ArticlePDF Available

Review of parasitoid wasps and flies (Hymenoptera, Diptera) associated with Limacodidae (Lepidoptera) in North America, with a key to genera.



Content may be subject to copyright.
BioOne sees sustainable scholarly publishing as an inherently collaborative enterprise connecting authors, nonprofit
publishers, academic institutions, research libraries, and research funders in the common goal of maximizing access to
critical research.
Review of Parasitoid Wasps and Flies (Hymenoptera, Diptera)
Associated with Limacodidae (Lepidoptera) in North America,
with a Key to Genera
Author(s) :Michael W. Gates, John T. Lill, Robert R. Kula, James E. O'Hara,
David B. Wahl, David R. Smith, James B. Whitfield, Shannon M. Murphy and
Teresa M. Stoepler
Source: Proceedings of the Entomological Society of Washington, 114(1):24-110.
Published By: Entomological Society of Washington
BioOne ( is a nonprofit, online aggregation of core research in the
biological, ecological, and environmental sciences. BioOne provides a sustainable online
platform for over 170 journals and books published by nonprofit societies, associations,
museums, institutions, and presses.
Your use of this PDF, the BioOne Web site, and all posted and associated content
indicates your acceptance of BioOne’s Terms of Use, available at
Usage of BioOne content is strictly limited to personal, educational, and non-commercial
use. Commercial inquiries or rights and permissions requests should be directed to the
individual publisher as copyright holder.
114(1), 2012, pp. 24–110
W. G
T. L
R. K
E. O’H
B. W
R. S
B. W
M. M
M. S
(MWG, RRK, DRS) Systematic Entomology Laboratory, USDA, ARS, PSI, c/o
National Museum of Natural History, Washington, DC 20013-7012, U.S.A. (e-mail:
MWG, RRK, DRS dave.; (JTL, TMS) The George Washington University, Department
of Biological Sciences, 2023 G Street, NW, Suite 340, Washington, DC 20052, U.S.A.
(e-mail: JTL, TMS; (JEO) Canadian
National Collection of Insects, Agriculture and Agri-Food Canada, 960 Carling
Avenue, Ottawa, Ontario, Canada K1A 0C6 (e-mail:;
(DBW) American Entomological Institute, 3005 SW 56
Ave., Gainesville, Florida
32608 U.S.A. (e-mail:; (JBW) Department of Entomology,
University of Illinois, Urbana-Champaign, Illinois 61801, U.S.A. (e-mail: jwhitfie@; (SMM) Department of Biological Sciences, University of Denver,
F. W. Olin Hall, 2190 E. Iliff Ave., Denver, Colorado 80208, U.S.A. (e-mail: Shannon.
Abstract.—Hymenopteran and dipteran parasitoids of slug moth caterpillars
(Lepidoptera: Limacodidae) from North America are reviewed, and an illustrated
key to 23 genera is presented. Limacodid surveys and rearing were conducted during
the summer months of 2004–2009 as part of research on the ecology and natural
history of Limacodidae in the mid-Atlantic region of the U.S.A. Parasitoid rearing
involved a combination of collecting naturally occurring larvae in the field (at least
14 host species) and placing out large numbers of “sentinel” larvae derived from
laboratory colonies of three host species. Species in the following families are
documented from limacodids in North America as primary or secondary parasitoids
(number of genera for each family in parentheses; number of genera included in key
but not reared through this research in brackets): Chalcididae ([1]; Hymenoptera:
Chalcidoidea), Eulophidae (3; Chalcidoidea), Pteromalidae ([1]; Chalcidoidea),
Trichogrammatidae (1; Chalcidoidea), Braconidae (3 [1]; Hymenoptera: Ichneumo-
noidea), Ichneumonidae (7 [3]; Ichneumonoidea), Ceraphronidae (1; Hymenoptera:
Ceraphronoidea), Trigonalidae (2; Hymenoptera: Trigonaloidea), Bombyliidae ([1];
Diptera: Asilioidea), and Tachinidae (3; Oestroidea). We recovered 20 of 28 genera
known to attack limacodids in North America. Records discerned through rearing in
the mid-Atlantic region are augmented with previously published host-parasitoid
relationships for Limacodidae in North America north of Mexico. New records are
reported for the following parasitoids (total new records in parentheses): Uramya li-
macodis (Walker) (1), U. pristis (Townsend) (5), Austrophorocera spp. (6), Ceraphron
sp. (1), Alveoplectrus lilli Gates (1), Playplectrus americana (Girault) (10), Pediobi u s
crassicornis (Thomson) (1), Trichogramma (1), Mesochorus discitergus (Say) (1),
Hyposoter fugitivus (Say) (1), and Isdromas lycaenae (Howard) (5). The male of
Platyplectrus americana (Hymenoptera: Eulophidae) is redescribed, and the fe-
male is described for the first time. Incidental and miscellaneous host-parasitoid
associations are discussed, and it is concluded that most of these records are likely
parasitoids of contaminants accidentally introduced during the limacodid rearing
process. Triraphis eupoeyiae (Ashmead), new combination, is transferred from
Rogas (Hymenoptera: Braconidae).
Key Words: hyperparasitoid, parasitic, slug moth caterpillar, Acharia,Acrolyta,
Orthogonalys,Packardia,Par a s a,Pediobius,Phobetron,Platyplectrus,
Slug moth caterpillars (Lepidoptera:
Limacodidae) are mostly polyphagous
external foliage feeders on a broad array
of deciduous trees and shrubs in eastern
North America (Wagner 2005, Lill 2008).
Their broad diets, rather distinctive larval
morphologies, and often bright coloration
make them interesting subjects for eco-
logical and behavioral studies (e.g., Lill
et al. 2006, Murphy et al. 2010). Unlike
many more mobile caterpillars, limaco-
did larvae tend to remain on an individual
host plant for the duration of their de-
velopment: typically about two months,
going through numerous instars (7–12)
depending on the species, with some
species requiring more than 100 days
to complete development (J. Lill and
S. Murphy pers. obs.). Larvae tend to
occur in low densities (<1 caterpillar/
foliage; Lill et al. 2006, Lill 2008)
and can be highly cryptic, especially in
early instars, but they produce a rather
distinctive pattern of feeding damage
that facilitates their capture. Outbreaks
of North American limacodids are re-
ported rarely, but many tropical species
are important agricultural pests, partic-
ularly in banana and palm plantations
in South America and Southeast Asia
(Ostmark 1974, Cock et al. 1987, Godfray
and Chan 1990). Because many species
of Limacodidae possess stinging spines
during all or a portion of their larval de-
be a nuisance when they occur in planta-
tions (Ostmark 1974) or on ornamental
palms near tourist attractions (e.g., Conant
et al. 2002). Their protracted larval de-
velopment period exposes the caterpillars
to attack from a diverse assemblage of
larval and larval-pupal parasitoids. How-
ever, research to-date on host-parasitoid
interactions in Limacodidae has focused
almost entirely on biocontrol of pest li-
macodids in tropical agricultural systems
(e.g., Cock et al. 1978) with the notable
exception of the massive caterpillar rear-
ing database compiled for tropical forests
of Area de Conservacio
´n Guanacaste in
Costa Rica (Janzen and Hallwachs 2009).
*Accepted by John W. Brown
DOI: 10.4289/0013-8797.114.1.24
By contrast, the hymenopteran/dipteran
parasitoid community that attacks North
American limacodids has been described
only anecdotally, and concerted rearing
efforts are mostly lacking (but see Le
Corff and Marquis 1999 and Stireman
and Singer 2003a, b for examples where
limacodids were reared as part of larger
community sampling).
In general, dipteran (i.e., Tachinidae)
and hymenopteran parasitoids are essen-
tial in regulating native forest macrolep-
idoptera and are reasonably well studied
(LaSalle 1993, McCullough et al. 1999).
Conversely, there is little information
regarding bombyliid parasitoids (i.e., in
Systropus) and their roles in regulating
native forest macrolepidoptera, likely
due to their relative rarity compared to
parasitic hymenopterans and tachinids.
Most hymenopteran parasitoids of mac-
rolepidoptera belong to the superfamilies
Ichneumonoidea and Chalcidoidea, and
species of Tachinidae are the most com-
mon dipteran parasitoids of microlepi-
doptera (Askew 1971, Quicke 1997). Two
recent studies in North America have
focused on Tachinidae and Hymenoptera
specifically and their host relationships in
eastern North American forests (Strazanac
et al. 2001, Petrice et al. 2004). Most
research on parasitoids of native forest
macrolepidoptera has focused on out-
break species such as fall webworm
(Hyphantria cunea Drury; Arctiidae),
eastern tent caterpillar (Malacosoma
americanum (F.); Lasiocampidae) (Kulman
1965, Morris 1976, Witter and Kulman
1979), and gypsy moth (Lymantria
dispar (L.); Lymantriidae) (Barbosa et al.
1975, Elkinton and Liebhold 1990 and
references therein). Few studies have
documented parasitoids in less abundant/
non-outbreak forest macrolepidoptera (in-
cluding limacodids) (Schaffner and Griswold
1934, northeasternU.S.A.; Raizenne 1952,
Ontario, Canada), and fewer still have
focused on Limacodidae (as mentioned
Tachinidae and parasitoid Hymenoptera
are diverse in forest canopy communi-
ties in North America with new host-
parasitoid associations being recorded with
some frequency (Butler 1993, Strazanac
et al. 2001, Petrice et al. 2004). Bio-
logically, tachinids are recorded from
15 arthropod orders (Wood 1987), whereas
parasitoid Hymenoptera have been docu-
mented from 16 (LaSalle 1993). As a
group, parasitoid Diptera have been
viewed as secondary to hymenopterans
as effective parasitoids (Askew 1971,
Quicke 1997); however, both groups
are abundant in North American forests
where limacodids occur most commonly.
North American hymenopteran and dip-
teran parasitoid-host associations have
been cataloged by Krombein et al. (1979)
and Arnaud (1978), respectively. A world
catalog to Tachinidae is in preparation
(JEO), and databases are available for
Ichneumonoidea (Yu et al. 2005) and
Chalcidoidea (Noyes 2003). Recent large-
scale rearings of forest macrolepidoptera
by Butler (1993) documented 115 new
hymenopteran host associations, and
Strazanac et al. (2001) reported 60 new
tachinid host associations.
Many of the tachinid and hymenop-
teran parasitoids reported herein are
known to attack lepidopterans (or their
primary parasitoids) more generally, but
several specialize on Limacodidae, and
others are facultative (see Pediobius;
Peck 1985) or obligatory hyperparasitoids
(see Conura Spinola; Delvare 1992).
Given their specialization on Limacodidae,
many of these taxa, particularly certain
Chalcidoidea such as Platyplectrus Ferrie
and Alveoplectrus Wijesekara and Schauff,
were encountered infrequently in North
America (Wijesekara and Schauff 1997).
This can be ascribed to the lack of major
limacodid pests in North America, the
difficulty of locating/handling their larvae,
and the lack of financial resources avail-
able to study non-outbreak pests. Most
of the parasitoids documented as a re-
sult of the rearing portion of this study
parasitize early limacodid instars.
The purpose of this study is to review
the parasitoids associated with limacodids
in America north of Mexico, including
introduced limacodids (i.e., Monema
flavescens Walker) and exotic parasitoids
recorded from limacodids elsewhere that
could potentially attack limacodids in
North America. We assemble these vari-
ous records herein and add host-parasitoid
associations generated from six years of
intensive rearing of larval limacodids in
the greater Washington, DC metropolitan
area (Table 1).
Collection and Rearing
Beginning in mid- to late-June of each
summer of 2004–2009, we searched
manually for limacodid larvae on the
undersides of leaves of common woody
trees at the following field sites near
Washington, DC: Little Bennett Regional
Park, Plummers Island (Montgomery
County, MD), Patuxent National Wildlife
Refuge (Prince George’s County, MD),
Rock Creek Park (Washington, DC), and
the U.S. National Arboretum (Washington,
DC). More than a dozen host plants have
yielded limacodids, but most of our efforts
were focused on searching six common
host plants used by most species of
Limacodidae: American beech (Fagus
grandifolia Ehrh.), white oak (Quercus
alba L.), northern red oak (Quercus rubra
L.), black gum (Nyssa sylvatica Marsh.),
black cherry (Prunus serotina Ehrh.), and
pignut hickory (Carya glabra Miller).
Additional host plant species were sam-
pled less intensively (Table 1). For each
wild-caught larva, we recorded the species,
collection date, and host plant. In addition
to these wild-caught larvae, we also con-
ducted a series of field experiments as part
of a different project examining tri-trophic
interactions in Limacodidae that involved
placing out “sentinel” larvae on each of the
six common host plants described above
at the Little Bennett Regional Park site.
These larvae came from laboratory colo-
nies established through a combination of
larval and adult (ex ovo) collections; thus,
larvae were unparasitized when placed in
the field. These experimental larvae were
left exposed to parasitoid attack in the field
for one to several weeks depending on the
experiment and then brought back to the
lab for rearing.
Collected larvae were reared individu-
ally in 16 oz. clear plastic deli containers
containing a disk of moistened filter paper
to prevent host foliage from drying out.
Fresh leaves from the various host plants
were replaced as needed (typically twice
per week). Larvae showing signs of para-
sitism were checked routinely for parasit-
oid larvae/pupae, and emerging adults were
either placed in 95% ethanol (Hymenop-
tera) or frozen (Diptera) for later mounting/
pinning and identification. One genus of
tachinid flies (Austrophorocera spp.) exclu-
sively contains larval-pupal parasitoids, so
adults eclosed from overwintering cocoons
the following year (pupae were kept in an
environmental chamber in moist peat moss
during the winter months and then exposed
to spring conditions to induce fly pupation
and emergence). In addition, several of
the hymenopteran parasitoids reared from
limacodid hosts late in the season dia-
paused as pupae and emerged the follow-
ing summer.
Hymenopteran Parasitoid
Preparation and Imaging
Specimens in ethanol were dehydrated
through increasing concentrations of
Table 1. Parasitoids reared from host plant/limacodid host pairs in greater Washington, D. C., 2004–2009.
Limacodid Species Plant Species Parasitoid Species Notes
Acharia stimulea Acer negundo Uramya pristis
A. saccharinum U. pristis
P. americana*
Cotesia empretiae
Asimina triloba U. pristis
Pl. americana*
P.s crassicornis Hyperparasitoid of Pl. americana
Carya glabra U. pristis
A. cocciphila*
Pl. americana*
Pe. crassicornis Hyperparasitoid of Pl. americana
Fagus grandifolia A. cocciphila*
U. pristis
Pl. americana*
Pe. crassicornis Hyperparasitoid of Pl. americana
T. discoideus
Lindera benzoin Pe. crassicornis
Ly. mandibularis Pseudohyperparasitoid through
Nyssa sylvatica U. pristis
Austrophorocera sp.*
Pe. crassicornis Hyperparasitoid of Pl. americana
Pl. americana*
Prunus serotina U. pristis
A. cocciphila*
Pl. americana*
Pe. crassicornis Hyperparasitoid of Pl. americana
Co. empretiae
Quercus alba U. pristis
A. cocciphila*
Pe. crassicornis Hyperparasitoid of Pl. americana
Pl. americana*
Co. empretiae
Quercus rubra Austrophorocera sp. Only egg observed
Ceraphron sp.* Hyperparasitoid, likely via
Pl. americana or Al. lilli
Pl. americana*
Acharia stimulea Quercus rubra T. discoideus
Pe. crassicornis Hyperparasitoid of Pl. americana
Acharia stimulea Quercus rubra Is. lycaenae*
Robinia pseudoacacia Pl. americana*
Co. empretiae
Adoneta spinuloides Carya glabra Pl. americana*
Fagus grandifolia U. pristis*
Nyssa sylvatica Pl. americana
Pe. crassicornis* Hyperparasitoid of Pl. americana
Quercus rubra U. pristis*
Al. lilli*
Pl. americana*
T. discoideus
Table 1. Continued.
Limacodid Species Plant Species Parasitoid Species Notes
Euclea delphinii Carpinus caroliniana Co. empretiae
Carya glabra U. pristis
Pl. americana*
T. discoideus
Fagus grandifolia U. pristis
Pl. americana*
T. discoideus
Nyssa sylvatica A.einaris*
Com. concinnata
T. discoideus
Taeniogonalos gundlachii Hyperparasitoid of Com. concinnata
Pl. americana*
Prunus serotina A.einaris*
U. pristis
Com. concinnata*
Pl. americana*
Quercus alba A.einaris*
Pl. americana*
Pe. crassicornis*
Trichogramma sp.* Egg parasitoid
T. discoideus
Quercus rubra A.einaris*
Euclea delphinii Quercus rubra Com. concinnata*
U. pristis
Al. lilli
Pl. americana*
Euclea delphinii Quercus rubra Pe. crassicornis* Hyperparasitoid of Pl. americana
T. discoideus
Diospyros virginiana Co. empretiae
Isa textula Acer saccharum U. pristis
Carya glabra U. pristis
Fagus grandifolia U. pristis
A. lilli
T. discoideus
Quercus alba U. pristis
Austrophorocera sp.*
Pl. americana*
Pe. crassicornis* Hyperparasitoid of Pl. americana
Is. lycaenae
T. discoideus
Quercus prinus U. pristis
Orthogonalys pulchella Hyperparasitoid of U.pristis
Quercus rubra U. pristis
Ceraphron sp. Hyperparasitoid of Pl.
americana*or Al. lilli*
Al. lilli
Pl. americana*
T. discoideus
Is. lycaenae* Likely hyperparasitoid of Triraphis
O. pulchella Hyperparasitoid of U.pristis
Table 1. Continued.
Limacodid Species Plant Species Parasitoid Species Notes
Isa/Natada Nyssa sylvatica Al. lilli
Fagus grandifolia U. pristis*
O. pulchella Hyperparasitoid of U.pristis
Quercus rubra U. pristis*
Lithacodes fasciola Acer negundo U. pristis*
Carya glabra U. pristis*
Pl. americana*
Fagus grandifolia Austrophorocera n. sp.*
Pl. americana*
Lithacodes fasciola Nyssa sylvatica Pl. americana*
Prunus serotina Austrophorocera n. sp.*
Pe. crassicornis* Hyperparasitoid of Pl. americana
T. discoideus
Quercus alba Austrophorocera sp.* Only egg observed
Pl. americana*
Quercus rubra Pl. americana*
T. discoideus
Lithacodes fasciola Quercus velutina Al. lilli
Lithacodes/Packardia Nyssa sylvatica Pl. americana
Natada nasoni Carya glabra Triraphis discoideus
Fagus grandifolia T. discoideus
Is. lycaenae*
Nyssa sylvatica T. discoideus
Prunus serotina Is. lycaenae*
Quercus alba T. discoideus
Quercus prinus Pe. crassicornis* Possible primary parasitoid
Quercus rubra Pl. americana*
T. discoideus
Is. lycaenae
Packardia geminata Fagus grandifolia U. pristis*
Pl. americana
Parasa chloris Quercus rubra Austrophorocera sp.* Only egg observed
T. discoideus
I. lycaenae*
Mesochorus discitergus Hyperparasitoid of Triraphis;
Phobetron pithecium Fagus grandifolia Pl. americana*
Prolimacodes badia Carya glabra H. fugitivus*
Diospyros virginiana A.imitator*
Fagus grandifolia A.imitator*
Pl. americana*
Is. lycaenae*
Me. discitergus Hyperparasitoid of Triraphis
Nyssa sylvatica Austrophorocera sp.* Only egg observed
Prunus serotina A.imitator*
T. discoideus
Quercus alba Pl. americana*
H. fugitivus*
ethanol to hexamethyldisilazane (HMDS)
(Heraty and Hawks 1998) before point- or
card-mounting. Images of specimens were
produced by scanning electron micros-
copy (SEM) and an EntoVision Imaging
Suite. Card- and point-mounted specimens
were examined using stereomicroscopes
with 10X or 25X oculars and fiber optic
light sources. Mylar film was used to dif-
fuse glare from fiber optic light sources to
reduce glare from the specimens. Scan-
ning electron microscope (SEM) images
3.2 mm Leica/Cambridge aluminum SEM
stubs with carbon adhesive tabs (Electron
Microscopy Sciences, #77825-12). Stub-
mounted specimens were sputter coated
using a Cressington Scientific 108 Auto
with gold from at least three differ-
ent angles to ensure complete coverage
(;20–30 nm coating). Wings were re-
moved and slide-mounted in polyvinyl
alcohol prior to imaging. Wing and
habitus images were captured using an
EntoVision Imaging Suite, which includes
a firewire JVC KY-75 3CCD digital cam-
era mounted on a Leica M16 zoom lens
via a Leica z-step microscope stand. Ad-
ditionally, a GT-Vision Lw11057C-SCI
digital camera attached to a Leica DMRB
compound scope was used to feed image
data to a desktop computer. The program
Cartograph 5.6.0 (Microvision Instru-
ments, France) was used to merge an
image series (typically representing 15
30 focal planes) into a single in-focus,
composite image. Lighting was achieved
using techniques summarized in Buffington
et al. (2005), Kerr et al. (2008), and
Buffington and Gates (2008).
Several images (see below) were ob-
tained with a Visionary Digital imaging
station. The station consists of an Infin-
ity Optics K2 long distance microscope
affixed to a Canon EOS 40D digital SLR
camera. Lighting was provided by a Dyna-
lite M2000er power pack and Microptics
ML1000 light box. Image capture soft-
ware is Visionary Digital proprietary ap-
plication with images saved as TIF with
the RAW conversion occurring in Adobe
Photoshop Lightroom 1.4. Image stacks
were montaged with Helicon Focus
4.2.1 for images of Conura nortonii
(Cresson), C. immaculata (Cresson),
Orthogonalys pulchella (Cresson), and
Taeniogonalos gundlachii (Cresson),
as well as habitus shots of Psychophagus
omnivorus (Walker), Platyplectrus ameri-
cana (Girault), and Pediobius crassicornis
Final image plates for hymenopteran
and dipteran figures were prepared using
Table 1. Continued.
Limacodid Species Plant Species Parasitoid Species Notes
Quercus rubra A.imitator*
Pl. americana*
T. discoideus
Tortricidia sp. Quercus alba Pl. americana*
Fagus grandifolia T. discoideus
Prunus serotina U. pristis*
Quercus rubra U. pristis*
Unknown limacodid Carya glabra Pl. americana
(too small to ID) Pe. crassicornis
Unknown limacodid Fagus grandifolia Pl. americana
Unknown limacodid Quercus rubra T. discoideus
InDesign CS4. Figures 14–17, previously
unpublished, were used with the permis-
sion of D. M. Wood (Agriculture and
Agri-Food Canada, Ottawa). Figures 38
and 43–44 were reproduced from Townes
(1970) with permission.
Diptera Imaging
The image of Systropus macer Loew
was captured with a Nikon Coolpix 8800
and adapters through the ocular of a
Leica MZ9.5 stereoscope. Tachinid images
were taken with a Canon EOS 40D digital
SLR camera mounted on a Kaiser RS1
copy stand. A Canon EF 100 mm f/2.8
macro lens was used for full body images,
and a Canon MP-E 65 mm 1–5X macro
lens was used for images of body struc-
tures. A ring light consisting of 80 LEDs
and covered with a reflective dome
provided the lighting. Image stacks were
montaged using Syncroscopy’s Auto-
Montage, and the resultant images received
further treatment in Adobe Photoshop
Hymenopteran terminology for sur-
face sculpture follows Harris (1979) and
for morphology follows Wahl (1993a),
Gibson (1997), Sharkey and Wharton
(1997), and Deans et al. (2010). Several
measurements for chalcidoids were taken,
including the following: body length, in
lateral view from the anterior projection
of the face to the tip of the metasoma;
head width through an imaginary line
connecting the farthest lateral projection
of the eyes; head height through an
imaginary line from the vertex to the
clypeal margin bisecting both the median
ocellus and the distance between the tor-
uli; malar space, in lateral view between
the ventral margin of the eye and lateral
margin of the oral fossa; posterior ocellar
line (POL), the shortest distance between
the posterior ocelli; ocular ocellar line
(OOL), the shortest distance between the
lateral margin of the posterior ocellus
and the eye orbit; marginal vein, the
length coincident with the leading fore
wing edge to the base of the stigmal
vein; stigmal vein, the length between its
base on the marginal vein and its apex;
postmarginal vein, the length from the
base of the stigmal vein to its apex on
the leading fore wing edge. Mesosomal
and metasomal sclerites were measured
dorsally along the midline. The use of
“[ ]” in descriptions denotes structures
that are not visible in the specimens upon
which the description is based (observed
via SEM), whereas their use in the ma-
terial examined section refer to author
notes. For braconids metasomal terga
1, 2, 3, etc. are abbreviated as T1, T2, T3;
antennal flagellomeres are abbreviated
as F1, F2, F3, etc. The ovipositor of
ichneumonids was measured from the
structure’s base (observed or inferred)
to its apex. The ovipositor sheaths must
sometimes be separated by a fine needle
to expose the ovipositor valves. The junc-
ture of the occipital and hypostomal carina
above the mandibular base is measured
in posterolateral view; it is sometimes
necessary to remove the head to properly
measure. Dipteran terminology follows
McAlpine (1981).
The portion of the key pertaining to
Chalcidoidea is based upon the keys in
Gibson et al. (1997). That for Braconidae is
based on the keys of Sharkey (1997) and
Whitfield (1997). That for Ichneumonidae
is based on the key of Wahl (1993b). The
diagnosis for Cotesia Cameron is based on
the key in Whitfield (1997); the diagnosis
for Ascogaster Wesmael is based on the
key in Shaw (1997a). The diagnosis for
Triraphis Ruthe is based on the diagnoses for
Triraphis and Rogas Nees in van Achterberg
(1991), as well as the keys in van Achterberg
(1991) and Shaw (1997b).
All binominals for hosts and parasitoids
are reported in their current nomenclatural
combinations. Although this is not
necessarily straightforward in any single
publication, we cite the most recent/
comprehensive nomenclatural authorities
where required. For Limacodidae, nomen-
clatural references include: Fletcher and
Nye (1982), Davis (1983), and Becker and
Epstein (1995). Host records from the lit-
erature for Tachinidae list the current
combination, as well as the combinations
under which the hosts and tachinids were
originally cited. New host records result-
and TMS for a particular parasitoid are
denoted with *inthetextandTable1.
Those newly reported herein from other
sources are denoted with
host records at the parasitoid species level
only rather than higher taxonomic levels.
Thus, if a parasitoid species is newly re-
corded for a particular host genus or spe-
cies, we report it as new even if congeneric
parasitoids were previously recorded from
that host genus or species.
Abbreviations for collections are as
follows: AEI (American Entomological
Institute, Gainesville, Florida, U.S.A.),
ANSP (Academy of Natural Sciences,
Philadelphia, Pennsylvania, U.S.A.), BMNH
(The Natural History Museum, London,
United Kingdom), CNC (Canadian
National Collection of Insects, Ottawa,
Ontario, Canada), IRSNB (Institut Royal
des Sciences Naturelles de Belgique,
Brussels, Belgium), MZLU (Zoological
Museum, Lund University, Lund, Sweden),
MCZ (Museum of Comparative Zoology,
Harvard University, Cambridge, Massa-
chusetts, U.S.A.), MSUC (Michigan State
University, East Lansing, Michigan,
U.S.A.), NHMW (Naturhistorisches
Museum Wien, Vienna, Austria), SEMK
(Snow Entomological Museum, Univer-
sity of Kansas, Lawrence, Kansas, U.S.
A.), ULQC (Universite
´Laval, Quebec,
Canada), USNM (National Museum of
Natural History, Smithsonian Institution,
Washington, DC, U.S.A.), and ZMUC
(Zoologisk Museum, Copenhagen,
Key to Primary and Secondary
Parasitoids Known From Limacodids
in North America
1. Hind wing modified as haltere (Fig. 1),
hamuli absent, mouthparts sponging (Fig. 1)
1’ Hind wing not modified as haltere, hamuli
present (Figs. 92–93), mouthparts mandib-
ulate (e.g., Figs. 39, 82) . .............
................... 5(Hymenoptera)
2. (1) Abdomen narrow and elongate, >2.0X
as long as broad and swollen apically
(Fig. 3). Wings shorter than abdomen.
Antennae and proboscis much longer than
head ..........................
..Systropus (macer Loew; Bombyliidae)
2’ Abdomen stout, less than 2.0X as long as
broad (Figs. 1, 11). Wings longer than ab-
domen. Antennae and proboscis at most as
long as head . . .........3(Tachinidae)
3. (2’) Facial ridge bare except for a few small
and decumbent setae on lower third or less
(Fig. 2). Metathoracic spiracle fringed with
plumose hairs of about equal length along
both anterior and posterior edges, leaving a
V-shaped middorsal opening (Fig. 4). Pros-
Uramya (pristis (Walker), limacodis (Townsend))
3’ Facial ridge with row of stout setae on lower
one-half or more (Fig. 9). Metathoracic
spiracle with posterior lappet much larger
than anterior one (Fig. 5). Prosternum haired
4. (3’) Ocellar seta vestigial or absent (Fig. 9).
Abdominal terga 3 and 4 each with 1 pair of
median discal setae. Bend of vein M obtuse
(Fig. 8). Female with sickle-shaped, pierc-
ing ovipositor (Fig. 8) . . .............
......Compsilura (concinnata (Meigen))
4’ Ocellar seta well developed, similar in size
to outer vertical seta (Fig. 10). Abdominal
terga 3 and 4 without median discal setae
(Fig. 11). Bend of vein Malmost a right angle
(Fig. 11). Female with short, non-piercing
..Austrophorocera (cocciphila (Aldrich and
Webber), coccyx (Aldrich and Webber), A.
einaris (Smith), A. imitator (Aldrich and
Webber), A. n. sp.)
5 (1’) Fore wing venation complete, with at
least 2 closed cells (Figs. 18, 28, 34) . . . . .
. . . . . . 6 (Trigonaloidea, Ichneumonoidea)
5’ Fore wing venation reduced, with fewer
than 2 closed cells (Figs. 77–78, 96, 113)
6. (5) Fore wing with veins C and SC+R
touching/fused, costal cell absent (Figs.
63, 65). Metasomal sterna less strongly
sclerotized than terga (Figs. 19, 23, 27,
29) .............7(Ichneumonoidea)
6’ Fore wing with veins C and SC+R separate,
costal cell present (Fig. 18). Metasomal sterna
and terga equally sclerotized (Figs. 70–73) . .
7. (6) Fore wing with vein 2m-cu present
(Fig. 28, 35) ........8(Ichneumonidae)
7’ Fore wing with vein 2m-cu absent (Fig.
56) ............... 14(Braconidae)
8. (7) Areolet of fore wing closed, large and
rhombic (Figs. 20, 26). Ovipositor delicate,
needlelike, sheaths thick and rigid (Figs. 19,
22). Female hypopygium prominent and tri-
angular in lateral view (Fig. 22). Male gon-
oforceps produced into elongate process.
Spiracle of metasomal segment 1 near or just
behind middle, glymma large and deep
(Fig. 21) . . . ......................
.........Mesochorus (discitergus (Say))
8’ Areolet of fore wing open (vein 3rs-m ab-
sent) or closed, if closed then cell obliquely
quadrate and petiolate (Figs. 26, 34).
Ovipositor always stouter than above, sheaths
thin, often curved if ovipositor ;2.0X as
long as metasomal apical depth. Female
hypopygium small and quadrate in lateral
view (Fig. 23). Male gonoforceps not
produced. Spiracle of metasomal segment
1 beyond middle, glymma small or absent
(Fig.27) .......................9
9. (8’) Ventral posterior corner of pro-
pleuron with strongly produced, more or
less angulate lobe touching or overlapping
pronotum (Fig. 24). Sternaulus about 0.3X
as long as mesopleuron (Fig. 30). Clypeus
not separated from supraclypeal area by
distinct groove (Fig. 32). Areolet of fore
wing closed, obliquely quadrate and peti-
olate (Fig. 26) . . . ................10
9’ Ventral posterior corner of propleuron not
developed as distinct lobe, not angulate, at
most with weak groove delimiting it from
main area of propleuron (Fig. 25). Sternaulus
of mesopleuron present and reaching middle
coxa (Fig. 31). Clypeus separated from su-
praclypeal area by groove (Fig. 33). Areolet
of fore wing open and pentagonal (Fig. 34).
10. (9) Petiole of fir st met asomal segment
long and cylindrical in cross-section;
midpoint of petiole with tergo-sternal
suture at midline; T1 without trace of glymma
(Fig. 28) . . . .Casinaria (grandis Walley)
10’ Petiole of first metasomal segment shorter
and quadrate in cross-section; midpoint of
petiole with tergo-sternal suture close to
ventral margin; T1 with glymma present as
pitlike impression (Fig. 27) . . . .......
...........Hyposoter (fugitivus (Say))
11. (9’) Body color (excluding legs) black with
white markings (Fig. 29). Mesosoma with
coarse punctures (Fig. 37). Dorsal margin
of pronotum with strong swelling at dorsal
end of epomia (Fig. 37). Fore wing 5.0–8.5
mm long . . ......................
........Baryceros (texanus (Ashmead))
11’ Body color (excluding legs) ranging from
uniformly black/dark brown to having
brownish red areas on T2–T3 (Figs. 34–36).
Mesosoma with punctures ranging from
fine to absent. Dorsal margin of pronotum
without strong swelling at dorsal end of
12. (11’) Apical 0.3 of clypeus turned inward
at 90°(Fig. 43). Vein 2-Cu of hind wing
basally incomplete . . ...............
....Lysibia (mandibularis (Provancher))
12’ Apical 0.3 of clypeus flat, not turned in-
ward (Fig. 44). Vein 2-Cu of hind wing
complete . . ....................13
13. (12’) Occipital and hypostomal carinae
meeting at mandibular base (Fig. 41) . . .
Acrolyta (nigricapitata (Cook and Davis))
13’ Occipital carina meeting hypostomal ca-
rina above mandibular base, juncture sep-
arated from base by about 0.2X basal
mandibular width (Fig. 42) ...........
........ Isdromas (lycaenae (Howard))
14. (7’) Labrum visible through gap between
ventral margin of clypeus and mandibles
and concave (as in Fig. 39) (cyclostome
Braconidae). Epicnemial carina present
(Fig. 47); median carina present on T1 and
usually extending posteriorly from dorsal
carinae; T2 striate to striate-rugose (Figs.
52, 57); fore wing vein 1m-cu basad or in
line with fore wing vein 2RS (Fig. 56) . .
.......... Trira ph is Ruthe (eupoeyiae
(Ashmead), discoideus (Cresson), harri-
sinae (Ashmead))
14’ Labrum concealed by clypeus or if vis-
ible not concave (Figs. 40 (as in), 60)
(non- cyclostome Braconidae). Without
combination of characters found in
Triraphis .......................
........Cheloninae, Microgastrinae, 15
15. (14’) Occipital carina present, fore wing
vein RS reaching wing margin as tubular
vein, T1–T3 fused into carapace cover-
ing all other terga (Figs. 48, 63). Other
characters not in combination found in
Cotesia (seebelow) ..............
..Ascogaster (quadridentata Wesm a el )
15’ Occipital carina absent, fore wing vein RS
not reaching wing margin as tubular vein
(Fig. 65), T1–T3 variable (Microgastrinae).
Fore wing vein r-m absent, areolet ab-
sent (Fig. 65). Ovipositor and sheaths
short, weakly extending beyond tip of
hypopygium (Fig. 67). Hypopygium evenly
sclerotized medially. Propodeum with-
out areola, often with distinct medial
longitudinal carina and usually rugose
.. Cotesia (empretiae (Viereck), phobetri
(Rohwer), schaffneri (Rohwer))
16. (6’) Antenna black with white or light
yellow band at center. Head and mesosoma
with black and white pattern. Metasoma
and legs mostly orange (Figs. 70–71).
Metasoma thin, smooth, impunctate . . .
.....Orthogonalys (pulchella Cresson)
16’ Antenna yellowish brown without lighter
band at center. Head and body black with
yellow markings (Figs. 72–73). Meta-
soma black and yellow, legs yellow with
dark brown to black on femur, tibia
dusky in apical third. Metasoma stout,
..Taeniogonalos (gundlachii (Cresson))
17. (5’) Fore wing with tubular submarginal
vein basally on anterior margin, stigmal
vein curving distally (Fig. 77). Female an-
tenna with 7–8 flagellomeres (Fig. 76), male
with 8–9 (Fig. 74). Dorsum flat in lateral
view ............................
......Ceraphron Jurine (Ceraphronidae)
17’ Fore wing lacking tubular vein basally on
anterior margin (Fig. 78). Other features
not as above . ..................18
18. (17’) Hind femur enlarged (<3.0X as long
as broad) (Figs. 79–80). Prepectus reduced
to a small sclerite along dorsal margin of
mesopleuron (Fig. 83). Gaster petiolate
(Figs. 79–80) . ....................
....Conura (camescens (Cameron), nig-
ricornis (F.), nortonii (Cresson))
18’ Hind femur not enlarged (>3.0X as long as
broad). Prepectus larger, triangular (Figs.
87, 109). Gaster indistinctly or not petiolate
...... Eulophidae,Trichogrammatidae,
Pteromalidae, 19
19. (18’) Legs with 5 tarsomeres. Clypeus
shallowly bilobed apically (Fig. 88).
Pedicel ;2.0X as long as broad, antennal
formula 11263 (Fig. 89). Propodeum re-
ticulate-rugose, with median carina in-
complete (Fig. 90). Gaster subcircular to
.....Psychophagus (omnivorus (Walker))
19’ Legs with 3 or 4 tarsomeres. Other features
not as above . ....................
. . . . Eulophidae, Trichogrammatidae, 20
20. (19’) Legs with 4 tarsomeres. Fore wing
lacking setal tracks (Fig. 113) .........
....................Eulophidae, 21
20’ Legs with 3 tarsomeres (Fig. 94). Fore
wing with setal tracks, broad with sig-
moid venation and distinctive Rs1 setal
...... Trichogramma (minutum Riley)
21. (20’) Scutellu