec

Walker. Synonomy by Viggiani & Mazzone 1979:44.

Prospaltoides Brethes 1914:12. Type-species: Prospaltoides howardi Brethes by

original designation, monotypic. Synonomy with Aspidiotiphaqus by

Brethes 1916:429. Ferriere (1965) considered type species a synonym of

Encarsia citrina (Craw).

Trichaporus Foerster 1856:2. Nomen nudum, LaSalle & DeBach 1982:297.

The genus Encarsia is one of 16 genera included in the subfamily

Coccophaginae, sensu De Santis (1948) and Hayat (1985), and has been placed

in the tribe Pteroptricini (Hayat, 1989). Monophyly of the genus Encarsia is

supported by the short axillae, by the pattern of mesoscutal setae (Polaszek and





33
Hayat, 1992) and by the spoon-shaped phallophase of the male genitalia (Viggiani

and Mazzone, 1982). The genera Coccophagoides Girault, Coccophaqus Howard,

Dirphys Howard and Encarsiella Hayat are considered to be its closest relatives

(Polaszek and Hayat, 1992).

Foerster (1878) erected the genus Encarsia with E. tricolor as its only

included species. Walker described the genus Prospalta and designated

murtfeldtae as its type species. Ashmead (1904) replaced the name Prospalta with

Prospaltella as it was preoccupied by the genus Propalta Walker (Lepidoptera).

Mimatomus Cockerell and Doloresia Mercet were synonomized with Prospaltella

by Girault (1917) and Mercet (1930a) respectively. Viggiani and Mazzone (1979)

synonomized AspidiotiphaCqus Howard, Prospaltella and Trichoporus (Foerster) with

Encarsia. Hayat (1983) synonomized Aleurodiphilus DeBach & Rose with Encarsia.

The synonomy of Aspidiotiphagus with Encarsia was not accepted by all workers

(DeBach & Rose 1981), but Hayat (1989) agreed that the characters used to

separate the genera were not considered of generic value and suggested that the

differences only warrant the designation of a separate species group. The

synonymy of Encarsiella Hayat with Encarsia proposed by Shafee and Rizvi (1984)

was not accepted by Hayat (1989) because it was based only on the literature and

not of study of relevant material. The generic name Trichaporus (Foerster) is

considered a nomen nudum because Foerster who created the genus, failed to

assign a species to it. Subsequently, Ashmead designated Trichoporus

columbianus in 1900 as the type species for the genus Trichoporus. Both





34
Trichaporus Foerster and Trichoporus Ashmead are objective synonyms. LaSalle

and DeBach (1982) stated that these genera are properly placed in Eulophidae,

not in the Aphelinidae, where they are both senior objective synonyms of

Galeopsomvia Girault (a eulophid) and sought suppression of the name by the

International Commission of Zoological Nomenclature (ICZN).

Hayat (1989) provided a detailed description of the genus Encarsia and of

the diagnostic characters that separate it from other closely related genera. Briefly,

antennae 8-segmented, excluding the radicle and anellus, and occasionally 7-

segmented in some males in which the F5 and F6 are partially or completely

fused; tarsal formula 5-5-5 rarely 5-4-5; mesoscutum with 0-18 setae (usually 4-12)

arranged in bilateral symmetry; scutellum with 2 pairs of setae (extranumeral setae

rarely occur); hypopygium not prominent and rarely extending beyond the cercal

plates; two setae on the submarginal vein (except in lounsburyi, which has one,

and pulliclava, which has several); strong seta (mvb) at the juncture of the

submarginal vein and marginal vein; forewing never with speculum; gaster with

seven tergites; male genitalia with the phallophase several times longer than wide,

with a truncate or rounded apex and without digiti; aedeagus longer than the

phallophase; body color variable with males generally darker than females.

Encarsia may be distinguished from Coccophaqus and Coccophaioides by having

only two setae on the submarginal vein and scutellum whereas in the latter two

genera, there are at least three setae on the submarginal vein and usually three

(rarely two) setae on the scutellum (Figure 6.1i,k;6.3e,f;6.5b,c) Also, the marginal





35
vein is less than or equal to costal cell, the antennae are spindle-shaped with apex

pointed; and the hypopygium is prominent, reaching at least to level of cercal

plates in Coccophaqoides.

Encarsia may be distinguished from Dirphys and Encarsiella, which have the

axillae long, mesally separated by a distance of less than the length of an axilla.

The mesoscutal setae are dense (more than 25) and scattered (Figure 6.4g,h).

The scutellum width is less than 1.5 times its length, the submarginal vein has two

prominent setae and usually with one to five setae at its base, and the club is often

oblique and much wider than the funicle segments (Figure 6.1c,d). (Refer to pp.

231-232 of Hayat [1985] for a taxonomic key to separate these genera).










CHAPTER 4


MATERIALS AND METHODS


4.1. Natural Enemy Survey

4.1.1. Collection

A statewide survey of the natural enemies of the SPWF was initiated in

1988 to gather information on the parasitoid complex of the SPWF in Florida and

the humid neotropics, and to identify promising natural enemy species from the

neotropics (and other areas of the world) for eventual introduction into Florida and

the southern United States. The objectives of the survey were to:

1) identify the species of parasitoids commonly found parasitizing SPWF.

2) measure the ratio between emergent SPWF and parasitoids as a comparative

indicator of parasitism.

3) identify important SPWF wild host plant reservoirs.

4) identify changes in the parasitoid complex composition attributable to seasonal,

geographic and host plant variation.

5) determine if SPWF and its parasitoids can overwinter in northern Florida

In addition, parasitoids were reared from several other whitefly species in order to

determine the host range for each of the parasitoid species reared from SPWF.

(Bennett et al. 1990)






37

Samples of SPWF were collected from over 117 crop and weed species in

28 counties (Figure 4.1) in Florida by Dr. Fred D. Bennett, Dr. David Schuster,

Emily Vasquez and other University of Florida cooperating scientists and field

personnel. In addition, collections of parasitized SPWF were made in many

Caribbean and Latin American countries (Figure 4.2), primarily by Dr. Bennett

while traveling on other projects.

Host plant material was examined for the presence of whitefly pupae, after

which the leaves containing higher numbers of whitefly pupae were preferentially

selected and placed in 1/2-pint, pint or quart size paper cans (Gainesville Paper

Co.) for parasitoid emergence. The inside lid of the paper can was covered with

a paper tissue or fine netting before closing to prevent the parasitoids from

escaping upon emergence. This method was preferred because it requires less

handling time and increased the probability of parasitoid emergence by confining

a larger number of parasitized hosts with less manipulation. However, erroneous

host records may result when more than one host species are present in the

sample. The plant material was examined carefully under the dissecting

microscope for any other insects present, and were removed before placing the

plant material into the container. To ensure accurate host records, each individual

host should be isolated by removing a small leaf-disc bearing a single host with a

hole punch and placing it into an individual vial. Whether the method used to

collect and hold SPWF nymphs for parasitoid emergence equally affects the

number of adult whiteflies and parasitoids or the species of parasitoid that emerge,






38
is not known. The emergence of parasitoid species, such as Eretmocerus that

attack earlier nymphal stages and have a longer life cycle than Encarsia species,

may be more affected as the host plant material held for parasitoid emergence,

deteriorates. Data presented of the relative abundance of each parasitoid species

in foreign countries are based on collections made on certain host plants at certain

time of the year and at certain locations. They may not be representative of the

true relative abundance of each species in the area.

Each collection was given an accession number and the following collection

data recorded: 1) locality country, state or department, city or town, property; 2)

date of collection; 3) collector; 4) host insect; 5) host plant; 6) infestation level -

very high, high, medium, low, and very low; 7) number of leaves per sample; and

8) field notes. Rating the infestation levels was done subjectively, and varied with

the size of the leaf.

The contents of each container were examined three to four weeks after

collection. The dry, dead wasps were sorted to species, counted, sexed (for some

localities), then removed with a fine brush or a minute pin glued to the end of a

2-mm diameter dowel rod. Microscope slide preparations were made of the host

and parasitoids, when necessary to identify or confirm the identification of the

species. Specimens representing new host or distribution records or other special

cases were processed similarly. Additional parasitoid specimens were stored dry,

in small glass vials or gelatin capsules with some loose cotton to prevent their

movement and breakage. Records were kept of the number of adult whiteflies, and






39

of females and males of each parasitoid species that emerged. Collections of other

whitefly species were processed similarly, with the exception that only the identity

of each parastiod was recorded. Slide labels were printed on dry gum paper with

the laser printer using Word Perfect 5.1 software. Collection data and parasitoid

identifications were input into PARADOX 3.5 database software for storage,

retrieval and analyses of data.

The number of leaves collected per sample sometimes varied for different

plant species. For example, 30 or more leaves per sample were collected of

Chamaesvce hyssopifolia, which has small leaves, while only 3 to 4 leaves per

sample were collected of Brassica oleracea, a large leaf host. Collections were

made periodically in specific areas from certain plants while other collections were

made opportunistically whenever an infestation of SPWF was encountered.

Collections were not made at random since the primary focus of the survey was

to determine the identity of the parasitoid species that attack the SPWF.

Leaves containing higher numbers of late-instar whitefly pupae were preferentially

selected to be held for parasitoid emergence, in order to maximize the number of

adult parasitoids that emerged. Most collections were made in areas that were not

treated with pesticides, such as roadsides, organic gardens and field plots that

were not treated. Nevertheless, there were several collections made in areas that

had been sprayed with pesticides.

Percent parasitism was calculated by dividing the number of parasitoids by

the sum of the number of SPWF and parasitoid adults that emerged. McAuslane





40
(personal communication) found that direct observation of parasitized SPWF late-

instar pupae on peanut leaves, determined by the displacement of the mycetomes

in the nymphal body, overestimated the percent parasitization or emergence data

underestimated the percent parasitization. The data, although highly variable,

showed that the number of adult whiteflies that emerged was about 50% that of

the number of 4th instar nymphs and red-eyed nymphs (pharate adults) present

on the leaf. Percent parasitization was about 16% of the number of observed

parasitized SPWF pupae.



4.1.2. Mounting and Preservation

Whitefly late instar nymphs were cleared in 10% KOH, stained in "Wilkey's

double-stain", and passed through baths of 70 and 95% alcohol, a 1:1 mixture of

95% and clove oil, and pure clove oil before mounting them in Canadian balsam.

They were then sent to Dr. Avas Hamon, Division of Plant Industry, Gainesville,

Florida for identification. Most of the slide-mounted parasitoid specimens were

processed by clearing them in lactophenol for three to four days. They were then

bathed in Nesbitt's formula to remove excess lactophenol, and mounted, with

wings and appendages extended, on microscope slides in Hoyer's medium (see

Krantz (1978) for formulae for Hoyer's and Nesbitt's solutions). Slides with

mounted specimens were placed on a hot plate at 500C for about two weeks or

until dry before sealing the edge of the coverslip with a ring of GLYPT solution.

Hoyer's medium was preferred because it requires less manipulation of the





41

specimen and processing time, better preserves the color of the specimen,

provides a better resolution and reveals more details than using balsam. Also,

antennae and other body structures may collapse while processing specimens for

balsam mounts unless special care is taken in processing the specimen. However,

since Hoyer's is not a permanent medium, it is uncertain how long the mount will

last before becoming cloudy or crystallized as the Hoyer's medium deteriorates.

Therefore, it is recommended that specimens be mounted in balsam to ensure

their longevity, especially those representing new species or other specimens of

special interest. Noyes (1982) gives a thorough procedure for mounting

specimens in balsam. Specimens are dissected and the head, wings, antennae

and body are placed under four separate 3-mm microcoverslips. More accurate

measurements of the body parts are achieved and better photographs are taken,

because each dissected body part is flat and on the same focal plane.

Storage in alcohol, for conventional taxonomy, was avoided. Specimens

stored in alcohol when placed directly in lactophenol do not clear sufficiently and

make unsatisfactory slide mounts. If specimens have been preserved in alcohol,

they should first be dried before beginning the slide mounting process. Specimens

were not pointed or mounted on cards because of their small size. Also, they tend

to shrivel and many of the characters used for identification cannot be observed.

However, for DNA analyses or Scanning Electron Microscopy, specimens should

be preserved frozen at a temperature of no higher than minus 200C, preferablyy

minus 800C). Specimens can be preserved in 80% ethyl alcohol for Scanning





42
Electron Microscopy. Alcohol storage of specimens is not highly recommended for

DNA analyses; when used, specimens should only be stored for short periods of

time.

4.2. Systematic Studies

Several thousand slide-mounted specimens of parasitoids reared from the

SPWF and at least forty other whitefly species were examined. The majority of the

specimens were collected in Florida, but specimens from California, Georgia,

Louisiana, Maryland, New York, South Carolina, Tennessee, Texas, Mexico,

Guatemala, Honduras, El Salvador, Costa Rica, Puerto Rico, Jamaica,

Guadeloupe, Grenada, Venezuela, Colombia, Bolivia and Brazil, were also

examined. Type specimens and additional specimens of several described species

were borrowed from the U. S. National Museum and El Museo Nacional de La

Plata, Argentina, for examination. Extralimital specimens reared from the SPWF

and other hosts collected in India, Thailand, China, Hong Kong, Egypt and Sudan

were also examined.



4.3. Morphology and Explanation of Terms and Measurements

Rosen and DeBach's (1976) have discussed in detail the morphology of the

genus Aphytis. With few exceptions, most of their discussion is relevant to the

genus Encarsia. Body measurements were taken as prescribed by Hayat (1989)

from slide-mounted specimens. The body is divided into three well-defined regions

(Figure 4.3): 1) the head, including the eyes, mouthparts and antennae; 2) the

mesosoma, comprising the three thoracic segments (pronotum, mesonotum and






43
metanotum), legs and wings. The mesonotum is divided into the mesoscutum (or

midlobe), the axillae and the side lobes (or parapsides). The metanotum consists

of the scutellum and metanotal ridge. Between the metanotum and the gaster is

the propodeum, which is actually the first abdominal segment; 3) the gaster which

is separated from the propodeum by the petiole, is composed of the seven

subsequent abdominal segments designated as tergites 1-VII. Tergum VII is

sometimes referred to as the syntergum.

The body length was measured from the anterior margin of the head to the

most posterior point of the gaster. The head was measured across its maximum

width. Setae found on the head are prefixed by an "h". Setae found on the

postoccipital bars (hb) are designated as hbl (central pair), hb2, hb3, etc. (moving laterally toward
the compound eyes). Posterior to the postoccipital bars is a row of prominent setae designated as

"hp" setae (Figure 4.3). The mandibles are usually tridentate and the maxillary palp 1-

(rarely 2) segmented. The number of setae between the antennal bases and Table

occipital triangle, the number of labial and maxillary palp segments and sculpturing

of the occiput and stemmaticum were also noted.

When counting the number of antennal segments, the radicle and anellus

are excluded. Therefore, for female Encarsia species the antennae is 8-

segmented, consisting of the scape, pedicel and flagellum. The flagellum is 5 or

6-segmented consisting of 2 to 4 funicle segments and 2 to 4 club (or clava)

segments. For simplicity, the funicle and club segments will be labeled

consecutively as F1, F2, F3, F4, F5 and F6 (Figure 4.4 b,c). The club may be 3-

segmented (formed by F4, F5, F6), 2-segmented (F5, F6), or not clearly








Table 4.1. Morphological and Setal Abbreviations Used.

Ax Axilla
as axilla setae
dl disk length
dw disk width
F1 Funicle (1st segment)
F2-6 Funicle (segments 2-6)
Gs Gaster
gc5 gastral central setae (tergite V)
gc6 gastral central setae (tergite VI)
gc7 gastral central setae (tergite VII)
gll gastral lateral setae (tergites I)
gl2-7 gastral lateral setae (tergites II-VII)
gs genital setae (ovipositor plate)
hb occipital bar
hbl occipital bar setae (central pair)
hb2 occipital bar setae (2nd lateral pair)
ho occipital triangle setae
hp postoccipital bar setae
ISD intersensillar distance
MC Mesoscutal posterior central setae (primary)
mcl mesoscutal setae (1st central pair (=1st longitudinal row)
mclR mesoscutal seta (1st central seta, right side, unpaired)
mcLL mesoscutal seta (1st central seta, left side, unpaired)
mc2-4 mesoscutal setae (2nd-4th pair of central setae)
mdl mesoscutal setae (1st pair in 2nd longitudinal row)
md2-4 mesoscutal setae (2nd-4th pairs in 2nd longitudinal row)
mel-4 mesoscutal setae (3rd longitudinal row)
mfl-4 mesoscutal setae (4th longitudinal row)
ML Mesoscutal anterior lateral setae (primary)
Ms Mesonotum
mf marginal fringe
my marginal vein
mvb marginal vein basal seta
oc occiput
P Pedicel
pob postoccipital bars
PSD Placoid sensillum diameter
R Radicle
S Scape
Sc Scutellum
Sc Scutellar setae (anterior pair)
Sc2 Scutellar setae (distal pair)
sm stemmaticum
smv submarginal vein
smvl submarginal vein setae mesiall pair)
smv2 submarginal vein setae (apical pair)
sv stigmal vein
VII 2nd valvular segment
VIII 3rd valvular segment
vs ventral setae





45
differentiated. Males of some Encarsia species have 7-segmented antennae in

which the F5 and F6 segments are fused together. Funicle segments may bear

several or no linear sensilla and/or short, club-like sensilla. The number of linear

sensilla on each funicle segment is given in sequence for F1-F6, respectively. For

example, (0,0,2,2,2,3) indicates that there are no linear sensilla on F1 and F2, 2

sensilla on F3, F4 and F5, and 3 on F6. At the apical end of F6 there are usually

2 to 3 finger-like sensilla. The male F2 antennal segment may exhibit various kinds

of specialized sensory processes. For example, the F2 antennal segment of E.

niqricephala males has round, pit-like sensorial processes and the males of E.

cibcensis have short arrow-shape processes. The length of the pedicel, by

convention, is measured along the ventral side, since in side view it rarely forms

a perfect isosceles triangle, making its dorsal length usually greater than its ventral

length (Figure 4.4d). Unlike Hayat, I prefer to compare the relative lengths of

antennal segments to the F1 segment because of difficulties in accurately

measuring the pedicel.

The mesoscutum was measured from the center of the anterior margin to

the center of the posterior margin; the width was measured at the level of, but

excluding, the tegulae. The number of setae, setal type and sculpturing of the

mesoscutum are useful characters for distinguishing species. Setal number and

position on the mesoscutum are often designated in the literature in the form of

setal formula. Some ambiguity exists because authors that have used this method

give no mention of how they derived the formula and the same methodology has






46

not been used consistently throughout the literature. An example of a formula

commonly used would be (4+2+2+2), which indicates that there are 4 setae

(including the large pairs of setae in the upper lateral corners of the mesoscutum)

across the anterior margin of the mesoscutum, followed by 3 pairs of central setae.

Alternately, the following method is proposed to name and better pinpoint

the position of a particular seta. Setae are prefixed by the first letter of the body

part on which they are found. Primary setae are designated by capital letters and

secondary setae by small letters. Setae found on the head, mesoscutum,

scutellum, axilla and gaster begin with "h", "m", "s", "a" and "g", respectively (refer

to figure 4.3). With few exceptions, the mesoscutum of Encarsia species

possesses at least the primary setae ML and MC located in the anterior corner

close to the tegulae and the posterior margin, respectively. The secondary

mesoscutal setae are designated as follows: the central setae are designated as

"mc" setae, followed by longitudinal rows of "md", "me" (rarely "mf") setae moving

toward the lateral margins. The setae are numbered 1, 2, 3 or 4, based on their

lateral position. The setae found along the anterior margin or the anterior quarter

of the mesoscutum are designated as "1" setae. The setae found between the

anterior quarter and the middle, between the midline and the posterior three-

quarters, and the posterior three- quarters and the MC setae of the mesoscutum

are designated as 2, 3 and 4, respectively. The name and position of a particular

seta can be designated by using a combination of the longitudinal and lateral

positions. For example, mcl indicates the pair of setae in the central position along






47

the anterior margin of the mesoscutum. The remainder of the secondary setae are

designated following the same scheme for each of the longitudinal rows of setae

(md, me and mf). Unpaired setae can be referred to by adding an "R" (right) or an

"L" (left) to the designated seta. For example mc3R, would indicate an unpaired

central seta on the right side between the midline and posterior three-quarters of

the mesoscutum. For the setal formula, each seta is prefixed in the setal formula

by the number of setae of that type. The formula (2ML;2mc1,2,4+2MC;2md1,2)

indicates one pair of ML setae, three pairs of mc setae plus the MC setae,

followed by 2 pairs of setae adjacent to the central setae.

The scutellum was measured at its maximum length from the center of the

anterior to the center of the posterior margin, and its maximum width across the

transverse midline. The ratio of the intersensillar distance to the placoid sensillum

diameter (ISD:PSD) was determined by dividing the distance between the bases

of the placoid sensillae (ISD) by the width (diameter) of one sensillum (PSD). The

scutellar setae are designated as Sc and Sc2, for the anterior pair and posterior

pair of setae, respectively.

Each axilla always bears one seta in Encarsia, designated as "as1," which

is usually located toward the apex of the axilla but may be found centrally in some

species. The axillae were measured from their anterior to their base and the

position of the axilla seta from its base to the base of the axilla.

All of the legs are 10-segmented (rarely 9), consisting of the coxa,

trochantellus, trochanter, femur, tibia, and 5 (rarely 4) tarsal segments. The stigil,






48
a comb-like structure used for grooming, is present on the foreleg. The tibia of leg

II, referred to as the midtibia or tibia II, and the hindleg bear a saltatorial spur,

which is usually much larger in the former. The coxae of the hindlegs are more

robust than those of the forelegs or midlegs. The tarsi in some species have the

two apical tarsal segments of the midleg fused together (i.e. 4-segmented). Tibia

II was measured from the apex of the femur to the apex of the basitarsus.

Basitarsus II was measured along its longest axis. The endophragma was

measured from the anterior edge of the scutellum to its most posterior point.

The gaster comprises segments II-VIII of the abdomen (excluding the

propodeum, which is the first abdominal segment). The segments of the gaster are

designated, starting with the first segment posterior to the petiole, as tergites 1-VII.

The cercal plates located at the base of tergum VI are useful for orientation. The

gaster may bear one or several lateral setae designated as "gl" setae followed by

1-7, depending on the tergum on which they are found. Central setae are usually

found on tergites V, VI and VII and are designated as gc 1-7, depending upon

which tergum they are located on. One to several rows of ventral setae (vs) are

located between coxa III and the anterior margin of the ovipositor.

The ovipositor may be exserted or not. The ovipositor shaft is composed of

three parts: a pair of serrated stylets (the first valvulae); an unpaired smooth

component (the second valvulae); and the sheaths, or third valvulae (also called

gonostyli) (Figure 4.4e). The ovipositor field or plate is a sunken area on the

venter, demarcated by a ridge running between the first valvulae and the posterior






49

margin of the gaster; genital setae (gs) are usually found in transverse rows and

are designated by a formula that counts the number of pairs of setae in each row

beginning with the most anterior pair of setae. Setae located on the third valvular

segment are designated as "vc" and "vd" for the central and distal setae,

respectively. The ovipositor was measured from its anterior margin (the anterior

edge of the valvular I) to its tip. Valvular II was measured from the same anterior

edge as the ovipositor to the apex of valvular III. The length of valvular III was

measured from its juncture with valvular II to the most posterior point, and the

maximum width was measured across the anterior margin including both papillae

and the ovipositor (Figure 4.4e).

The venation of the forewing is reduced, consisting of a submarginal vein

(smv), marginal vein (mv), stigmal vein (sv), parastigmal vein (pv) (Figure 4.4a).

The parastigmal vein is greatly reduced in Encarsia and bears 1 setae (pv). Just

posterior to the base of the parastigmal vein and along the posterior margin of the

submarginal vein, are the basal setae (bs). The submarginal vein is usually shorter

than the marginal vein and almost always bears only two setae, designated as

smvl and smv2. The marginal vein is equal to or longer than the submarginal vein.

The setae along the anterior border of the marginal vein were designated with the

most mesially-directed seta designated as mvl followed by mv2, mv3, etc. toward

the apex of the forewing.

The length of the forewing was measured from the wing base to the furthest

distal point of its apex; the submarginal vein from the wing base to the pvl seta;





50
the marginal vein from the pvl seta to its apex. The length of the forewing disk,

or discal cell, (dl) was measured by first determining the intercept of the bisect that

divides the forewing into anterior and posterior halves, with a line drawn from the

stigmal vein, running perpendicular to the bisect line; then measuring the line

drawn from the intercept to the apex of the forewing (Figure 4.4a). The disk width

(dw) was measured at the maximum width of the disk.

Figures of new and described species were not drawn to scale, so their

dimensions cannot be compared with those of other drawings. Figures 5.12b,c;

5.15c and 5.24b,d were drawn free-hand by Kurt Ahlmark, Division of Plant

Industry, Gainesville, Florida. Other figures were drawn by the author with the aid

of a drawing tube or modified from those found in the literature. Acronyms used

in the text for holotype depositories following those given by Heppner and Lamas

(1982) are given below.


Table 4.2.

AMEI
BPBM
FSCA
IEUN

MCNG
MNN
NHM
UCR

UNLP
UNP
USNM


Acronyms Used for Holotype Depositories.

American Entomological Institute, Gainesville, Florida, USA.
Bernice P. Bishop Museum, Honolulu, Hawaii, USA.
Florida State Collection of Arthropods, Gainesville, Florida, USA.
Istituto de Entomologia Agraria, Universita degli Studi di Napoli,
Portici, Italy.
Museo Civico di Storia Naturale, Genoa, Italy.
Museo Nacional de Ciencias Naturales, Madrid, Spain.
Natural History Museum (British Museum), London, England.
Division of Biological Control, University of California, Riverside,
California, USA.
Universidad Nacional de La Plata, Argentina.
University degli Studi di Napoli, Portici, Italy.
United States National Museum of Natural History, Washington, DC.





























Okahumpka


Leesburg


-West Palm
Beach


*ompano
Beach

napperr
Creek


homestead


Figure 4.1. Florida Counties and Major Sites Sampled for SPWF Parasitoids.



































RICO
GUADELOUPE'

* GRENADA"


COSTA


Figure 4.2. USA, Caribbean and Latin American Countries Sampled for SPWF
parasitoids.
















































Figure 4.3. Female Encarsia morphology and setal nomenclature.
















































Figure 4.4.


Female Encarsia A) forewing B) antenna with 3-segmented club
C) antenna with 2-segmented club D) pedicel E) ovipositor
F) superior view of head G) frontal view of head.











CHAPTER 5


KEYS TO GENERA, SPECIES GROUPS AND SPECIES



5.1. Introduction

The following taxonomic keys are based on characters found in the adult

female, unless otherwise indicated. Two keys are presented. The first is a key to

the hymenopterous parasitoid genera that attack whiteflies, followed by a brief

discussion of each genus. The second key is to the Encarsia species groups and

a key to the native and exotic Encarsia species parasitizing aleyrodids in the New

World. Encarsia species that parasitize non-aleyrodid hosts are keyed only to the

level of the species group. It is necessary to mount the specimens on microscope

slides and examine them under a compound microscope with at least 100x power

to observe many of the characters used in the key.



5.2. Key to the Parasitoid Genera that Parasitize Aleyrodids



1. Antennae 9 to 11-segmented. Marginal vein lacking or very

short.(Figures5.1 :a,b;5.2:a,b)................................................................... 2

1b. Antennae 8-segmented or less. Marginal vein long and well developed.....3





56
2(1) Antennae 10-segmented, including fused 3-segmented club (Figure 5.1:a).

Marginal and stigmal veins lacking (Figure 5.2:a). Pronotum reaching

tegula (Figure 5.5:a). Midtibial spur short and slender. Male club segments

not fused. Body dark, robust and heavily sclerotized [Proctotrupoidea:

Platygasteridae].................................................................................... Am itus

2b. Female antennae 11-segmented, male antennnae 9-segmented (Figure

5.1:b). Marginal vein very short, about as long as stigmal vein (Figure

5.2:b). Axillae with their apices meeting in the center of the body (Figure

5.4:b). Pronotum separated from tegula. Midtibial spur large and thick.

Body less robust and weakly sclerotized [Encyrtidae].

............................................. .................. ..............................M etaphycus



3(1b). Antennae 3 to 6-segmented, apical segment greatly elongate, usually more

than 4 times as long as the penultimate segment (Figure 5.1 :f,g,j,l,m).......4

3b. Antennae 7 to 8-segmented, apical segment less than two times as long as

penultimate segment (Figure 5.1 :c,d,e,h,k)......................... ............ 6



4(3). Tarsal formula 5-5-5; forewing disk bare or with one seta (Figure 5.2:d),

marginal fringe longer than maximum width of disk. Antennae with F1, F2,

and F3 ring-like (Figure 5.1:j).[Signiphoridae].

............................................. .................. ...............................Si niphora

4b. Tarsal formula 4-4-4; forewing disk setose, at least with several rows of

setae....................................................................................... ............... 5






57
5(4b). Forewing very narrow and with 3-4 rows of discal setae, disk length and

marginal fringe about 2 times as long as maximum disk width (Figure

5.2:e). Antennae 6-segmented, F1 and F2 ring-like. Male antennae 4-

segmented. (Figure 5.1 :m,l) [Aphelinidae]........................................... Cales

5b. Forewing broad and uniformily setose, disk about as long as wide,

marginal fringe less than 0.5 times maximum disk width (Figure 5.2:c).

Female antennae with F1 and F2 short and F3 elongate. Male antennae

3-segmented (Figure 5.1:f,g).[Aphelinidae].................................Eretmocerus



6(3b). Tarsal formula 4-4-4. Antennae 7-segmented, F1 ring-like, very short, more

than twice as wide as long (Figure 5.1:h). Stigmal vein very elongate

(Figure 5.3:c). Known species are dark colored (Figure 5.4:c). [Eulophidae]

.............................................................................................. Euderom phale

6b. Tarsal formula 5-5-5 (rarely 5-4-5). Antennae 7 or 8-segmented, F1 not

ring-like, usually longer than wide. Stigmal vein not very elongate

[Aphelinidae]........................................................................... ............... 6



7(6b). Antenna 7-segmented, often with dark and pale funicle segments, club one

segmented (Figure 5.1:e). Forewing hyaline with dark markings and small

area of long coarse setae under the stigmal vein (Figure 5.2:f). Male

antennae uniformily colored.... ............................. ..........................Azotus






58
7b. Antennae 8-segmented (7-or 8-segmented in male) usually uniform in

color. Forewing setae uniform (lacking small area of coarse setae

underthe stigm al vein............................................................. .............. 8



8(7b). Axillae usually relatively short, mesally separated by a distance greater

than the maximum length of one axilla. Mesoscutum with fewer than 25

setae (usually less than 15) arranged in bilateral symmetry (Figure 5.5:e).

F5 straight, not oblique (Figure 4.4:c). Tarsal formula 5-5-5 rarely 5-4-5.

Two submarginal vein setae, rarely 1 or more than 2....................Encarsia

8b. Axillae long, mesally separated by a distance less than maximum length of

one axilla. Mesoscutum densely setose with more than 25 setae not

arranged in bilateral symmetry. F5 segment oblique or straight (Figure

5.1:c,d). Tarsal formula 5-5-5. Submarginal vein with 2-3 large setae plus

a variable number of smaller setae at the distal end (Figure 5.3:a,b).....9



9(8). Mesoscutum sculpture aciculate (Figure 5.4:g). Intersensillar distance:

placoid diameter ratio less than one. Side lobes divided.................Dirphys

9b. Mesoscutum sculpture imbricate/reticulate, intersensillar distance: placoid

diameter ratio greater than 2. Side lobes not divided (Figure 5.4:h)

.................................................................................................. E ncarsiella







5.3. Biological Characteristics of the Genera

5.3.1. Family Platygasteridae (Proctrupoidea)

Genus Amitus Haldeman 1887

The genus Amitus is cosmopolitan in distribution. There are 11 described

species. However three new species have been discovered during the course of

the Florida survey of Bemisia parasitoids. Females of the new species reared from

Bemisia tabaci in Puerto Rico are parthenogenetic, solitary parasitoids (F. D.

Bennett, personal communication); males occur but are extremely rare. All species

are known only from aleyrodid hosts. According to Flanders (1969), females of A.

hesperidum develop gregariously while males develop as solitary parasitoids. This

parasitoid was instrumental in bringing the citrus blackfly, Aleurocanthus woqlumi,

under control in many areas. Refer to Fouts (1924), MacGowen and Nebeker

(1978) and Viggiani and Mazzone (1982) for descriptions of species.



5.3.2. Eulophidae (Chalcidoidea)

Genus Euderomphale Girault 1916

The genus Euderomphale contains 12 species. Euderomphale species have

been reported primarily from whiteflies of the genus Aleyrodes. However, Viggiani

(1977) described Euderomphale bemisiae from Bemisia citricola. J. LaSalle and

M. Schauff are currently preparing a manuscript on the eulophid genera parasitic

on aleyrodids. They treat six genera, four of them newly erected (M. Schauff,

personal communication).







5.3.3. Encvrtidae (Chalcidoidea)

Genus Metaphycus Mercet 1925

The new Metaphycus species reared from Bemisia tabaci by F. D. Bennett

from Venezuela is the only bonafide whitefly host record for this large genus. Most

species of Metaphycus are parasitoids of soft and armoured scale insects (Noyes

1984). Dr. John Noyes of the Natural History Museum (NHM) confirmed the record

by identifying the pharate (ready to emerge) Metaphycus adults still inside the late

instar Bemisia tabaci pupae as well as individuals that later emerged from the host

material.



5.3.4. Signiphoridae (Chalcidoidea)

Genus Signiphora Ashmead 1880

Woolley (1988) in his review of the phylogeny and classification of the family

Signiphoridae, included 36 described species plus several undescribed species in

the genus Signiphora. Most of the described species are hyperparasites but a few

are primary parasitoids of aleyrodids and diaspine scale insects. Signiphora

aleyrodis (Girault) was reared several times from collections of Bemisia tabaci and

other whiteflies in the survey, but is probably a hyperparasite of Encarsia or

Eretmocerus larvae developing in the same host.



5.3.5. Aphelinidae (Chalcidoidea)

5.3.5.1. Genus Azotus Howard 1898

The genus Azotus is cosmopolitan in distribution and contains around 26





61
described species world-wide (Hayat 1983). Most species develop as

hyperparasites, but occasionally they may develop as primary parasitoids. (Viggiani

1984). Azotus species were not reared from whitefly hosts collected in this survey,

but occasionally were reared from samples containing diaspine scales.



5.3.5.2. Genus Gales Howard 1907

The genus Gales contains 3 species (Viggiani & Carver, 1988), C. noacki

has been the only species reported from the New World; it is a primary parasitoid

of the woolly whitefly, Aleurothrixus floccosus (Maskell) and Aleurotuba ielineki

(Fraunf.).



5.3.5.3. Genus Dirphys Howard 1914

The genus Dirphys contains 4 species all are gregarious, primary

parasitoids of whitefly species of the subfamily Aleurodicinae from the neotropics.

Polaszek and Hayat (1992) in their revision of the genus, described two new

species and provided a cladistical analysis of Dirphys and Encarsiella.



5.3.5.4. Genus Encarsiella Havat 1983

Six species have been assigned to the genus Encarsiella (Polaszek and

Hayat (1992), three of Neotropical and three of Oriental distribution. All but

Encarsiella boswelli (Girault) which parasitizes eggs of Heteroptera, are recorded

as parasitoids of the whitefly subfamily Aleurodicinae. All species are believed to

develop as solitary parasitoids.







5.3.5.5. Genus Eretmocerus Haldeman 1850

The genus Eretmocerus contains about 32 species worldwide. Both males

and females are solitary, endoparasitic, primary parasitoids of aleyrodids. Most

species are arrhenotokous but thelytokous species have been recorded (Gerling

1966). Unlike Encarsia species, eggs are deposited underneath the aleyrodid

nymph and no meconium is cast at the end of larval development. Eretmocerus

species often constitute a large percentage of the parasitoid complex of an area.

Eretmocerus mundus Mercet was introduced into Florida from Israel and Sudan

for the biological control of Bemisia tabaci. Like Encarsia, species identification is

complicated by limited knowledge of genus, especially of the New World fauna.

Eretmocerus is currently being revised by Michael Rose of Texas A & M

University. (See Compere (1936) and Hayat (1972) for species descriptions and

taxonomic keys).



5.4. Key to Encarsia Species Groups of the World and Encarsia

Species that Parasitize Aleyrodids in the New World.



1. Tarsal formula 5-4-5 (Figure 5.7:a;5.8:h,k)...................................... ...2

lb. Tarsal form ula 5-5-5.(Figure 5.13:c,f,i).......................................................



2(1). Forewing with a large, round asetose (bare) area under stigmal

vein (Figure 5.6:a,b,c). Mesoscutum usually with 2 to 3 pairs of setae.

Male F5 and F6 of segments fused (Figure 5.7: c,f).(Cubensis Group)..15






63
2b. Forewing uniformily setose (Figure 5.8:a). Mesoscutum usually with more

than 4 pairs of setae. Club usually 2-segmented. F5 and F6 segments

fused or not.(Formosa Group)...............................................................17



3(1b). Forewing narrow, disk more than 1.5 times as long as wide. Marginal

fringe long, at least 0.6 as long as maximum disk width (Figure

5.11:e;5.25:b,e). Mesoscutum with 0 to 4 pairs of setae (usually

2)........ ......... ....................................................................... ....... ......4

3b. Forewing moderate to broad, area under stigmal vein setose, marginal

fringe variable but usually less than maximum width of disk (Figure

5.17b). Number mesoscutal variable, but usually more than 4 pairs.........6



4(3) Forewing with large, round asetose area under the stigmal vein. Petiole

striated or not (Figure 5.11:e). Parasitoids of aleyrodids and diaspine

scales................................................................................... ............... 5

4b. Forewing uniformily setose. Petiole rarely striated (Figure 5.26:h,i).

Parasitoids of diaspine scales.........................................(Inuirenda Group)



5(4). Petiole with striate lines. Forewing often sharply inflexed (angled)

posteriorly. Valvular III gonostyli short and thick, diaspine scales

parasitoids (Figure 5.25:a,b,c).............................................(Citrina Group)





64

5b. Petiole smooth. Forewing not sharply inflexed. Valvular III gonostyli long

and narrow (Figure 5.11). Parasitoids of aleyrodids....(Parvella Group).24



6(3b). Scutellum with intersensillar distance less than diameter of one sensillum

(Figure 5.13). F1 usually cylindrical at least 1.5 times as long as wide and

0.75 as long as F2........................................................ (Strenua Group).7

6b. Scutellum with intersensillar distance greater than the width of one

sensillum. F1 length variable in length............................................ 9



7(6). Club 3-segmented. Gaster orange or yellow (dark in bella). Forewing

hyaline (Figure 5.13)............................................................................ ... .8

7b. Club 2-segmented, gaster usually dark brown or black. Forewing

infuscate (Figure 5.16)...................................... (Quercicola Subgroup).31



8(7). Body yellow to orange (some tergites may have dark transverse bands.

Placoid sensillae kidney shaped. Ovipositor often long and/or extruded.

Parasitoids of aleyrodids (Figure 5.13)....................(Strenua Subgroup).27

8b. Body dark brown. Placoid sensillae round. Ovipositor moderate in length.

Parasitoids of diaspine scales (Figure 5.26:a,b,c,d) .........(Bella Subgroup)



9(6b) Pedicel distinctly longer than F1 (Figure 5.18:a,i)....................................10

9b. Pedicel subequal in length to, or shorter than F1 (Figure 5.21 :c)............13





65

10(9). Ovipositor short to moderate (less than 0.8 times as long as midtibia) and

valvular III short to moderate. Conical setae absent on basitarsus II

(Figure 5.23:c)....................................................................................... 11

10b. Ovipositor long (equal to or longer than midtibia) and valvular III long.

Conical setae present on basitarsi of many species (Figure 5.18:g,k)

................................................................................. (O pulenta G roup).34



11(10). Valvular III dark brown, in sharp contrast to pale ovipositor plate (Figure

F1 quadrate, shorter than F2. Marginal fringe about 0.33 times as long

as maximum width of disk. Male with F1-3 laterally expanded, and F5

and F6 fused (Figure 5.23:a-g)............................................(Lutea Group)

Only one species reported in the New World..........................lutea (Masi)

11b. Valvular III pale. F1 cylindrical subequal to F2. Length of marginal fringe

variable. Male F1-3 not laterally expanded.............................................12



12(11b). Ovipositor plate quadrate (Figure 5.25:f), mostly diaspine scale

parasitoids...........................................................................(Aurantii G roup)

One species parasitizing New World aleyrodids. Body dark brown to

black. Marginal fringe short about 0.10 times as long as maximum width

of disk (Figure 5.22:a-e)..................................................eltata (Cockerell)

12b. Ovipositor plate rectangular, aleyrodid parasitoids. (Figure 5.23:k,l)

.........................................................................................(PorteriG roup).43





66
13(9b). F1 longerthan F2................................................................................ 40

13b. F1 equal to or shorter than F2, funicle segments subequal in length......14



14(13b). Club 3-segmented. Maxillary palps 2-segmented. Male with large oval

sensorial process on F2 antennal segment (Figure 5.22:f-i)

(Plaumanni Group). One species, mesoscutum imbricate with 14-16 pairs

of setae........................................................................... laum anni Viggiani

14b. Club 2-segmented. Maxillary palps 1-segmented. Males lacking oval

sensorial process. Mesoscutum with less than 10 pairs of setae.

(Figure 5.20)........ ........................................................(Inaron Group).42



15(2). Gaster, axillae and scutellum pale. Head and mesoscutum dark brown

with posterior 0.25 to 0.50 of mesoscutum pale and with two pairs of

setae. Male F2 antennal segment longer than F3 (Figure 5.6:b;5.7:c,d,h)

................................................ .................................niqricephala D ozier

15b. Gaster, axillae, head and mesoscutum dark brown. Scutellum pale to

bright yellow, mesoscutum with 2 to 4 pairs of setae.............................16



16(15b). F1 short, about 0.5 times as long as F2. Mesoscutum with 2 pairs of

setae (Figure 5.6:a;5.7:b,g). Males unknown..................cubensis Gahan

16b. F1 and F2 subequal in length. Mesoscutum with 3 pairs of setae. Male

F2 shorter than F3 segment (Figure 5.6:c;5.7:a,e,f,i)....quaintancei Howard





67

17(2b). Entire body yellow to orange (sometimes dusky)...................................18

17b. At least head and thorax dark brown to black, gaster variable................19



18(17). F1 about 0.5 times as long as F2 (Figure 5.8:1). Hind femur fuscous.

Marginal fringe about 0.10 as long as maximum disk width.

............................................................................................haitiensis Dozier

18b. F1 length slightly shorter than F2. Hind femur pale. Marginal fringe more

than 0.2 times as long as maximum width of disk (Figure 5.28:i-k).

.......................................................................................m eritoria G ahan

(E. gallardoi Marelli is indistinguishable from E. meritoria based on

Marelli (1933) description and figures).

19(17b). Gaster dark brown or black, not banded..........................................20

19b. Gaster pale or sometimes banded....................................................... 21



20(19). Club 3-segmented and distinctly clavate. Mesoscutum with 4 pairs of

setae. Marginal vein short about 0.5 times as long as disk. Ovipositor as

long as hind tibia (Figure 5.8:a,b)......................................brunnea Howard

20b. Club 2-segmented (not distinctly clavate). Mesoscutum with 9 pairs of

setae. Marginal vein about equal to disk length. Ovipositor 1.14 as long

as midtibia (Figure 5.9:a-e)........................................uadeloupae Viggiani



21(19b). Gaster with brown transverse or lateral bands. Midtibial spur long, about

equal to corresponding basitarsus. F1 short less than 1.5 times as long





68
as wide (Figure 5.5:k).............................................................................22

21b. Gaster pale, midtibial spur short, only 0.50 to 0.75 as long as basitarsus

II. F1 elongate at least two times as long as wide (Figure 5.8:h)............23



22(21). F1 about 0.5 times as long as F2. Lateral margin of tergites I-VII with

dark brown markings. Head and mesoscutum dark brown. Scutellum pale

with anterior margin dark brown. Forewing disk broad with rounded apex

(Figure 5.10:a)................................................................ varie ata Howard

22b. F1 only slightly shorter than F2. Tergites V and VI with brown transverse

bands. Head, mesoscutum and scutellum lighter brown, midtibial spur

longer and forewing disk not as broad or apex as rounded as in above

species (Figure 5.8:c-e).................................................desantisi Viggiani



23(21b). Head and thorax dark brown, gaster bright yellow. Linear sensillae

present on F1 and F2. Interior of mesoscutum and axillae areolate rather

smooth. Males rare and with scutellum dark brown (Figure 5.8:f,g).

.........................................................................................form osa G ahan

23b. Head and thorax light brown to dark orange, gaster pale yellow. Linear

sensillae absent on F1 and F2. Interior of mesoscutum and axillae

areolae with rugose lines. Males common and with light brown scutellum

(Figure 5.10:b,c).................................................................... uteola Howard





69
24(5b). Mesoscutum with inverted brownish triangle spot and with more than 3

to 4 pairs of setae (Figure 5.11 :c-e).......................... ercqandiella Howard

24b. Mesoscutum color uniform with 2 pairs of setae....................................25



25(24b). Base of gaster with prominent, dark transverse band. F1 short about

equal to 0.5 times the length of F2 (Figure 5.11 :f-h)........basicincta Gahan

25b. Gaster uniform in color. F1 and F2 subequal in length..........................26



26(25b). Valvular III apical setae lanceolate. Forewing setae very sparse (large

bare areas on anterior and posterior margins) (Figure 5.12).

..................................................................................lanceolata spec. nov.

26b. Valvular III apical setae slender. Forewing setae more profuse (Figure

5.11 :a,b).. .................................................americana DeBach and Rose



27(8). Forewing with infuscate band under the marginal vein. Tergites Ill-V with

transverse bands or spot (Figure 5.14:a).........................................28

27b. Forewing hyaline. Gaster yellow or orange, not banded........................29



28(27). Transverse bands of gaster very well defined and reaching lateral

margins. Funicle segments increasing in length. Valvular III moderate in

length about two times as long as wide, basitarsus II very short less than

length of midtibial spur (Figure 5.14a-e)...........................citrella (Howard)

28b. Dark spot in center of gaster with margins not well defined. Funicle





70

segments subequal in length. Valvular III very short, about as long as

wide. Basitarsus II moderate length longerthan tibial spur (Figure 5.15:a-d)

.......................................................................pseudocitrella spec. nov.



29(27b). Forewing posterior margin with small area of long coarse setae. Occiput

rugose. F6 elongate longer than F4 or F5, conical setae on basitarsus II

absent. Ovipositor moderate in length, less than or equal to length of

midtibia, syntergum not elongate. Five basal setae not extending mesially

beyond smv 2 setae. Midtibial spur distinctly shorter than basitarsus II

(Figure 5.13:g-j)......................................................... transvena (Timberlake)

29b. Forewing setae uniform (without area of long coarse setae). Occiput

reticulate. F6 shorter than F4 or F5. Row of conical setae on basitarsus

II (Figure 5.13:c,f). Ovipositor much longer than midtibia and syntergum

elongate. Forewing with 7-8 basal setae extending mesially beyond smv2

setae. Midtibial spur about equal to length of basitarsus II....................31



30(29b). Ovipositor more than 1.7 times as long as midtibia. F6 conical about

equal to F5 with F4 longer than F5. Basitarsus II with 2-3 conical setae,

axilla setae elongate reaching almost to base of axilla (Figure 5.13:

a-c)...................................................................................arm ata (Silvestri)

30b. Ovipositor less than 1.7 times as long as midtibia, F6 short and stubby

with F5 longer than F4. Axilla setae shorter than in former species, not

reaching axilla base. Basitarsus II with row of 5 large conical setae.






71

(Figure 5.13:d-f)............................................................. strenua (Silvestri)

31(7b). F1 elongate, about 3 times as long as wide, longer than pedicel and

subequal to other funicle segments. Thorax yellow (Figure 5.16:

e-f).......................................................................... lycopersici De Santis

31b. F1 short, less than 2 times as long as wide and shorter than pedicel,

color variable........................................................................................... 32



32(31b). Forewing hyaline, funicle uniform in color F4 shorter than F5. Midtibial

spur about equal to basitarsus II. Five pairs of mesoscutal setae.

Ovipositor robust originating at base of the gaster (Figure 5.16:a-c).

.................................................................................portoricensis How ard

32b Forewing infuscate, F5 and/or F6 brown, contrasting with yellow

F 1-F4 .................................................................... ................................ 33



33(32). Mesoscutum dark brown. F1 cylindrical, subequal to length of pedicel.

Antennae yellow with scape and F6 antennal segments brown. Hind

coxae and femur dark brown. Axilla setae located centrally. Ovipositor

shorter than midtibia (Figure 5.16:g-i)........................ uercicola (Howard)

33b. Mesoscutum orange. Antennae yellow, F5 and F6 brown, pedicel short,

less than length of F1. Axilla setae at apex of axilla. Hind coxae pale and

ovipositor longerthan midtibia (Figure 5.17:f-h)............catherinae Dozier







34(10b). Gaster completely brown to dark brown (tergum VII may be lighter),

forewing hyaline (except in smithi)........................................................35

34b. Gaster not completely brown, at least one other tergum yellow. Forewing

infuscate............................................................................................. 37



35(34). F1 and F2 short, quadrate and subequal in length, each about 0.5 times

as long as F3. Tergum VI with spiracular plates distinctly separated.

Head and thorax yellow with axillae dark brown (Figure 5.5.19:a-d).

..........................................................................................m erceti Silvestri

35b. At least F2 cylindrical more than 0.5 times as long as F3. Tergum VI

spiracular plates not separated. Color variable......................................36



36(35b) F1 usually wider than long and less than 0.5 times as long as F2.

Forewing hyaline. Axilla setae locate centrally. Syntergum elongate,

ovipositor strongly exserted............................................................... 37

36b. F1 longer than wide, and more than 0.5 times as long as F2. Forewing

infuscate. Syntergum not elongate and ovipositor not strongly exserted.

(Figure 5.19:e,f)...................................................................sm ithi (Silvestri)



37(34b). Ovipositor as long as midtibia. Marginal fringe about 0.25 times as long

as maximum disk width. Stigmal vein with narrow neck. Ovipositor not

extruded (Figure 5.18:c-f)...............................................diverens (Silvestri)






73

37b. Ovipositor longer than midtibia. Marginal fringe less than 0.20 times as

long as maximum disk width. Stigmal vein neck not narrow. Syntergum

elongate and ovipositor extruded.......................... ...........................39

38(37b). Clypeus with triangular projection. Mesoscutum with 10 setae. Basal half

of mid femur and entire hind femur dark brown. Sc setae relatively short,

not reaching base of Sc2 setae (Figure 5.19:g-1)........... clypealis (Silvestri)

38b. Clypeus lacking triangular projection. Mesoscutum with 8 setae. Femur

II and III orangish. Sc 1 setae very long and stout, reaching the base of

Sc 2 setae (Figure 5.18:a,b)..........................................townsendi Howard



39(37b). Gaster dark brown with tergum VII (sometimes I and II) yellow.

Mesoscutum with 10 pairs of setae and small hexagonal areolae in a

spiral design. Valvular III yellow with distal half dark brown. Midtibial spur

large about as long as basitarsus II (Figure 5.18:g-k)....opulenta (Silvestri)

39b. Gaster dark brown with tergum IV-VI dark, other characters similar to

opulenta (Figure 5.17:a-e)..........................................brasiliensis (Hempel)



40(13). Mesoscutum with two pairs of setae and areolae arranged in a dome-

shaped pattern. Sc setae minute. Forewing uniformily setose with bare

strip along the posterior margin (Figure 5.21 :a-d)........lahorensis (Howard)

40b. Mesoscutum very setose (11 pairs of setae) with areolae in longitudinal

rows. Sc1 setae long. Forewing setae underthe marginal vein longer and

coarser than disk setae............................................. ciliata (Gahan)





74

41(14b) Gaster entirely dark, hind femur with dark brown band in the middle.

Ovipositor subequal in length of midtibia. Axilla setae large and stout.

(Figure 5.20:f-i)...............................................................coquilletti Howard

41 b. Gaster pale (sometimes banded), hind femur without dark brown band.

Ovipositor shorter than midtibia. Axilla setae not particularly large and

stout........................................................................................................ 42



42(41b) Gaster pale (may have transverse bands on tergum V and VI), head,

mesoscutum and scutellum dark brown. Tergites II-IV imbricated with 1

pair of lateral setae, all funicle segments subequal in length (Figure 5.20:

a-c)....................................................................................... aron (W alker)

42b. Gaster yellowish with central area of tergites IV-VII brownish. Tergites II-

IV imbricate with 3 pairs of setae. Mesoscutum brown with lateral margins

yellowish. F2 and F4 longer than F3 and F5, respectively (Figure 5.20:

d-f)..................................................................................... o ezi Blanchard



43(12b) Body yellow. Basitarsus II elongate about 6-7 times as long as wide.

(Figure 5.23:k-m).. ............................................................orteri (Mercet)

43b. Body yellow, with central region of mesoscutum with large, round dark

brown spot, lateral margins of gaster dark brown (Figure 5.24:a-e)

................................................................................. polaszeki spec. nov.














































Figure 5.1.


Parasitoid Antennae. A) ? Amitus B) ? Metaphycus C) ? Encarsiella
D) ? Dirphys E) ? Azotus F) 9 Eretmocerus G) c" Eretmocerus
H) V Euderomphale I) q Coccophaqus J) Siqniphora
K) 9 Coccophaqoides L) ci Gales M) ? Gales.






76



_--
A .














E

'^-^ ^ ^\ *r ^ 1 / '- -- -_"- -^^ ^ -4
F IN


































Figure 5.2. Forewing 9 A) Amitus B) Metaphycus C) Eretmocerus
D) Sigqniphora E) Cales F) Azotus.
~ ... .- -















-l -t=- -

'C' XX
Figure~~~~~~~~~~I 52 Foe Ing~A mtsB eahcsC rtoeu
D)Siiipor ) aesF)Aots






77
















z.~~~~~~--, ',-,,;"-"- ---_---- .

-----------. ;.-.~ ''--. ~-.,;,.4i'-.._- .- -._-_-_ .--. -
I










CC) "

,, .; -.%_ __....
." -. -
AI .-. : .


































Figure 5.3. Forewings 9. A) Dirphys B) Encarsiella C) Euderomphale
D) Pteroptrix E) .Coccopha~Qoides F) Coccophapus.

































G







Figure 5.4.


SThorax. A) Amitus B) Metaphycus C) Euderomphale
D) Eretmocerus E) Signiphora F) Gales
G) Dirphys H) Encarsiella















































Figure 5.5.


9 Thorax. A) Azotus B) Coccophaqoides C) Coccophaqus
D) Pteroptrix E) Encarsia.




80














^- //,./_ \\ I^j7/\




t <
iii



























Figure 5.6. t Forewing and Mesothorax. A) cubensis B) niqricephala
C) quaintancei.
-\ N\









NB K
r- ~
~ IJ .

NI~ '
I
\x~




= ~~ ~ ~ ~~~ /';S)~~~l;;~


Fiue5..7 Freigan eohoa.A cueni 8)\V~ nticieh





Figure C gua 9Foeint ancei. hoa.A cbnisB irieh






































G HI






Figure 5.7. A) quaintancei ? midtarsus. B-G Antennae. B) 8 cubensis
C) d, niqricephala D) ? niqricephala E) 8 quaintancei
F) d' quaintancei. G-I ovipositor. G) cubensis H) niqricephala
I) quaintancei.






























//-

I'''


14


L h


E. brunnea A) 9 forewing B) -9 antenna. desantisi C) !? antenna D)
cP antenna E) V forewing. formosa F) ?. antenna G) cr antenna H) !?
tarsus 11. meritoria 1) !? antenna J) !? forewing K)!? tarsus 11. haitiensis_
L) 9 antenna.


~c~
A .
.1.
z


Figure 5.8.


1












































D



^ ^ /'
I-.. -


Figure 5.9.


Female. Encarsia quadeloupae A) thorax B) ovipositor C) forewing
D) stigmal vein E) antenna

















































Figure 5.10. Female. Encarsia variegata A) antenna. luteola B) thorax C) antenna















































Figure 5.11. Female. Encarsia americana A) thorax B) antenna. perqandiella
G) antenna D) thorax E) forewing,basicincta F) antenna G) ovipositor
H) F6 antennal apex.












--- -- ---'--



,, l .' --
/ I












D







Figure 5.12. Female. Encarsia lanceolata A) thorax B) ovipositor C) antenna D)
forewing.















































Figure 5.13.


Encarsia armata A) ? habitus B) 8 antenna C) ? tarsus II. strenua
D)._ habitus E) ? antenna F) ? tarsus II. transvena G) 9 antenna
H) d, antenna 1) 9 tarsus II J) 9 thorax















































Figure 5.14. Female. Encarsia citrella. A) habitus B) forewing C) antenna
D) ovipositor E) thorax F) tarsus II.















B /


I / I

I ~ -, -
/ I -
,I I I ----_

I' ~ --
/I ',


Figure 5.15. Female. Encarsia pseudocitrella. A) thorax B) ovipositor C) antenna
D) forewing.


---_(



























































































Figure 5.16. Encarsia portoricensis A) habitus B) forewing C) antenna. lycopersici

D) forewing E) ? antenna F) dc antenna. quercicola G) thorax

H) forewing I) antenna.


~-'r=; :

-=----;
,, j '- --
'
,,.-.~ ~i .'----I-~--:-~--.-- -~- F-

--
B L- ----- ----,-,-
-i---
---





,,












A C




;\ /

If D
E ^X^j J
E H




\G
IF







Figure 5.17. Female. Encarsia brasiliensis A) habitus B) forewing C) antenna
D) valvular III E) tarsus II. catherinae F) forewing G) tarsus II H) antenna.