Forest Service Northern Research Station General Technical Report NRS-179

United States Department of Agriculture

June 2018

Laura M. Blackburn and Ann E. Hajek

Gypsy Moth Larval Necropsy Guide
Manuscript received for publication 4 December 2017

Abstract

Since the early 1900s, a number of parasitoids have been released for classical biological control 

of the introduced destructive forest insect, Lymantria dispar (gypsy moth), in North America. 

During this time, two pathogens were accidentally introduced. These pathogens and several of 

the parasitoid species are now commonly found in North American gypsy moth populations. 

The aim in creating this guide was to provide laboratory techniques for distinguishing between 

two common pathogens, the fungus Entomophaga maimaiga and the gypsy moth multiple 

nucleopolyhedrovirus, and provide illustrations and images for adults, puparia, and cocoons 

of established gypsy moth parasitoids commonly found in the larval or pupal stages of gypsy 

moth in North America. Gypsy moth collection and rearing techniques are also reviewed, and 

a technical glossary and summary table highlighting the affected life stage by gypsy moth 

parasitoids in North America are included in this guide.

The Authors

LAURA M. BLACKBURN is an operations research analyst with the USDA Forest Service, Northern 

Research Station, Morgantown, WV 26505.

ANN E. HAJEK is a professor of insect pathology, disease ecology, and biological control at 

Cornell University, Department of Entomology, Ithaca, NY 14853.

Cover

North American range of the gypsy moth as of 2017, showing suitable habitat and some 

common pathogens and parasitoids of the gypsy moth. Photo credits from left to right: 

Entomophaga maimaiga conidia on cadaver (photo by Ann Hajek, Cornell University, used with 

permission), gypsy moth larva killed by the viral pathogen LdMNPV (photo by Ruth Plymale, 

Ouachita Baptist University, used with permission), Blepharipa pratensis (photo by Christophe 

Quintin; CC BY-NC 2.0 Generic), Parasetigena silvestris-parasitized gypsy moth larva (photo by 

Gyorgy Csoka, Hungary Forest Research Institute, via Bugwood.org).

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Gypsy Moth Larval Necropsy Guide

General Technical Report NRS-179

CONTENTS

Introduction......................................................................................................... 1

Pathogen Versus Parasitoid........................................................................... 2

Common Pathogens ......................................................................................... 2

Fungal Pathogen......................................................................................... 2

Viral Pathogen.............................................................................................. 5

Less Common Pathogens ............................................................................... 6

Fungal Pathogens....................................................................................... 6

Common Parasitoids ........................................................................................ 7

Hymenoptera (Wasps).............................................................................. 8

Diptera (Flies) .............................................................................................10

Egg Parasitoids ..........................................................................................13

Uncommon Parasitoids .................................................................................13

Hymenoptera (Wasps)............................................................................13

Diptera...........................................................................................................15

Gypsy Moth Collection And Rearing Techniques ...............................17

Field Collections........................................................................................17

Rearing Gypsy Moth Larvae.................................................................18

Pathogen Identification ................................................................................19

Laboratory Dissections ..........................................................................19

Molecular Methods..................................................................................21

Summary .............................................................................................................22

Acknowledgments ..........................................................................................22

Literature Cited.................................................................................................22

Glossary................................................................................................................29
Gypsy moth defoliation of hardwood trees along the Allegheny Front near Snow Shoe, Pennsylvania, in July 2007. 

Photo by Dhalusa (CC BY-SA 3.0), via Wikimedia Commons.
1

INTRODUCTION

Gypsy moth (Lymantria dispar) is a major pest of 

deciduous trees in the northern hemisphere. Native 

to Eurasia, this species was introduced to eastern 

Massachusetts by 1869 (Forbush and Fernald 1896), 

and despite many efforts to control gypsy moth spread, 

range expansion continues. Gypsy moth now occurs 

from Canada (southern Ontario, southern Quebec, 

southwestern New Brunswick, and southwestern 

Nova Scotia), south to North Carolina, and west to 

Wisconsin, Illinois, Indiana, Ohio, and West Virginia 

(Fig. 1). Since the early 1900s, intensive efforts have been 

made to introduce biological control agents against the 

gypsy moth, including 34 parasitoids, 1 predator, and 5 

pathogens (Fuester et al. 2014). Successful establishment 

of parasitoid introductions has been limited, as has the 

effects of parasitoids on the regulation of gypsy moth 

populations (Hajek and van Nouhuys 2016). While 

parasitoids contribute to the overall mortality of gypsy 

moth populations, pathogens often play a more vital role.

This guide is intended to assist researchers involved in 

gypsy moth life table studies. A life table accounts for 

stage-specific mortality in a study population. Life tables 

can provide key information on the magnitude of host 

mortality caused by specific agents, information that 

is critical to understanding the role of such agents in 

host population dynamics. Knowledge of the processes 

driving gypsy moth population dynamics is crucial for 

understanding gypsy moth outbreaks and management 

practices.

This publication focuses on the larval stage, where 

mortality due to natural enemies can be substantial. 

Death of field-collected larvae can generally be 

attributed to one of four causes:

• A fungal pathogen: most commonly 

Entomophaga maimaiga

• A viral pathogen: Lymantria dispar multiple 

nucleopolyhedrovirus (LdMNPV)

• Parasitoids: flies and wasps

• Some combination of these agents

Descriptions of the most common pathogen and 

parasitoid species that cause death in gypsy moths are 

provided in this document, along with descriptions of 

some less common agents. Information on identifying 

selected hyperparasitoids is also included for species 

commonly emerging from field-collected gypsy moth 

larvae or pupae. This guide focuses on collecting gypsy 

moth larvae and pupae, thus only limited information 

on egg parasitoids is provided. Collection and rearing 

techniques for gypsy moth larvae and pupae and 

laboratory techniques for identifying the most likely 

causes of mortality are also reviewed.

Figure 1.—Range of gypsy moth 

in North America as of 2017.
2

PATHOGEN VERSUS PARASITOID

The cause of death in gypsy moth can be narrowed 

down by considering the life stage at time of death. For 

example, infections by Entomophaga maimaiga and 

LdMNPV can occur in all larval stages but are usually 

most prevalent in late instars (Hajek 1994, Hajek and 

Snyder 1992). Parasitoids are limited by their ability to 

only develop in specific life stages. Table 1 provides a 

list of affected life stages for commonly occurring gypsy 

moth parasitoids.

Physical characteristics of the cadaver also provide 

useful clues that can aid in identifying the cause of 

death. Be sure to note distinguishing characteristics 

such as a sugar coating on larval hairs (Hajek and 

Roberts 1992), a thin cuticle (Hajek and Roberts 1992), 

an oozing wound (Campbell 1963), or large white eggs 

on the dorsum of late instar larvae (Fuester et al. 2014, 

Ticehurst et al. 1978). When gypsy moth-parasitized 

larvae or pupae are held in the laboratory, eventually a 

life stage of the parasitoid may emerge. However, when 

field-collected larvae die during rearing and no other 

insect eventually appears in the container with the 

cadaver, the cause of death will likely be a pathogen.

COMMON PATHOGENS

Fungal Pathogen

Entomophaga maimaiga Humber, Shimazu 

& Soper (Phylum Zoopagomycota; Order 

Entomophthorales; Family Entomophthoraceae)

Native to Asia, Entomophaga maimaiga was deliberately 

released in the United States in 1910-11 and again 

in 1985-86 (Nielsen et al. 2005). Though attempts to 

establish this fungal pathogen in the United States 

were thought to have failed, epizootics were first seen 

during the very wet spring of 1989, predominantly 

in New England (although not near areas of the 

1985-86 releases), after which this fungus spread 

Table 1.—North American gypsy moth parasitoids and hyperparasitoids and affected host life stages. Species are included 

in order of host stages initially parasitized.

Host stages affected

Parasitoid Initial parasitization Exit

Hymenoptera, Eupelmidae: Anastatus japonicus Ashmead eggs eggs

Hymenoptera, Encyrtidae: Ooencyrtus kuvanae (Howard) eggs eggs

Hymenoptera, Ichneumonidae: Phobocampe unicincta (Gravenhorst) early instars middle instars

Hymenoptera, Braconidae: Cotesia melanoscela (Ratzeburg) early, middle instars middle instars

*Hymenoptera, Braconidae: Aleiodes indiscretus (Reardon) early, middle instars late instars

Diptera, Tachinidae: Compsilura concinnata (Meigen) early, middle, late instars middle, late instars, pupae

Diptera, Tachinidae: Blepharipa pratensis (Meigen) early, middle, late instars late instars, pupae

Diptera, Tachinidae: Parasetigena silvestris (Robineau-Desvoidy) middle, late instars late instars, pupae

*Diptera, Tachinidae: Exorista larvarum (Linnaeus) late instars late instars

*Diptera, Tachinidae: Exorista mella (Walker) late instars late instars

*Diptera, Tachinidae: Lespesia frenchii (Williston) late instars late instars, pupae

*Hymenoptera, Perilampidae: Perilampus hyalinus Say late instars pupae

*Diptera, Sarcophagidae: Agria housei Shewell late instars, pupae pupae

Hymenoptera, Ichneumonidae: Theronia atalantae (Poda) prepupae, freshly formed pupae pupae

Hymenoptera, Ichneumonidae: Pimpla disparis Viereck prepupae, pupae prepupae, pupae

*Hymenoptera, Ichneumonidae: Pimpla pedalis Cresson prepupae, pupae prepupae, pupae

*Hymenoptera, Ichneumonidae: Itoplectis conquisitor (Say) prepupae, pupae pupae

*Diptera, Sarcophagidae: Arachnidomyia aldrichi Parker prepupae, pupae pupae

Hymenoptera, Chalcididae: Brachymeria intermedia (Nees) pupae pupae

*Hymenoptera, Torymidae: Monodontomerus aereus Walker pupae pupae

*Indicates species that are uncommon or rare parasites of gypsy moths. Many of the species released for classical biological control are not well 

established or are only established in certain areas.
3

rapidly throughout the contiguous gypsy moth range 

(Andreadis and Weseloh 1990, Hajek 1999, Hajek et 

al. 1990). Like all fungi, E. maimaiga is greatly affected 

by moisture and temperature. Thriving in cool and wet 

spring and early summer weather, this fungus is an 

important source of larval mortality in both low- and 

high-density gypsy moth populations.

Extensive laboratory and field tests have demonstrated 

that while E. maimaiga can infect some other 

lepidopteran species in the laboratory, high levels of 

infection in any species other than gypsy moth have 

never been reported from the field (Hajek et al.1995, 

Hajek et al. 1996, Hajek et al. 2000a, Hajek et al. 

2004). This fungus can be grown outside of insects (in 

vitro) in the laboratory under special conditions, but 

this process is difficult and viable methods for mass 

production have not been developed.

Entomophaga maimaiga produces two kinds of spores 

(conidia [singular = conidium] and resting spores 

[azygospores]), but spores are only produced after 

host death. The type of spore produced is dependent 

on host age. Infective conidia (asexual short-lived 

spores) are usually produced from early instar cadavers, 

while resting spores, and sometimes conidia also, are 

produced from later instar cadavers and occasionally 

from pupae (Hajek 1999). Resting spores, produced 

near the end of the larval season, overwinter in soil and 

can survive for at least 6-12 years (Hajek et al. 2000b, 

Weseloh and Andreadis 1997). During May and early 

June, resting spores germinate and produce infective 

germ conidia. The conidia (either from cadavers 

or resting spores) are actively discharged and are 

dispersed by the wind, where they infect gypsy moth 

larvae with which they come into contact. Once they 

land on a host, the conidia grow through the cuticle 

and start consuming the living insect. After the fungus 

consumes the inside of the larva and the larva is dead, 

under humid conditions conidiophores can grow out 

through the cuticle. Conidia that are produced from the 

ends of the conidiophores are shot into the air where 

they are dispersed and can then infect other larvae. If 

no host is found, a conidium can germinate and expel 

a secondary conidium. This infection cycle continues 

throughout the season. Resting spore production 

occurs in the fourth to sixth instar larvae, which allows 

the spores to overwinter and start the cycle again the 

next year or in subsequent years (Fig. 2).

Entomophaga maimaiga conidia—Conidia are 

produced externally on infected gypsy moth larvae 

after they die. Whether an infected larva will produce 

conidia depends on a variety of factors, including larval 

age, fungal isolate, and environmental conditions. If 

conidia are produced from the cadaver, they will be 

formed within hours to a few days after larval death 

(Fig. 3A). Conidia are actively expelled from the 

cadaver, and E. maimaiga is spread within surrounding 

gypsy moth populations during the current season, 

although some spores may travel longer distances.

Conidia (Fig. 3B) produced by cadavers are clear, 

pear-shaped spores measuring approximately 20 μm x 

25 μm. Conidia are usually produced from cadavers of 

early instars, which typically are found with the prolegs 

gripping a twig or branch and the anterior portion of 

the body at a 90o angle hanging downward compared 

Figure 2.—Multiplicative cycles of Entomophaga 

maimaiga infection during one field season that 

create epizootics. Models have indicated that during 

epizootics, four to nine infection cycles can occur 

during one gypsy moth generation. Illustration from 

Hajek 2004, used with permission.
4

with the posterior part of the body (Fig. 3A). Once 

conidia have been discharged, some spores can be 

retained on the larval hairs and may appear similar to 

a sugar coating. The surface of a cadaver from which 

conidia are being ejected appears moist and white to 

light brown. After conidial discharge, these wet spores 

and conidiophores will decompose quickly, and soon 

there will be little to no indication that they were ever 

present. Therefore, in order to detect these fungal 

stages, daily monitoring of larval death and of cadavers 

(for ~3 days after death) is very important.

Entomophaga maimaiga resting spores—Later in the 

larval season when gypsy moths are in the fourth- to 

sixth-instars, E. maimaiga will produce thick-walled 

resting spores inside of cadavers. The bodies of 

recently killed larvae containing resting spores are 

soft, and the contents appear to be liquid. Over time, 

the cadavers become dry and stiff with a firm, but not 

fragile, cuticle. Cadavers containing resting spores can 

often be found hanging vertically on tree trunks with 

prolegs usually extended at 90o from the body (Fig. 4). 

After some time, the cadavers fall from the trees 

and disintegrate on the ground. Most resting spores 

overwinter in the top layers of the soil and germinate 

during the following year(s) where they infect gypsy 

moth larvae in successive generations.

Figure 4.—Symptomatic expression of resting sporeproducing cadaver with legs extended and stiff posture. 

Cadavers of late instars killed by E. maimaiga are typically 

seen in this position on tree trunks during epizootics. 

Illustration by Laura Blackburn, USDA Forest Service.

Figure 3.—Entomophaga maimaiga conidia on cadavers (A) are produced relatively shortly after the larva dies. Conidia 

shot off from the cadaver adhere to the larval hairs, making the dark larval hairs appear white. (B) A microscopic view of 

E. maimaiga conidia (20 μm x 25 μm) stained red to improve viewing. Photos by Ann Hajek, Cornell University, used with 

permission. 

A

B
5

Resting spores develop from immature to mature 

spores in the days following larval death (Fig. 5). When 

first formed, resting spores possess a single thin wall 

and have a granular interior. As the spores mature, 

they develop a thickened wall and the granular interior 

coalesces into a small number of large lipid droplets. 

Normally, mature resting spores are observed in 

cadavers of larvae that have died at least several days 

before, but occasionally immature spores are also still 

visible at that time. Resting spores are approximately 30 

μm in diameter (Fig. 6A) and at times can be mistaken 

for air bubbles (Fig. 6B).

Viral Pathogen

Lymantria dispar multiple nucleopolyhedrovirus

(LdMNPV)

LdMNPV is a highly host-specific virus that infects 

gypsy moth larvae when they consume foliage 

contaminated with viral occlusion bodies. This virus 

most likely arrived in North America before 1907, 

along with parasitoids being introduced for biological 

control (Howard and Fiske 1911). Prior to the arrival 

of E. maimaiga, LdMNPV was well known for causing 

epizootics resulting in the collapse of outbreak gypsy 

moth populations, but this usually only occurred after 

populations had reached high densities. On its own, 

LdMNPV only increases in abundance in high density 

populations (Doane and McManus 1981). LdMNPV 

is now found infecting insects in high density gypsy 

moth populations, but its prevalence is often low, 

and E. maimaiga infections have become much more 

common, at times leading to the collapse of gypsy moth 

populations at a variety of densities (Hajek et al. 2015, 

Hajek and van Nouhuys 2016, Liebhold et al. 2013).

After being ingested by a gypsy moth larva, viral 

occlusion bodies (environmentally persistent protein 

packets containing the actual virus) dissolve in the 

alkalinity of the insect’s gut, releasing the virions 

(infective virus particles). The virions infect the midgut 

cells and infection subsequently spreads to the rest of 

the body. The virus replicates in the nuclei of infected 

cells, forming two types of progeny: budded virus and 

occluded virus. The budded virus leaves an infected 

cell and spreads the infection to additional cells within 

the insect. Toward the end of the infection cycle, new 

occlusion bodies are made that contain the occluded 

virus. Subsequently, infected cells burst, releasing 

the occlusion bodies. This systematic destruction 

of infected cells eventually destroys the internal 

organs of the host, leaving the cadaver as a slurry of 

occlusion bodies and cell fragments. The virus also 

destroys cells beneath the larval cuticle and produces 

cuticle-degrading enzymes, making the cuticle very 

thin and fragile (Miller 2013). After the host dies and 

the cadaver decomposes, the cuticle breaks and viral 

occlusion bodies are released into the environment.

Figure 5.—Progression of Entomophaga maimaiga resting 

spore maturation following gypsy moth larva death (adapted 

from Hajek et al. 2008).

Figure 6.—Mature E. maimaiga resting spores (A) are typically found 

with a thick wall and a single large lipid droplet. Younger resting spores 

will contain multiple lipid droplets, giving them a granular appearance. 

However air bubbles under slide covers (B) can easily be mistaken as 

immature spores. Use the fine focus dial on the microscope to overcome 

this by simply zooming in and out. The air bubbles will have a thick dark 

wall. The wall on mature resting spores is thick and white/clear. Photos by 

Ruth Plymale, Ouachita Baptist University, used with permission.
6

Larvae that die from LdMNPV often hang in an upside 

down V shape (an “inverted V”; Fig. 7) and have a very 

thin cuticle. The cadavers may break when collected 

or before being dissected, so if you have to pull to tear 

the cuticle apart, it is unlikely that the larva died from 

LdMNPV alone. A larva killed by LdMNPV alone will 

have a large number of occlusion bodies present within 

them (Fig. 8A). The occlusion bodies are 1-10 μm in 

diameter and are generally spherical but with slightly 

uneven sides. Occlusion bodies under a cover slip 

on a microscope slide will appear bright and sparkly 

under a compound microscope using phase contrast. 

(The view in phase contrast has been described as 

similar to the appearance of the Milky Way on a dark 

night). While viewing the occlusion bodies at 400x 

and focusing up and down, the occlusion bodies will 

often have a dark center and bright halo. To diagnose 

LdMNPV in cadavers, add potassium hydroxide (1 M 

KOH), an alkaline solution, to the side of the cover slip, 

and watch for the dissolution of the occlusion bodies 

as the KOH moves under the cover slip (Fig. 8B). Refer 

to the section on pathogen identification for detailed 

techniques on microscopic examination.

LESS COMMON PATHOGENS

Fungal Pathogens

Isaria farinosa–(Phylum Ascomycota; Order 

Hypocreales; Family Cordycipitaceae)

Long known as Paecilomyces farinosus, this 

entomopathogenic fungus has a relatively broad host 

range and is found throughout the world in both 

tropical and temperate zones. It can be isolated from 

water, air, arthropods, plants, and other fungi. This 

ubiquitous fungus also infects and kills lepidopteran 

larvae and pupae and is most commonly found in 

forest soils with pine (Pinus spp.), oak (Quercus spp.), 

poplar (Populus spp.), and acacia (Acacia spp.) trees 

(Domsch et al. 1980). It is a moderately fast-growing 

fungus, beginning with white colonies that turn yellow 

and form woolly aerial mycelium. The conidia are 

transparent and circular to fusiform (2-3 μm x 1-1.8 

μm). After death, fungal mycelia grow out from infected 

gypsy moth larvae, often covering cadavers, which can 

look white and flocculent. See Zimmermann (2008) for 

Figure 8.—LdMNPV occlusion 

bodies (OBs) viewed with 

phase contrast microscopy at 

(A) 100x and at (B) 400x while 

dissolving from the addition 

of KOH, which was added at 

the lower right (lower right 

OBs have dissolved while 

upper left OBs have not). 

Photos by Ruth Plymale, 

Ouachita Baptist University, 

used with permission.

Figure 7.—Gypsy moth larvae killed by LdMNPV in 

the typical upside down (inverted) V-shape. Photo 

by Ruth Plymale, Ouachita Baptist University, used 

with permission.

A B

KOH

direction

Undissolved OB

Not OB

OB in process of dissolving

Dissolved OB
7

an in-depth synthesis of information on the biology, 

ecology, and use of I. farinosa in biological control.

Prior to the first reports of E. maimaiga in North 

America, Majchrowicz and Yendol (1973) found I. 

farinosa to be the most common fungal pathogen of 

diseased gypsy moth larvae and pupae, infecting over 

12 percent of a study population in Berks County, 

Pennsylvania in 1971. After E. maimaiga arrived, Hajek 

et al. (1997) found 4.9-12.2 percent I. farinosa infection 

in gypsy moth larvae from the Mid-Atlantic region, 

with no association, either positive or negative, between 

E. maimaiga and I. farinosa.

Beauveria bassiana (Phylum Ascomycota; Order 

Hypocreales; Family Cordycipitaceae)

First observed in 1835 in silkworms (Bombyx mori) 

reared for silk production, this fungus causes larvae 

to harden after death, and white mycelia grows out of 

the dead insect (Kleespies et al. 2008). This contagious 

condition, termed “white muscardine,” was found to 

be caused by the fungus B. bassiana. This fungus has 

a wide host range of over 200 insect species, mainly 

Lepidoptera and Coleoptera (Feng et al. 1994), and it is 

generally associated with shaded areas such as forests 

and uncultivated hedgerows (Bidochka et al. 1998, 

Meyling and Eilenberg 2006). As with E. maimaiga 

and I. farinosa, infection is caused by conidia that 

attach to the host cuticle and germinate in humid 

environments. The host cuticle is penetrated by germ 

tubes growing from the conidia. Fungal growth 

increases upon entering the hemocoel, and the host 

dies due to depleted hemolymph, nutrients, or toxemia 

(Khachatourians 1991). When moisture levels are high, 

B. bassiana produces aerial conidia on the cuticle of the 

host. Diagnostic characteristics include conidiogenous 

cells that are typically densely clustered or whorled with 

toothed rachises that extend apically and bear a single 

conidium per denticle (Humber 1997). The nearly 

globose conidia are <3.5 μm in diameter.

Although B. bassiana is common, infections by the 

fungus are low in prevalence. For example, Majchrowicz 

and Yendol (1973) isolated B. bassiana from 7 percent 

of dead gypsy moth larvae and pupae collected in Berks 

County, Pennsylvania in 1971, and infections were rare 

in a study of four Mid-Atlantic states, after E. maimaiga

had become established (Hajek et al. 1997). Trials 

applying B. bassiana against gypsy moth larvae in 

Slovakia and Maryland did not yield control (see Hajek 

et al. 1997).

COMMON PARASITOIDS

The term “parasitoid” refers to insects that spend part 

of their life, usually immature stages, attached to or 

within an insect host where they feed on and ultimately 

kill the host. Since 1906, parasitoid species have been 

introduced from Europe and Asia as part of a large 

classical biological control program for the gypsy moth. 

These parasitoids belong to the orders Hymenoptera 

(wasps) and Diptera (flies). The greatest number of 

parasitoid species belong to the order Hymenoptera 

and generally attack immature stages (including eggs) 

of host insects (Eggleton and Belshaw 1993). Species 

of parasitoids in the order Diptera have a wide host 

range and attack all stages of insect hosts except the 

egg (Eggleton and Belshaw 1993), with members of the 

family Tachinidae being the most beneficial (Clausen 

1972).

Parasitoids and hyperparasitoids of gypsy moth larvae 

and pupae that are established in North America are 

described below. For a review of historic (1906-1959) 

gypsy moth parasitoid releases see Clausen (1978), 

and for parasitoid releases between 1961 and 1977 

see Reardon (1981). A list of introduced Eurasian 

and North African parasitoids that are known to 

be established in North America is presented in 

Fuester et al. (2014). Simons et al. (1979) provide a 

more exhaustive key to gypsy moth parasitoids and 

hyperparasitoids (including species that are rare or 

were released but are not considered established). Keys 

to gypsy moth parasitoids in specific families are also 

available: Braconidae (Marsh 1979), Ichneumonidae 

(Dasch 1971, Gupta 1983), and Tachinidae (Sabrosky 

and Reardon 1976).

The most commonly encountered parasitoids reared 

from gypsy moth larvae include Cotesia melanoscela, 

Phobocampe unicincta, Compsilura concinnata, 

Parasetigena silvestris, and Blepharipa pratensis (Hajek 

and van Nouhuys 2016, Williams et al. 1993). Common 

parasitoids reared from gypsy moth pupae include 

Brachymeria intermedia, Pimpla disparis, Parasetigena 

silvestris, and Theronia atalantae fulvescens (Fuester 

et al. 1997). Most of these commonly encountered 

parasitoids are multivoltine and will transform into 

adults within the same season that gypsy moth larvae 

are collected, which simplifies identification. The 

exceptions include P. unicincta, B. pratensis, and P. 

silvestris, univoltine species that overwinter as cocoons 

or puparia and emerge as adults the following spring. 
8

These three parasitoids, however, are very easy to 

identify based on their puparia.

Parasitoid assemblages vary with gypsy moth 

population density. When gypsy moth densities 

are low, the tachinids Compsilura concinnata and 

Blepharipa pratensis contribute more to overall larval 

mortality (Doane and McManus 1981). When gypsy 

moth densities are high, the chalcidid Brachymeria 

intermedia causes high levels of mortality (Doane and 

McManus 1981, Ticehurst et al. 1978). During a gypsy 

moth outbreak, Cotesia melanoscela was found to be 

most abundant (Williams et al. 1992). After a gypsy 

moth outbreak, the tachinid Blepharipa pratensis can be 

a great source of mortality (Ticehurst et al. 1978). And 

2 years post-outbreak, Parasetigena silvestris reaches 

peak density (Sabrosky and Reardon 1976).

Forest type, ecological relationships, and site 

characteristics play a major role in the dominance and 

assemblage of parasitoids. Exorista larvarum is more 

prevalent in the willow-poplar (Salix spp.-Populus

spp.) forest type where its alternate host, the satin moth 

(Leucoma salicis), is present (Sabrosky and Reardon 

1976). Compsilura concinnata causes gypsy moth 

mortality in oak forests near orchards where many 

alternate hosts are available (Sabrosky and Reardon 

1976). Brachymeria intermedia and Cotesia melanoscela 

prefer dry sites, while Parasetigena silvestris and

Phobocampe unicincta are commonly recovered from 

mesic sites (Skinner et al. 1993).

Hymenoptera (Wasps)

Brachymeria intermedia (Nees)

This chalcidid was introduced from Europe and 

repeatedly released for biological control between 1908 

and 1963 (Clausen 1978, Hoy 1976). It is rarely found 

in low density gypsy moth populations (Williams et al. 

1993). This parasitoid prefers open sunny areas (Hoy 

1976) and exhibits delayed density dependence, causing 

high gypsy moth mortality in outbreak populations 

where defoliation is severe (Ticehurst et al. 1978, 

Williams et al. 1993). Brachymeria intermedia is a pupal 

endoparasitoid with a bias for male gypsy moth pupae 

(Fuester and Taylor 1996). Known to be polyphagous, 

Dowden (1935) notes that this species may also be 

a hyperparasite, attacking puparia of the tachinid 

parasitoids Compsilura concinnata and Exorista 

larvarum in the laboratory, though this is thought to be 

a rare occurrence in the field.

Dowden (1935) found that in Europe, B. intermedia

completes one generation on the gypsy moth and a 

second generation on other lepidopteran hosts; due to the 

comparatively delayed host development in New England, 

this parasitoid likely completes a first generation on other 

lepidopteran hosts and a second generation on gypsy 

moth. The adult emerges from a misshapen exit hole, 

typically between the middle to the anterior end of the 

host pupal shell (Dowden 1935). The adult female (Fig. 9) 

overwinters with other mated females found clustered in 

the litter and under loose bark (Dowden 1935). Adults are 

small and stout with a large hind femur and a body length 

of 5 mm (Howard 1889). For a key to Brachymeria of the 

United States and Canada refer to Burks (1960).

Cotesia melanoscela (Ratzeburg)

This braconid parasitoid completes two generations 

a year. It was introduced to the United States in 1912, 

specifically for biological control of the gypsy moth 

(Burgess and Crossman 1929). Discovered in North 

Africa and India, this oligophagous species thrives in 

xeric sites (Liebhold and Elkinton 1989, Skinner et al. 

1993). While Burgess and Crossman (1929) considered 

this parasitoid to be one of the most valuable enemies 

of the gypsy moth, its abundance is limited by 

host availability and a number of hyperparasitoids 

(Muesebeck and Dohanian 1927). Females lay from 

50-1,000 eggs, depositing a single egg in first or second 

instar larvae. During egg deposition, the females 

also insert a virus (Family Polydnaviridae; Genus 

Bracovirus), which prevents molting and suppresses 

the immune response of the host (Lavine and Beckage 

1995, Shelby and Webb 1999, Stoltz et al. 1988, Stoltz 

and Xu 1990). The small parasitoid larva emerges 

from a parasitized (generally living) gypsy moth larva 

Figure 9.—Brachymeria intermedia adult on gypsy moth 

pupa. Photo by Pedro Barbosa, University of Maryland, 

used with permission.

2 mm
9

and spins a white cocoon (about the size of a grain 

of rice) nearby (Fig. 10A). The virus often keeps the 

gypsy moth larva alive for many days after parasite 

emergence (Fleming 1992). Therefore, field-collected 

small instars that live for many days in rearing without 

molting should be inspected closely for exit holes of C. 

melanoscela. A second generation of wasps will emerge 

from these cocoons in 4-11 days to attack second or 

third instar host larvae (Reardon et al. 1973). Again, 

a parasitoid larva emerges from the parasitized gypsy 

moth larva and spins a white to yellowish cocoon 

attached to tree trunks and the undersides of branches. 

These overwintering cocoons produce adult wasps 

during the following April or May, when gypsy moth 

eggs are at peak hatch (Fuester et al. 2014). Adult C. 

melanoscela are shiny and black with a body length 

of 2.5-3 mm (Fig. 10B). See Crossman (1922) for a 

detailed description of this parasitoid.

Phobocampe unicincta (Gravenhorst)

Phobocampe unicincta is a univoltine, ichneumonid 

parasitoid released in Massachusetts between 1907 and 

1912 to help control the gypsy moth (Clausen 1956). 

Rates of parasitism are typically low but can be heavier 

in dense woodlands (Gupta 1983). This parasitoid 

(historically referred to as Hyposoter disparis) is of 

minimal value for controlling the gypsy moth due to 

heavy hyperparasitism, high overwintering mortality, 

and high mortality of egg and first instar larvae due to 

phagocytosis by the host immune system (Muesebeck 

and Parker 1933). This parasitoid oviposits in first and 

second instar larvae. Females can produce in excess of 

1,200 eggs over a period of 5-8 weeks (Clausen 1956). 

A week after oviposition, the egg hatches inside the 

host larva, and the larval parasitoid grows while feeding 

within the host for nearly a month. When the host 

reaches the fourth instar it dies and the fully grown 

wasp larva emerges as a large, slimy-looking, whitishgray larva. The larval parasitoid spins a hardened, dark, 

oval cocoon of 6 mm x 4 mm on a nearby leaf or twig. 

When formed, the dark cocoon has a very distinctive 

light band around the middle (Fig. 11A). This loosely 

attached cocoon then falls to the ground where 

pupation takes place and it remains over the winter, to 

emerge the next spring. Adults have a short thorax with 

an elongated abdomen. The body length is 4-7 mm 

(Gupta 1983, Viereck 1911) and the color is black with 

reddish-yellow legs (Fig. 11B).

Pimpla disparis Viereck

This ichneumonid wasp is a polyphagous endoparasitoid 

of lepidopteran pupae (Fuester et al. 1989). Native to 

Asia, this species was released between 1972 and 1987 

for biological control of gypsy moth in New Jersey, 

Figure 10.—Cotesia melanoscela (A) cocoon with 

gypsy moth larva and (B) adult. Photos by George 

Boettner (cocoon), University of Massachusetts and 

Michael Sharkey (adult), University of Kentucky, 

used with permission.

Figure 11.—Phobocampe unicincta

(A) cocoon and (B) adult. Photos 

by Yong-Lak Park, West Virginia 

University, used with permission. 1 mm

A B

1 mm

1 mm
10

Pennsylvania, and West Virginia (Reardon 1981, 

Schaefer et al. 1989). Since 2000, it has been commonly 

recovered from coastal Massachusetts and Maine.1 In 

the United States its primary hosts are gypsy moth, 

browntail moth (Euproctis chrysorrhoea), bagworm 

(Thyridopteryx ephemeraeformis), and tent caterpillars 

(Malacosoma americanum); in Asia it also attacks 

pupae of fall webworm, Hyphantria cunea (Fuester et 

al. 2014). Pimpla disparis is multivoltine, completing a 

single generation in 25-32 days (Schaefer et al. 1989) and 

overwintering as a mature larva in the host pupa. The 

adult parasitoid emerges from the pupal case by chewing 

a hole from which it exits (Hrabar et al. 2012). Body 

length is 12-15 mm (Gupta 1983, Viereck 1911). Adults 

are black with white pubescence, and the hind leg has 

a black coxa, reddish-yellow femur, a dark tibia and a 

dark tarsus (Fig. 12).

Theronia atalantae fulvescens (Cresson)

Theronia atalantae fulvescens is a primary pupal 

parasitoid of gypsy moth and a secondary parasitoid 

of ichneumonids and tachinids. This ichneumonid 

wasp is a hyperparasitoid of both Pimpla disparis and

Brachymeria intermedia (Fuester et al. 1997, Schaefer 

et al. 1989). While parasitism rates can be as low as 

2 percent in gypsy moth pupae (Howard and Fiske 

1911), up to four times as many pupae are stung and 

killed by T. atlantae (Campbell 1963). This species is 

native, but specimens from Europe have been released 

in New England prior to 1910 to control gypsy moth 

(Townes 1940). It is polyphagous and seems to prefer 

1 Elkinton, J.; Boettner, G. 2015. Personal communication. 

Professor and lab technician, UMASS-Amherst, 

Department of Environmental Conservation, 250 Natural 

Resources Rd., Amherst, MA 01003.

shady conditions (Campbell 1963) and dense forests 

(Gupta 1983). This wasp lays a single egg in either 

a prepupa or a newly formed pupa (Townes 1940). 

Transformation from egg to adult only takes 14-18 days 

(Townes 1940). Theronia atlantae fulvescens has 2-3 

generations a year and overwinters in the host pupa. 

Adults may be present through the spring and summer, 

but in the fall only females persist (Townes 1940). Adult 

females overwinter under the bark of logs suspended 

across creeks (Dasch 1971). The size of the adult varies 

depending on the host, with body length ranging from 

6 to 13 mm and forewing lengths ranging from 4 to 

11 mm (Gupta 1983) (Fig. 13). This species is easily 

identified by pure orange hind legs.

Diptera (Flies)

Blepharipa pratensis (Meigen)

Introduced to North America between 1905 and 1933 

for biological control of the gypsy moth (Grenier 

1988), these tachnid flies are most abundant when 

host populations are at low to intermediate densities 

(Ticehurst et al. 1978). This univoltine, oligophagous 

parasitoid aggregates on gypsy moth-damaged leaves 

where females lay tiny, microtype eggs. Oviposition 

typically occurs when gypsy moth larvae are in the 

late third or early fourth instars (Godwin and Odell 

1981, Williams et al. 1992). The eggs hatch after they 

are consumed by gypsy moth larvae. The maggot 

bores through the gut wall and into a longitudinal 

intersegmental muscle where it remains until the gypsy 

moth larva pupates. Then, the maggot completes its 

development and leaves its host to form a puparium 

in the soil where it will overwinter. Puparia are 

roughly 6-10 mm in length and can be identified by 

Figure 12.—Pimpla disparis adults. Photo by Michael Hrabar, 

Simon Fraser University, via flickr, used with permission.

Figure 13.—Theronia atalantae fulvescens adult. Photo by 

James Lindsey at Ecology of Commanster (CC BY-SA 2.5 

or 3.0), via Wikimedia Commons.

3 mm

2 mm
11

the distinctive posterior area of the spiracular plates 

(Fig. 14), which can easily be seen under a dissecting 

microscope. The puparium of Blepharipa pratensis 

has a large, prominent subspiracular protuberance 

with a triangular shaped ridge-like extension dorsally 

between and ventral to the spiracular plates (Sabrosky 

and Reardon 1976). Adults (Fig. 15) emerge to mate the 

following spring and begin oviposition. Adults are 10-

14 mm long, by far the largest and most robust of the 

gypsy moth parasitic flies occurring in North America 

(Sabrosky and Reardon 1976).

Compsilura concinnata (Meigen)

This generalist tachinid parasitoid was introduced 

for gypsy moth control as early as 1906 (Culver 1919, 

Sanchez 1995) and is well established throughout much 

of the northeastern and north central United States. It 

is common in low density gypsy moth populations and 

tends to prefer late instar larvae (Skinner et al. 1993). 

Compsilura concinnata has at least four generations 

per year, although only one of these generations (and 

a partial second generation) attacks gypsy moths. This 

species is a likely culprit in the decline of native giant 

silk moths of the northeastern United States (Elkinton 

and Boettner 2004). This fly is ovoviviparous, inserting 

between one and five larva(e) into the host’s midgut or 

body cavity (Culliney et al. 1992). A larva will complete 

two molts in roughly 2 weeks, emerging from the host 

as a white maggot, then forming a smooth, reddish 

brown puparium (Fig. 16). After about 10 days, adult 

flies emerge. Gould et al. (1990) demonstrated that 

this fly can be the dominant parasite of small patches 

of outbreaking gypsy moth larvae. However, the fly 

is dependent on rich forest habitats with abundant 

alternate hosts to complete its multiple generations per 

year, which limits its ability to respond to gypsy moth 

numbers in some years.

If rearing field-collected gypsy moth larvae individually 

in cups, at least one puparium will be found from 

any larva parasitized by C. concinnata, and from this 

an adult will emerge in a few weeks (Fig. 17). Adult 

length is 7-8.5 mm (Sabrosky and Reardon 1976), and 

the puparium is 6.5 mm long (Koch and Hutchinson 

2017) with a rounded posterior end in profile. The 

spiracular plates (Fig. 18), which have straight or very 

slightly curved slits, are vertical to the long axis of the 

puparium with little to no protuberance above the 

surrounding surface (Sabrosky and Reardon 1976).

Figure 14.—Line drawing of Blepharipa pratensis puparium. 

Illustration adapted from Figure 69 of Sabrosky and Reardon 

(1976) and Simons et al. (1979) by Laura Blackburn, USDA Forest 

Service.

Figure 15.—Blepharipa pratensis adult. Photo by Christophe 

Quintin (CC BY-NC 2.0 Generic).

Figure 16.—Puparium of Compsilura concinnata. Photo by © 

2010 Malcolm Storey, via www.bioimages.org.uk, used with 

permission.

4 mm

1 mm
12

host, and creates a puparium to pass the winter in the 

soil. In profile, the puparium (Fig. 20) has a rounded 

posterior end that is slightly depressed above the 

apex, with spiracular plates on the depressed surface 

above the apex and a bulge between the anus and the 

subspiracular protuberance (Sabrosky and Reardon 

1976). In larger pupae, the distance between the 

spiracular plates and the anal opening is much longer in 

P. silvestris than in C. concinnata, making these species 

easy to distinguish. Adult flies are 8-12.5 mm long and 

emerge in May and June (Fig. 21).

Figure 20.—Line drawing of Parasetigena silvestris

puparium. Illustration adapted from Figure 53 of Sabrosky 

and Reardon (1976) and Figure 63 of Simons et al. (1979) 

by Laura Blackburn, USDA Forest Service.

Parasetigena silvestris (Robineau-Desvoidy)

This tachinid fly is a univoltine, oligophagous larval 

parasitoid that was introduced for gypsy moth control 

in 1910 (Kenis and Vaamonde 1998). It is one of 

the most important natural enemies of gypsy moth 

throughout its global range (Elkinton and Liebhold 

1990, Kenis and Vaamonde 1998), exhibiting peak 

parasitism when host populations decline following 

an outbreak. Females lay large, white eggs on the 

dorsum of middle to late instar larvae (Fig. 19). The 

eggs hatch after 2 days, and the young maggot enters 

the host’s body, forming a respiratory funnel at the 

point of entry. This larval stage lasts from 16 to 35 days 

(Fuester et al. 2014). Then, the larva emerges from the 

fully-grown larval host, or occasionally from the pupal 

Figure 17.—Compsilura concinnata adult. Photo by Joyce 

Gross, UCB, via Bugwood.org.

Figure 18.—Line drawing of Compsilura concinnata puparium. 

Illustration adapted from Figure 61 of Sabrosky and Reardon 

(1976) and Figure 66 of Simons et al. (1979) by Laura 

Blackburn, USDA Forest Service.

Figure 19.—Parasetigena silvestris eggs attached to host, 

a late instar gypsy moth larva. Photo by Gyorgy Csoka, 

Hungary Forest Research Institute, via Bugwood.org

1 mm
13

Egg Parasitoids

Two egg parasitoids are established in North America: 

Ooencyrtus kuvanae (Howard) (Hymenoptera: 

Encyrtidae) and Anastatus japonicus Ashmead 

(Hymenoptera: Eupelmidae). Ooencyrtus kuvanae is the 

most common, often causing 10-40 percent of gypsy 

moth mortality, with higher mortality rates occurring 

in dense host populations (see Brown [1984] for a 

review). Ooencyrtus kuvanae has also been reported 

as a hyperparasite of Cotesia melanoscela, though this 

occurred at low rates (<0.2 percent) (Crossman 1925, 

Weseloh 1978). Anastatus japonicus is the more cold 

tolerant of these two egg parasitoid species (Leonard 

1974) and has the ability to parasitize at least 35 percent 

of gypsy moth eggs (Crossman 1925). Details on 

distinguishing characteristics of these egg parasitoids 

can be found in older publications (Burgess and 

Crossman 1929, Crossman 1925, Parker 1933, Tigner 

1974). For a key to Anastatus, including A. japonicus, 

refer to Narendran (2009).

UNCOMMON PARASITOIDS

Hymenoptera (Wasps)

Aleiodes indiscretus (Reardon)

This braconid is native to India where it is a parasitoid 

of Lymantria obfuscata, a relative of the gypsy moth 

(Shaw et al. 2013). In the late 1960s, it was introduced 

to the eastern United States for biological control of 

L. dispar (Reardon 1970) and can now be found from 

Massachusetts south to Maryland (Shaw 2006). Aleiodes 

indiscretus is an oligophagous endoparasitoid of second 

and early third instar larvae of Lymantria and Dasychira

species. As with all species of Aleiodes, the dead host 

becomes dried and preserved (i.e., mummified). The 

parasitoid larva pupates and overwinters inside the 

mummified larval host. The adult parasitoid emerges 

from the penultimate larval instar through a circular 

postero-dorsal hole (Reardon 1981, Shaw 2006). Adults 

are honey-yellow (Fig. 22) with a black ocellar triangle 

and a body length of 6-8 mm (Shaw 2006). Females 

have fulvous antennae and a slightly larger body than 

males, which have fuscous antennae (Reardon 1970).

Monodontomerus aereus Walker

This torymid pupal parasitoid was imported from 

Europe between 1905 and 1911 (Clausen 1956). It is 

oligophagous and univoltine, developing in the pupae of 

gypsy moth, browntail moth, and several native 

Lepidoptera. Monodontomerus aereus is also a secondary 

parasitoid of Hymenoptera and Diptera associated with 

gypsy moth and browntail moth (Clausen 1956, Fuester 

et al. 2014, Muesebeck 1931). While development is 

internal in a lepidopteran pupa, development in a 

tachinid puparium or braconid cocoon is external 

(Muesebeck 1931). This hyperparasitic habit likely causes 

more harm than good, though it has not been found in 

sufficient numbers to impact either the primary or 

secondary host (Clausen 1956, Hoy 1976). The adults 

overwinter in old cocoons and webs of browntail moths, 

with only females surviving winters (Muesebeck 1931). 

Figure 21.—Parasetigena silvestris adult. Photo by Patrick 

Derennes, used with permission.

Figure 22.—Aleiodes indiscretus parasitizing a gypsy 

moth larva. Photo by Scott Bauer, USDA. 

2 mm

2 mm
14

Females break hibernation in mid-April to mid-May and 

oviposit in June when a host cocoon, puparium, or pupa 

is found (Muesebeck 1931). A single female may lay up 

to 350 minutely-spined, grayish-white eggs over a 

two-month period. Eggs are 0.6-0.65 mm long by 0.2 

mm wide at their widest and tapered at both ends. The 

time from egg deposition to adult emergence ranges 

between 18 and 24 days. Up to 24 adults can emerge 

through a single, mostly circular hole in the host’s pupal 

envelope. Mating occurs before hibernation, though it is 

also possible for this species to reproduce 

parthenogenetically, in which case all offspring will be 

males (as is typical of all Hymenoptera). Female length 

is 1.5-3.5 mm, with males slightly smaller in size 

(Muesebeck 1931). Adults have a greenish-black head, 

thorax, and abdomen with brown tibiae and tarsi (Fig. 

23). See Muesebeck (1931) for life history and biological 

information on this parasitoid/hyperparasitoid.

Pimpla pedalis Cresson

This wasp is multivoltine and native to the United 

States. Pimpla pedalis is polyphagous and commonly 

uses tent caterpillar larvae as hosts, although historically 

it has been listed as a parasitoid of gypsy moth (Gupta 

1983). Even though parasitism of the gypsy moth by P. 

pedalis is rare, this wasp still kills many host prepupae 

and pupae through stinging alone (Campbell 1963). 

Pimpla disparis and P. pedalis (Figs. 12 and 24) are easily 

distinguished by their middle and hind coxae, which 

are orange in P. pedalis and black in P. disparis. Adults 

are 12-15 mm in length with a fore wing length of 8-12 

mm (Gupta 1983).

Itoplectis conquisitor (Say)

Native to the United States, this is a primary parasitoid 

of lepidopterans. This parasitoid is multivoltine 

and highly polyphagous, occasionally acting as a 

hyperparasitoid (Moser et al. 2008). Similar to Pimpla

and Theronia, this endoparasitoid 

readily stings gypsy moth pupae, though 

it rarely develops in them (Campbell 

1963). Females readily feed on the 

hemolymph or body fluids of their host, 

which leads to host mortality (Leius 

1961). Itoplectis conquisitor prefers an open canopy 

and is positively associated with highly defoliated 

areas (Campbell 1963). Adults are black with a banded 

appearance on the hind leg and abdomen; the legs are 

mostly reddish-brown while the hind tibia has a broad 

yellowish-white band (Gupta 1983), and the tarsus is 

black and white striped (Fig. 25). The size of an adult is 

typically 9-14 mm, though some specimens are shorter 

(5-6 mm).

Figure 25.—Itoplectis conquisitor adult (A) 

dorsal view and (B) side view. Photo by 

Gerald J. Lenhard, Louisiana State University, 

via Bugwood.org. 

Figure 24.—Pimpla pedalis adult. Photo by KNWR Entomology 

Collection (CC0), via Wikimedia Commons.

Figure 23.—Monodontomerus aereus adult. Photo by Josef 

Dvořák, via Biolib.cz, used with permission.

0.5 mm

2 mm

A B

3 mm 2 mm
15

Perilampus hyalinus Say

This is a primary parasitoid of some species of sawflies 

in the genus Neodiprion and a hyperparasitoid in 

association with Lepidoptera and Orthoptera. In 

gypsy moth and brown-tail moth, Perilampus hyalinus 

is a hyperparasitoid of the tachinids Compsilura 

concinnata, Exorista mella, and Lespesia frenchii, and 

the braconid Cotesia melanoscela. In 1912, P. hyalinus 

was reported as being regularly reared from cocoons 

and puparia of primary parasitoids (Smith 1912), yet 

today these species are not found throughout the gypsy 

moth range. These hyperparasitic adults generally 

emerge from their host puparia 4 to 7 days after the 

typical emergence of the host parasitoid (Rees 1973). 

Females can produce up to 367 eggs, ovipositing on 

foliage at least 4 days following emergence (Tripp 

1962). Eggs are elongate with rounded ends and are 

0.3 mm long by 0.07 mm wide (Tripp 1962). Planidia 

(highly mobile first instar larvae) emerge 8-10 days 

later and are less than 0.3 mm in length (Tripp 1962). 

Planidia seldom move far from their oviposition site, 

where they stand erect and wait to make contact with 

a host. Once they have found a host, they enter the 

integument in the soft area between the segments 

in search of a parasitoid on which to complete their 

development (Smith 1912). Perilampus hyalinus

completes four instars, and the pupa develops inside 

the host. Adults emerge from puparia or cocoons 

through a jagged hole roughly 1.9 mm in diameter. 

Adults are metallic green and 5 mm in length (Rees 

1973) (Fig. 26).

Diptera

Sarcophagidae

Referred to as scavengers, some sarcophagids are 

known to attack gypsy moth pupae already parasitized 

by ichneumonids (Campbell 1963). Arachnidomyia 

aldrichi Parker exhibits a delayed density-dependent 

response, increasing their population prior to host 

population collapse (Brown 1938). In one study, 

Campbell (1963) found that for 33-46 percent of gypsy 

moth pupae initially parasitized by ichneumonids, 

sarcophagids successfully emerged. This native, 

univoltine fly larviposits in prepupae and pupae of 

the forest tent caterpillar larva, but may also utilize 

the gypsy moth, satin moth, and spruce budworm 

(Choristoneura fumiferana) (Dodge 1961). Yellowishwhite maggots emerge from the host pupa 8-10 days 

after parasitism and then drop to the ground (or 

Figure 26.—Perilampus hyalinus adult. Photo by John Rosenfeld, 

via BugGuide.net, used with permission.

Figure 27.—Line drawing of Arachnidomyia aldrichi puparium. 

Illustration adapted from Figure 1 of Sippell (1961) by Laura 

Blackburn, USDA Forest Service.

burrow into the diet in the cup) to form a puparium 

and hibernate. Arachnidomyia aldrichi is easily 

identified in the maggot and puparial stage by the 

deep posterior depression where the stigmatal slits 

are located (Fig. 27). The spiracular slits are almost 

straight and nearly vertical in position (Tigner 1974). 

Adults measure 8-10 mm in length (Hallock 1940), 

and, as is characteristic of many Sarcophagidae, there 

are three prominent longitudinal stripes on the thorax 

1 mm
16

(Fig. 28). Another sarcophagid species that has been 

reared from gypsy moth late instar larvae and pupae 

is Agria housei Shewell (Tobin and Hajek 2012), and 

additional species of sarcophagids likely associate 

with gypsy moths and attack wounded and moribund 

individuals.2

Exorista larvarum (Linnaeus) and Exorista mella

(Walker)

Exorista larvarum (Fig. 29) is native to the Palearctic 

region and was released in New England as early 

as 1906 for biological control of the gypsy moth 

(Clausen 1956). This species is often confused with the 

native E. mella (Fig. 30). Both species of Exorista are 

multivoltine and polyphagous. Similar to Parasetigena 

silvestris, Exorista species lay eggs directly on the 

surfaces of late-instar host larvae (Sellers 1953). These 

eggs hatch and the maggots immediately burrow 

into the host’s body. However, E. larvarum forms 

its puparium outside the skin of the host, while E. 

mella forms its puparium inside the skin of the host 

(Sellers 1953). The puparia of both Exorista species

have a rounded posterior end, which is somewhat 

depressed above the apex, with very little elevation of 

the spiracular plates above the surrounding surface. 

The spiracular plates, located well above the apex of 

the puparium, have three slits each on a well-defined 

ridge (Fig. 31). Adults emerge roughly 15 days after 

the puparium is formed, nearly 1 month following 

oviposition. Adult size ranges from 5 to 13 mm, 

depending on the size of the host (Adam and Watson 

1971).

Lespesia frenchii (Williston)

Lespesia frenchii is another rarely occurring parasitoid 

that is native to the United States. This multivoltine 

and highly polyphagous parasitoid uses both gypsy 

moth and forest tent caterpillar larvae as hosts. Lespesia 

frenchii exhibits high rates of parasitism in fragmented 

forests and along the forest edge (Roth et al. 2006). 

Puparia are 8.5 mm long and have strongly sunken, 

sinuous spiracular slits with distinct loops (Fig. 32) 

(Sabrosky and Reardon 1976). Adults are 7-9 mm long 

(Fig. 33). For more information, see Rees (1973) or 

Sabrosky (1980).

2Pape, T. 2012. Personal communication. Associate 

Professor and curator, Natural History Museum of 

Denmark, Universitetsparken 15, 2100 København Ø, 11, 

Building: 02-4-411 Denmark.

Figure 30.—Exorista mella, 12 mm adult male. Photo by Ombor 

Mitra, Agriculture and Agri-Food Canada, via TachImage, used 

with permission.

Figure 29.—Exorista larvarum, adult. Photo by Janet Graham 

(CC-BY 2.0), via Wikimedia Commons.

Figure 28.—Arachnidomyia species, adult. Photo by Joseph 

Berger, via Bugwood.org.

1 mm

3 mm

2 mm
17

GYPSY MOTH COLLECTION AND 

REARING TECHNIQUES

Field Collections

Larvae can be collected in a number of ways, and two 

methods are outlined in this guide. The first method 

is to search for dead larvae. The shape of the cadaver 

is likely to reveal the underlying cause of death, so 

be sure to note this while collecting specimens. The 

second method is to collect live larvae and rear them 

individually in the laboratory (see section below 

on rearing larvae). While rearing, check frequently 

to detect larval death, after which a parasitoid 

might emerge or the cadaver may be examined 

microscopically for the presence of pathogens.

When collecting gypsy moth life stages, it is important 

to take time of year and behavioral characteristics into 

consideration. Gypsy moth larvae hatch in early spring, 

roughly coinciding with oak bud break, and their 

development is strongly influenced by temperature. 

In the northeastern United States, larvae are generally 

found from May to early July. Newly hatched larvae 

may remain near the egg mass from which they 

hatched if emergence occurs during rainy weather 

or if temperatures are below 7 °C (Leonard 1981). 

Larvae then ascend trees, where aerial dispersal, called 

ballooning, occurs as they release silken threads and 

are blown on the wind (McManus 1973).

Gypsy moth larvae in the early instar stages are 

the most specific when it comes to host selection, 

preferring Quercus, Populus, and Salix species 

Figure 31.—Line drawing of Exorista mella puparium. Illustration 

adapted from Figure 72 of Greene (1922) and Figure 58 of 

Sabrosky and Reardon (1976) by Laura Blackburn, USDA Forest 

Service.

Figure 32.—Line drawing of Lespesia frenchii puparium. 

Illustration adapted from Figure 10 of Sippell (1961), Figure 72 

of Sabrosky and Reardon (1976), and Figure 69 of Simons et al. 

(1979) by Laura Blackburn, USDA Forest Service.

Figure 33.—Lespesia frenchii, 8.6 mm adult male. Photo 

by A.J. Fleming, Agriculture and Agri-Food Canada, via, 

TachImage, used with permission.

1 mm
18

(Elkinton and Liebhold 1990). Except in outbreak 

populations, larvae feed during the day, mainly in early 

morning and late afternoon (Leonard 1981). During 

periods of inactivity, early instars can be found along 

the midrib on the undersides of leaves.

As the larvae grow, their feeding behaviors change. 

Late instars crawl down tree trunks, where they seek 

darker daytime resting sites, such as under the base of 

limbs, in bark crevices, and under the leaf litter or on 

the underside of objects on the forest floor. Late instars 

can be found resting on a wide variety of host and 

nonhost trees, and pupae are typically found in these 

same locations (Elkinton and Liebhold 1990). For an 

extensive list of gypsy moth preferred host species, see 

Liebhold et al. (1995).

Behavior of gypsy moth larvae can be dependent upon 

the density of the population. Campbell (1967) and 

Semevsky (1971) suggest that first instars from highdensity populations have higher rates of dispersal than 

larvae from low-density populations. Late larval instars 

in low-density populations are nocturnal, feeding 

during the night and spending the day in resting sites 

on the ground or in protected places on the trunks of 

trees. In high-density populations, late-larval instars are 

cathemeral, feeding in the tree canopy during both the 

day and night, only descending from the canopy when 

foliage no longer remains (Lance et al. 1987). All of 

these behaviors should be considered when collecting 

specimens.

Locating larvae can be a challenge depending on the 

density of the local population. Look for clues such as 

a build-up of frass pellets on the ground or damaged 

leaves on preferred species of shrubs or trees. Focus 

your search on new leaves, saplings, and preferred 

plants along a forest edge (Wagner et al. 1997). During 

the day, early instars will be in the foliage or on the 

undersides of twigs and branches if it is raining, while 

later instars will often be on tree trunks. Finding early 

instar larvae in stands with mature trees can be difficult 

because of the canopy height, so in this case, search for 

larvae on preferred understory shrubs.

In areas of high gypsy moth density, “beating” can 

be an efficient method for collecting early instar 

larvae from trees and shrubs. Spread a sheet over the 

ground and use a bat or sturdy stick to rap against 

a series of branches over the sheet, thus dislodging 

larvae. Recording the host species can be of value, and 

focusing your collection efforts on one tree species at a 

time will help you to keep more accurate records.

In areas of low to moderate gypsy moth density, it is 

often efficient to collect late instar larvae and pupae 

under burlap bands that provide daytime resting sites. 

Encircle a tree at chest height with a strip of burlap 

20 to 25 cm in width; staple along the top edge and 

cut vertical strips to allow the flaps to be lifted while 

inspecting for larvae. It should be noted that collecting 

larvae from bands may provide a biased sample of 

larval mortality agents because larvae resting beneath 

burlap bands will not crawl to the leaf litter during the 

day. Thus, their behavior is changed if burlap bands are 

used, and larvae then avoid resting under the leaf litter, 

a major site where infection by fungal pathogens, such 

as Entomophaga maimaiga, can occur.

Photographs may be taken of the insect in its natural 

setting, and recording notes about the tree species or 

substrate where the sample is collected from may also 

be desired. Field samples should be placed in individual 

containers. If live larvae are collected, place them in a 

container with an artificial diet (see section below on 

rearing larvae). If a cadaver or pupa is found attached 

to foliage or a twig, use a knife or pruners to remove the 

sample while it is still attached to its natural substrate.

Rearing Gypsy Moth Larvae

Gypsy moth larval specimens should be placed 

individually in closed containers and kept at 

approximately 22 °C at low to moderate moisture 

levels. Do not rear larvae at constant temperatures 

over about 25 °C because larvae may not tolerate high 

temperatures. Also, take care to store containers in 

the shade because exposure to direct sunlight can 

cause excessive heating. Live larvae should optimally 

be placed in 30 ml plastic cups containing an artificial 

diet. See Bell et al. (1981) for a recipe, or use a readyto-prepare gypsy moth artificial diet that is available 

commercially. When rearing larvae on an artificial diet, 

care should be taken not to touch the diet or introduce 

contaminants that can lead to fungal or bacterial 

growth. When larvae pupate, transfer day-old pupae 

to empty diet cups to avoid excessive moisture from 

the diet medium. Newly molted pupae are delicate and 

should be handled with care.

Larvae can also be kept in containers with live foliage. 

Foliage should be placed in a vial of water packed 

with cotton to anchor the stem or branch, and leaves 
19

should be replaced regularly (and checked daily) so that 

acceptable foliage is available. However, providing fresh 

leaves may not be efficient when rearing numerous 

samples or when parasitoid emergence requires a long 

waiting period. It is also possible that foliage may be 

contaminated with a virus or other pathogens.

For the 10 days after collection, all larvae should be 

checked for death daily. After larvae die, make sure that 

the larval container is at high humidity for 3 days to 

facilitate conidial production by E. maimaiga. Create 

a high humidity environment by closing the container 

with the artificial diet or by adding a piece of wet 

paper towel inside of a closed container. Larval death 

due to fungal infections usually occurs within one 

week of collection, although this can extend to 10 days 

(depending on rearing temperature and conditions). 

After the first 10 days, if a larva has not died, checks 

should be made weekly to look for host death. Because 

larval deaths caused by viral infections may occur for 

up to a month after collection, continue rearing and 

checking the larva for at least 1 month.

When checking rearings also look for parasitoids, 

making certain to examine the underside of the cup 

for any parasitoid larvae that may have emerged and 

burrowed into the artificial diet. If mold starts to grow 

on the artificial diet, transfer the specimen to a clean 

diet cup. It can take several weeks before a parasitoid 

emerges to form a cocoon (wasp) or a puparium (fly). 

Adults of some gypsy moth parasitoids emerge from 

cocoons or puparia in the following year, so pupal 

characteristics are a useful feature for parasitoid 

identification. Approximately 1 month after collecting 

larvae or pupae, all samples should be stored in a 

refrigerator and dissected to search for dead parasitoid 

larvae or pupae and microscopically examined for 

pathogens.

PATHOGEN IDENTIFICATION

Laboratory Dissections

To determine the cause of larval death, initially note the 

external condition of cadavers in the field. Both fungal 

and viral pathogens are more commonly found in later 

instar hosts. Bodies of larvae that die from LdMNPV 

are often loosely attached to surfaces (like tree trunks) 

by their anterior prolegs only (often creating an 

inverted V; Fig. 7). The cadavers are internally liquefied, 

so once the cuticle breaks, the cadaver drips and rapidly 

disintegrates. In contrast, late instar larvae killed by the 

fungus E. maimaiga will often hang vertically on the 

tree trunk with heads downward and prolegs at a 90o 

angle (Fig. 4), and their bodies will eventually appear 

dry and stiff (although if recently dead, the internal 

contents of these cadavers are not solid). However, the 

vertical orientation of cadavers alone cannot be used to 

identify the cause of death (Hajek and Roberts 1992). 

Usually within several days of larval death, cadavers 

fall to the ground and decompose, releasing spores 

or occlusion bodies which overwinter in the soil. A 

cadaver can contain either a single pathogen or both 

the fungus and the virus. Thus for accurate diagnosis, 

microscopic examination is necessary.

The size, structure, and odor of pathogens provide 

identifying characteristics. Spores of E. maimaiga can 

easily be seen since they are larger than viral occlusion 

bodies. Conidia may be produced by all instars. These 

ephemeral pear-shaped spores are approximately 20 

μm x 25 μm (Hajek and Snyder 1992) and are typically 

visible microscopically at 100x to 200x. Viral occlusion 

bodies are comparatively much smaller but readily 

visible under a compound microscope at 200x to 

400x. Viral structures vary between 1 and 10 μm in 

diameter and are polyhedral in shape (Hajek 1994). 

Cadavers killed by LdMNPV exude a characteristically 

unpleasant and unique odor, while larvae killed by E. 

maimaiga are typically odorless (Hajek and Roberts 

1992, Koyama 1954).

Equipment

• Dissecting microscope for parasitoids (6x-50x 

such as Wild M5 or M5A)

• Compound microscope with phase contrast

• Gloves

• Lighter

• Alcohol lamp with 95 percent ethanol*

• Small beaker with tap water

• Two dissecting probes 

• Bottle with 1M KOH and a pipet*

• Bottle with distilled water and a pipet*

• Glass microscope slides

• 18 mm x 18 mm cover slips

• Paper towels

* Bottles should be kept closed or covered to prevent 

evaporation.
20

Techniques for Pathogen Identification

1. Put on snug-fitting gloves for sterility.

2. Organize samples for dissection and prepare a 

notepad or other medium for recording results 

next to the microscope.

3. Flame two dissection probes by passing them 

into a flame (e.g., alcohol lamp) for several 

seconds.

4. Dip dissection probes into water to cool (they 

should sizzle). Wipe probes on paper towels to 

clean them.

5. Use both probes to remove a 1 to 2 mm-sized 

piece of cadaver from the cup and place it on a 

microscope slide. If saprophytic mold has taken 

over the sample and artificial diet (Fig. 34), 

look for hairs or the head capsule to identify 

the cadaver. If all you can find is the head 

capsule, transfer it and any surrounding bits of 

cadaver or hairs to the middle of a microscope 

slide. This technique works much better with 

less material rather than more.

6. Add 1-2 drops of distilled water from the water 

bottle to the cadaver piece on the slide (Fig. 35A) 

and gently pulverize the piece with the probes 

until the cadaver piece is homogenized and well 

mixed with the water (Fig. 35B).

7. Return large fragments from the cadaver back 

to the bioassay cup, leaving the smear in water 

on the slide (Fig. 36). Close the cup. Removing 

these larger fragments makes viewing under 

the compound microscope easier.

Figure 35.—Lab dissections may be needed to identify cause of death. (A) Add 1-2 

drops of water onto microscope slide with a dissected piece of a cadaver. (B) Smash 

the cadaver piece with probes to form a homogenized mixture (less material is better). 

Photos by Ruth Plymale, Ouachita Baptist University, used with permission.

Figure 36.—Slide with large fragments of the 

cadaver removed from the cadaver and water 

mixture. To observe viral occlusion bodies, the 

smear between the cover slip and slide must be 

thin and nearly transparent. Photo by Ruth Plymale, 

Ouachita Baptist University, used with permission.

Figure 34.—Gypsy moth cadaver covered in mold 

in a diet cup. Photo by Ruth Plymale, Ouachita 

Baptist University, used with permission.
21

8. Add an 18 mm x 18 mm cover slip by laying it 

on the sample, starting at one side to avoid air 

bubbles (Fig. 37A). Using the 18 mm x 18 mm 

cover slips, three samples will fit onto a normal 

glass microscope slide (Fig. 37B).

9. For typical samples, view under the compound 

microscope with phase contrast at 200x (10x 

eyepiece x 20x objective). Make sure the phase 

ring is on the appropriate phase (20x phase 

setting if you are using the 20x objective). Scroll 

from side to side across the cover slip and move 

the slide backward and forward, making sure to 

see the entire cover slip. Look for resting spores 

and then viral occlusion bodies and conidia 

(but remember that these are very different 

sizes). If only one resting spore is found, take a 

second sample from the cadaver to verify this 

diagnosis. Make sure to clean the dissecting 

probes and work area between samples so that 

the contents of one cadaver are not mistaken for 

the contents of another cadaver.

It can be difficult at high magnification to feel certain 

whether bright dots within the smear are viral 

occlusion bodies or not. Test each sample for LdMNPV 

occlusion bodies by adding 1M KOH and watching 

for the occlusion bodies to dissolve; but note that this 

only works with phase contrast, and doing this at 200x 

magnification is suggested. Add 1 small drop of 1M 

KOH from a small pipet to one side of the cover slip. 

As that drop of KOH moves under the cover slip and 

across the slide, any occlusion bodies that come into 

contact with the KOH will dissolve (i.e., they will turn 

from bright to black and often will then disappear). If 

the KOH does not seem to be moving under the cover 

slip, use a piece of fine tissue at the opposite edge of 

the cover slip to take up the liquid, thereby causing the 

KOH to flow under the cover slip. Watch closely under 

the microscope as the KOH moves across the slide 

in order to observe the dissolution of the occlusion 

bodies. If too much KOH is added, the cover slip gets 

flooded and the virus is washed away, losing the chance 

to see it dissolve, so be frugal with KOH usage. KOH is 

corrosive, so always take care to avoid contact between 

the KOH and the microscope. If you only observe the 

dissolving of a small number of occlusion bodies, take a 

second sample from the cadaver to verify this diagnosis.

Molecular Methods

Molecular methods can also be used to identify the 

pathogens/parasitoids in gypsy moth larvae, but there 

are several issues complicating the molecular analysis. 

For instance, developing the facilities needed for 

molecular analysis is expensive and requires high-end 

instrumentation. Also, molecular analysis is laborious 

and requires technical skill. With living larvae it is 

possible to pulverize individual larvae, extract DNA, 

and then use unique primers for the fungus, virus, and 

the parasitoids as a group. One would then use DNA 

fingerprinting to tell the different parasitoid species 

apart (Bashasab et al. 2006, Chatterjee et al. 2013, 

Greenstone 2006, Landry et al. 1993, Loxdale and 

Lushai 1998).

Dead larvae also require three separate analyses to 

identify fungus, virus, and parasitoids, and several 

issues can complicate the molecular analysis:

1. If the larva dies from E. maimaiga and 

produces only conidia, these spores decompose 

within a short time at room temperature, so 

a sporulating cadaver must be stored in 95 

percent ethanol or storage buffer (such as 

RNAlaterTM stabilization solution) during the 

relatively short time that the fungal cells are 

alive after host death.

Figure 37.—(A) Place the cover slip over the sample from the side to prevent air bubbles. (B) Three samples 

can be placed on one slide. Photos by Ruth Plymale, Ouachita Baptist University, used with permission.
22

2. If the larva dies from E. maimaiga and resting 

spores are only produced, these must be broken 

open in order to reach the fungal DNA. See 

Castrillo et al. (2007) for a method on how to 

break resting spores to harvest E. maimaiga 

DNA.

3. Cadavers need to be pulverized as for living 

larvae to detect viruses or parasitoids. 

4. In some cases, the parasitoid might have left 

the cadaver to pupate in the artificial diet, 

so rearing containers must always be closely 

searched for parasitoid larvae or pupae which 

can then be pulverized for DNA extraction and 

identification with DNA fingerprinting.

Potentially, identification with molecular methods may 

be more sensitive. However, there is uncertainty about 

the relative lengths of time for the different methods, 

and use of molecular methods may be more expensive.

SUMMARY

The introduction of natural enemies against the gypsy 

moth in the United States is likely the most intensive 

biological control effort directed against any individual 

species in the world. Over the past century there have 

been many unsuccessful attempts at biological control 

of gypsy moth. Of the 34 parasitoids introduced to 

control gypsy moth, only 12 are established in North 

America (Fuester et al. 2014), with the majority of these 

natural enemies becoming established prior to 1920 

(Burgess and Crossman 1929, Howard and Fiske 1911). 

Though keys to these parasitoids have been provided 

in the past (e.g., Sabrosky 1980, Sabrosky and Reardon 

1976), they are quite exhaustive and include many 

species that were released in North America but never 

became established. This guide helps to identify the 

most commonly occurring pathogens and parasitoids 

found today within the North American range of the 

gypsy moth.

ACKNOWLEDGMENTS

We thank Ruth Plymale and Patrick Tobin for initiating 

this project. We are grateful to Richard Reardon (USDA 

Forest Service, State and Private Forestry-Washington 

Office) for use of specimens, literature, and a review of 

this guide. We also thank George Boettner (University 

of Massachusetts Amherst), John Stireman (Wright 

State University), and Andrew Liebhold (USDA Forest 

Service, Northern Research Station) for their helpful 

comments in preparation of this document. For 

allowing the use of their images in the publication of 

this guide, we thank Ruth Plymale, Michael Sharkey, 

Michael Hrabar, James Lindsey, Christophe Quintin, 

Malcolm Storey, Joyce Gross, Gyorgy Csoka, Patrick 

Derennes, Scott Bauer, Josef Dvorak, Gerald Lenhard, 

John Rosenfeld, Joesph Berger, Ombor Mitra, Pedro 

Barbosa, Yong-Lak Park, Janet Graham, and A.J. 

Fleming.

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GLOSSARY

apex—the top or highest point of something.

cathemeral—irregularly active through the day and 

night, not falling into the strict definitions of diurnal or 

nocturnal or crepuscular.

cocoon—a papery or silken structure enclosing the pupa.

conidiophore—a specialized hypha upon which conidia 

develop.

conidium (sing.); conidia (pl.)—an asexual 

reproductive propagule formed in any manner that does 

not involve cytoplasmic cleavage. Conidia function as 

organs of reproduction and dissemination. Here, use of 

this term refers to fungi.

coxa (sing.); coxae (pl.)—the basal segment of a leg on 

insects.

diurnal—active mainly during the daylight hours.

endoparasitoid—a parasitoid that lives inside another 

insect and ultimately kills it.

epizootic—an unusually large number of cases of a 

disease within an animal population that is not human.

femur (sing.); femora (pl.)—the segment of an insect’s 

leg that is third from the body.

frass—excrement or debris produced by insects.

fulvous—reddish or brownish yellow; tawny.

fuscous—dark gray or grayish brown in color; dusky.

hemolymph—the circulatory fluid of various 

invertebrate animals that is functionally comparable to 

the blood and lymph of vertebrates.

hyperparasitoid—An insect that is a parasitoid of a 

parasitoid.

larva (sing.); larvae (pl.)—the active immature form 

of an insect, especially one that differs greatly from the 

adult. This is the stage between egg and pupa (e.g., a 

caterpillar, maggot, or grub).

larviposit—to bear and deposit living larvae instead of 

eggs.

maggot—a soft-bodied legless stage that is the larva of a 

dipterous insect (e.g., the housefly).

mesic—a type of habitat with a moderate or wellbalanced supply of moisture.

multivoltine—having more than one generation per 

year.

nocturnal—active mainly during the night.
30

ocellar triangle—a three sided space, sharply defined 

in many insects, on which the ocelli (or simple eyes) are 

located.

oligophagous—feeding on a limited number of foods, 

usually within one taxonomic family of plants or 

animals.

oviposit (verb); oviposition (noun)—to lay eggs, 

especially by means of an ovipositor (a tubular 

structure, usually concealed but sometimes extending 

outside the abdomen, with which many female insects 

deposit eggs).

parasitoid—an insect, and especially a wasp or fly, that 

completes its larval development feeding on the body of 

another insect, eventually killing it, and is free-living as 

an adult.

parthenogenetic (adj.); parthenogenesis (noun)—a 

form of reproduction in which an unfertilized egg 

develops into a new individual; commonly occurs 

among insects and certain other arthropods.

pathogenic (adj.)—causing or capable of causing 

disease.

penultimate—occurring immediately before the last 

one; next to the last.

phagocytosis—the engulfing and usually the 

destruction of non-self matter by individual immune 

system cells that perform this service; this term often 

refers to engulfing and killing microbes that have 

invaded a host.

planidium (sing.); planidia (pl.)—a first-stage larva of 

various parasitic hymenopteran and dipteran insects, 

known as a dispersal stage.

polyphagous—feeding on a broad array of plant or 

animal species.

postero-dorsal—relating to, or situated at, the back of 

the upper surface of an organism or object.

prepupa (sing.); prepupae (pl.)—a stage in the 

development of many insects immediately preceding 

the change to a pupa and usually marked by cessation 

of feeding and reduction of activity.

protuberance—usually a rounded part that sticks out 

from a surface.

pubescence—a covering of fine soft short hairs.

pupa (sing.); pupae (pl.)—the inactive stage in some 

insects, such as true flies and wasps, that falls between 

the larval and adult stages, during which the insect 

typically undergoes complete transformation within a 

protective cocoon or hardened case.

puparium (sing.); puparia (pl.)—a rigid outer shell 

formed from the larval skin that covers some pupae (as 

in dipteran parasitoids discussed above).

rachis (sing.); rachises (pl.)—a bent or sometimes zigzag extension at the tip of a conidiogenous cell produced 

by lateral branching of the elongating extension beneath 

each successive conidium that is formed.

resting spores—a persistent spore created by some 

fungi which has a thick wall in order to survive through 

stressful times, such as drought or periods when 

hosts are not active. It protects the spore from biotic 

(microbial, fungal, viral), as well as abiotic (wind, heat, 

xeric conditions) factors.

spiracular plates—In Diptera, the flattened tip of each 

tube that bears the posterior spiracles (or breathing 

holes) for puparia.

stigmatal slits—narrow breathing pores in fly larvae or 

puparia.

tarsus (sing.); tarsi (pl.)—the distal part(s) of the 

limb(s) of an insect.

tibia—the fourth segment of the leg of an insect, 

between the femur and tarsi.

univoltine—having one generation per year.

viral occlusion bodies—a crystalline protein matrix 

surrounding the virions of some insect viruses. For 

nucleopolyhedroviruses, occlusion bodies are produced 

within the nuclei of host cells while the insect is still 

alive, and the occlusion body protects the infectious 

virions after death of the host.

virion—a complete virus particle that consists of an 

RNA or DNA core with a protein coat and sometimes 

with an external envelope; this is the extracellular 

infective form of a virus.

viviparous—giving birth to live young (not eggs).

xeric—of, relating to, or adapted to a dry environment.
Blackburn, Laura M.; Hajek, Ann E. 2018. Gypsy moth larval necropsy guide. Gen. Tech. Rep. 

NRS-179. Newtown Square, PA: U.S. Department of Agriculture, Forest Service, Northern 

Research Station. 30 p. https://doi.org/10.2737/NRS-GTR-179.

Since the early 1900s, a number of parasitoids have been released for classical biological 

control of the introduced destructive forest insect, Lymantria dispar (gypsy moth), in North 

America. During this time, two pathogens were accidentally introduced. These pathogens 

and several of the parasitoid species are now commonly found in North American gypsy 

moth populations. The aim in creating this guide was to provide laboratory techniques for 

distinguishing between two common pathogens, the fungus Entomophaga maimaiga and 

the gypsy moth multiple nucleopolyhedrovirus, and provide illustrations and images for 

adults, puparia, and cocoons of established gypsy moth parasitoids commonly found in the 

larval or pupal stages of gypsy moth in North America. Gypsy moth collection and rearing 

techniques are also reviewed, and a technical glossary and summary table highlighting the 

affected life stage by gypsy moth parasitoids in North America are included in this guide.

KEY WORDS: Lymantria dispar, pathogens, parasitoids, hyperparasitoids, life table studies, 

cause of death, Entomophaga, Tachinidae, Ichneumonidae

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