MASARYK UNIVERSITY
Faculty of Science
Veronika Michalková
A STUDY OF HOSTS AND BIOLOGICAL ADAPTATIONS OF
TACHINID LARVAE (DIPTERA, TACHINIDAE) IN
CONNECTION WITH HOST-PARASITOID INTERACTIONS.
Dissertation thesis
Supervisor: Prof. RNDr. Jaromír Vaňhara, CSc. Brno 2009
Bibliographic identification
First name and surname of author: Veronika Michalková
Title of dissertation thesis: A study of hosts and biological adaptations of tachinid larvae
(Diptera, Tachinidae) in connection with host-parasitoid interactions.
Title of dissertation thesis in Czech: Studium hostitelů a biologických adaptací larev čeledi
Tachinidae (Diptera) ve vazbě na interakce hostitel-parazitoid.
Study program: Biology
Field of study: Zoology
Supervisor: Prof. RNDr. Jaromír Vaňhara, CSc.
Year of defence: 2009
Keywords: parasitoid, endogenous development, larval morphology, immune response, host
records, catalogue
Keywords in Czech: parazitoid, endogenní vývoj, morfologie larev, imunitní odpověď,
hostitelé, katalog hostitelů
 Veronika Michalková, Masaryk University, 2009
"Nothing shocks me. I'm a scientist."
Indiana Jones
Contribution of Ph.D. student Veronika Michalková
to papers presented in the thesis
Paper A
Michalková V., Valigurová A., Dindo M.L. & Vaňhara J., in press: Larval morphology and
anatomy of the parasitoid Exorista larvarum (Diptera: Tachinidae), with an emphasis on
cephalopharyngeal skeleton and digestive tract. Journal of Parasitology.
Student co-authored the original idea of the paper; took part in histological processing the
samples, in photomicrograph documentation; wrote the first version of the manuscript and
prepared its final version.
Paper B
Valigurová A., Michalková V., Koník P., Dindo M.L., Gelnar M. & Vaňhara J., in prep.:
Penetration and encapsulation of larval endoparasitoid, Exorista larvarum (Diptera:
Tachinidae) in the factitious host Galleria mellonella (Lepidoptera: Pyralidae).
Co-authored the original idea of the paper; took part in histological processing the samples,
photomicrograph documentation and immunological experiments; participated in the first
version of the manuscript and prepared its final version.
Paper C
Mückstein P., Tschorsnig H.-P., Vaňhara J. & Michalková V., 2007: New host and country
records for European Tachinidae (Diptera). Entomologica Fennica 18:179-183.
Took part in collecting the samples; contributed to the final version of the manuscript.
Paper D
Vaňhara J., Tschorsnig H.-P., Herting B., Mückstein P. & Michalková V., 2009: Annotated
host catalogue for the Tachinidae (Diptera) of the Czech Republic. Entomologica Fennica 20:
22-48.
Took part in mining of published data; participated in the finalization of manuscript and
contributed to the final version of manuscript.
Prof. RNDr. Jaromír Vaňhara, CSc.
supervisor
Acknowledgement
I would like to express my gratitude to my supervisor Prof. Jaromír Vaňhara for his
comprehensive help by supervising my study and to Prof. Rudolf Rozkošný for his valuable
comments by solving the problems. I greatly appreciate the help, advice and fruitful
cooperation of Assoc. Prof. Maria Luisa Dindo (DiSTA, University of Bologna) who gave me
the chance to work in experimental field of entomology.
For opportunity to get an experience in non-dipteran research, I would like to thank
Assoc. Prof. Stano Pekár who taught me how to be a professional. I am indebted to Dr.
Andrea Tóthová for introducing me into molecular techniques and phylogenetic analyses.
Special thanks go to my colleagues and friends: remarkable co-author and co-worker Dr.
Andrea Valigurová, field and lunch-mate M.Sc. Stanislav Korenko, my special co-beings
M.Sc. Natália Muráriková, M.Sc. Hanka Novotná and Dr. Miroslava Barančeková and M.Sc.
Sérgio Silva Henriques (for the Portuguese sun in the office); for their help and to stand by
me always and mainly in the hard times. I will never forget self-sacrifice of Bc. Patrick S.
Corbett for correcting my English.
Many thanks go to M.Sc. Jana Benešová (Department of Experimental Biology,
Faculty of Science, Masaryk University) for the help with the rearing of Galleria mellonella.
I am greatly indebted to the members from the Laboratory of Electron Microscopy (Institute
of Parasitology, Biology Centre, ASCR, České Budějovice) and to the members from
Department of Histology and Embryology (Faculty of Medicine, Masaryk University) for
their help and technical assistance. Catalogue and host-parasitoid coevolution work would be
never possible without help and material kindly provided by Dr. Cezary Bystrowski, Dr.
Vladimír Straka, Prof. Zdeněk Laštůvka, Dr. Peter Tóth (Tachinidae), Dr. Petr Kment
(Heteroptera), Dr. Ladislav Roller (Hymenoptera), Václav Týr and Dr. Igor Malenovský
(Coleoptera).
I would like to express a thousand thanks to my family and friends for neverending
support, help and love. Last but not least, I would like to thank my model organisms which
sacrificed everything for the science, their life and soul.
Dissertation related research activities were financially supported by the Ph.D.
Research Fellowship of the Czech Science Foundation GAČR 524/05/H536 and Research
Project of the Czech Ministry of Education MSM 0021622416.
ABSTRACT
Tachinidae represent a very diverse and ecologically important family of dipterous
parasitoids, which are important natural enemies of insect pests in most terrestrial ecological
communities and thus useful in biological control. Exorista larvarum (L.) is a polyphagous
gregarious larval endoparasitoid of lepidopterans and some hymenopterans of the family
Tenthredinidae. Its parasitization strategy includes direct oviposition of unincubated
macrotype eggs on the host body. The endogenous development of E. larvarum has been
analyzed in the last larval instar of a factitious host, the wax moth Galleria mellonella
(Lepidoptera, Pyralidae), using histological techniques and scanning electron microscopy.
This study focuses on the parasitoid internal body structures and their changes during the
larval development. The first and second instar larvae are enveloped by a host-derived
haemocyte capsule and attached to the respiratory funnel via a prominent anal hook located
between two anal lobes and spines of the last abdominal segment. The third instar larva
abandons the respiratory funnel and migrates free in the body cavity of the already dead host.
Emphasis is given to the prominent cephalopharyngeal skeleton, highlighting the
morphological aspects of its sclerotized as well as non-sclerotized components, and to the
extensive digestive tract, which occupies the major part of the body and overcome the vast
changes during the development. Subsequently, the host-parasitoid interactions were
observed; penetration of the parasitoid and gradual immune responses of the host, an
encapsulation. The larvae induce the formation of the integumental respiratory funnel while
penetrating into the host body and thus from the outset of their development they remain in
direct contact with atmospheric oxygen and avoid suffocation. Endoparasitoid penetration
evokes the host cellular defense resulting into the formation of the hemocyte capsule
consisting of multi-cellular sheath. This enveloping capsule later undergoes the melanization,
which is mostly obvious towards the posterior part of endoparasitoid. The tachinid larva uses
the encapsulation response to build a respiratory funnel, primarily built-up from the modified
host integument and this way benefits on host immune responses.
Tachinidae are diversified in many morphological and biological types with remarkable
diversity of hosts and large host ranges. Despite the abundance, diversity, and importance of
tachinids, relatively little is known about their life-history traits and host-records are known
for less than half of the species. Based on the parasitized host rearings and searching for the
hosts' data in museums, new host records were obtained from the different parts of Europe.
Annotated tachinid-host catalogue of the Czech Republic was prepared and summarizes
already verified data for this area from the end of 19th century.
ABSTRAKT
Čeleď Tachinidae představuje velmi různorodou a ekologicky významnou skupinu
dvoukřídlých parazitoidů, kteří jsou přirození nepřátelé hmyzích škůdců v mnoha
terestrických ekosystémech a stávají se tak vhodnými bioagens i pro případnou biologickou
kontrolu. Exorista larvarum (L.) je polyfágní druh, shlukující se larvální endoparazitoid
housenek motýlů a housenic blanokřídlých z čeledi Tenthredinidae. Strategií parazitace je
přímá ovipozice neinkubovaných makrotypických vajíček na tělo hostitele. Endogenní vývoj
E. larvarum byl analyzován v posledním larválním instaru nepřirozeného hostitele, zavíječe
voskového Galleria mellonella (Lepidoptera, Pyralidae), pomocí histologických technik a
skenovací elektronové mikroskopie. Výzkum byl zaměřen na studii vnitřní struktury
parazitoida a změn během larválního vývoje. První a druhý instar larev je obklopen kapsulou
tvořenou pozměněnými hemocyty hostitele a přichycen k dýchací nálevce pomocí análního
háku umístněného mezi dvěma análními laloky a rovněž pomocí trnů na posledním
abdominálním segmentu. Larva třetího instaru opouští dýchací nálevku a volně migruje
v tělní dutině již mrtvého hostitele. Práce je zaměřena na mohutný cephalopharyngeální
skelet, zvýrazněny jsou morfologické aspekty jeho sklerotizovaných i nesklerotizovaných
částí. Dále byl studován extremně dlouhý zažívací trakt, který zabírá převážnou většinu tělní
dutiny parazitoida a podléhá výrazným změnám během vývoje. Rovněž byly sledovány
interakce hostitel-parazitoid; penetrace parazitoida a následná imunitní odezva hostitele,
enkapsulace. Během penetrace do těla hostitele larvy indukují formování integumentální
dýchací nálevky, a tak od počátku jejich vývoje zůstávají v přímém kontaktu
s atmosférickým kyslíkem a zabraňují tak sufokaci. Penetrace endoparazitoida vyvolává
buněčnou odezvu hostitele vedoucí ke vzniku hemocytové kapsuly tvořící vícevrstvý obal
kolem parazitoida. Kapsula později podléhá melanizaci, která je nejvíce zřetelná směrem
k posteriórní časti endoparazitoida. Larva kuklice využívá imunitní odpověď hostitele
k výstavbě dýchací nálevky prvotně tvořené z pozměněného integumentu hostitele a tak
využívá imunitní odezvu hostitele ke svému prospěchu.
Tachinidae jsou diverzifikovány do mnoha morfologických a biologických typů
s mimořádnou diverzifikací a rozsahem hostitelů. Navzdory velkému rozšíření a jejich
významu, je poměrně málo známo o jejich životních cyklech a hostitelích. Na základě
dochování parazitovaných hostitelů i vyhledáváním muzejního materiálu s daty o hostitelích,
byly získány nové záznamy o hostitelích různých míst Evropy. Pro Českou republiku byl
rovněž zpracován komentovaný katalog hostitelů, který shrnuje ověřené záznamy pro toto
území od konce 19. století.
Contents
1. Introduction 10
1.1. Tachinid biology 10
1.1.1. Parasitization strategies 10
1.1.2. Larval development and pupation 11
1.1.3. Host range 12
1.2. Host - parasitoid interactions 14
1.2.1. Insect immune system 14
1.2.2. Parasitoid strategies to circumvent the host immune system 14
2. Aims of the study 17
3. Material and Methods 18
3.1. Insect rearing and experimental parasitization 18
3.2. Histological procedure 18
3.3. Scanning electron microscopy 18
3.4. Immunological experiments 19
3.4.1. Mouse immunization 19
3.4.2. Injection of polybead microspheres 19
3.4.3. Indirect immunofluorescence 19
3.5. Faunistic records and revisionary work 20
4. Summary of results 21
5. Literature cited 25
6. Appendix: Papers presented in the thesis 30
10
1. Introduction
1.1. Tachinid biology
Tachinidae represent the largest, the most diverse and ecologically important dipterous
family of entomophagous parasitoids (Mellini 1990). They are one of the most diversified
families of Diptera, with approximately 10,000 described species worldwide and second only to
the parasitic Hymenoptera in diversity and ecological importance as insect parasitoids (Stireman
et al. 2006). Tachinids play a significant role in regulating herbivore populations and structuring
ecological communities, both natural and managed (Grenier 1988). The family is divided into
four subfamilies, Phasiinae, Dexiinae, Exoristinae and Tachininae.
As tachinids exhibit an impressive diversity of morphologies and biological types, they
exploit remarkable number of parasitization and development strategies and wide diversity of
hosts. Despite the abundance, diversity, and importance of tachinids, relatively little is known
about their life-history traits and host-records are known for less than half of the species.
1.1.1. Parasitization strategies
Parasitization strategies involve direct oviposition (eggs are laid externally or injected into
the host) and indirect oviposition (eggs are laid away from the host), whether the eggs are
incubated and contain fully developed larvae when deposited or not.
By direct oviposition, oviparous species produce macrotype eggs highly specialized for
being fixed on the host body, usually deposited on specific parts of the host exoskeleton.
Ovoviviparous species deposit membranous eggs and larvae hatch at the moment of contact with
the host. First instar larvae are able to live free for a while and move to penetrate elsewhere on
the host body. In viviparous species, larva develops in reproductive tract in thin chorion in a
special uterine pocket. Ovoviviparous and viviparous tachinids are fairly numerous and take
advantage to not face the risk of being discarded. All of these species in the nature attack mobile
hosts. Endozoic deposition is also known by a few groups with the piercing ovipositor (generally
ovoviviparous and viviparous forms). The females perforate the tegument or deposit eggs
through the host natural openings (Mellini 1990).
Taxa with indirect oviposition can be also subdivided in ovoviviparous species, in which
larvae hatch soon after eggs are laid in the places frequented by the host and either wait for
passing hosts or actively search for the host; and species depositing microtype eggs on the trophic
11
substrate of the host. These are ingested by hosts, hatch in the gut, and the emerging first instar
larvae burrow into the hemocoel.
The number of produced eggs is proportional to the likelihood of a first instar to infect a
host. Predominantly larval hosts are attacked, but a significant fraction of tachinid species attacks
adults. Interestingly, the taxonomic diversity of the family is approximately evenly distributed
across oviposition strategies (Stireman 2006).
1.1.2. Larval development and pupation
Tachinidae have three larval endophagous instars with morphological structure perfectly
suited to the parasitic way of life (Thompson 1960, Tschorsnig & Richter 1998, Wood 1987).
Newly-hatched larvae, in exception by microtype eggs, penetrate the host tegument with
the mechanical use of cephalopharyngeal skeleton aided by an enzymatic substance in the saliva.
The larvae exhibit a remarkable way of assuring their respiration in which they form a respiratory
funnel by attaching their posterior body end with a pair of spiracles either to the host tegument or
to the victim's tracheal system to provide an adequate supply of atmospheric oxygen. First instars
which remain free in the hemocoel usually attach to the tracheal system or migrate to the specific
hypertrophy organs.
During the first instar, larvae are plasmophagous and feed mostly on host hemolymph to
avoid damaging of vital organs. The larger second instar is steatophagous and begins to feed
more aggressively on non-vital tissues, mainly on fatty stores. The third, sarcophagous instar,
abandons the respiratory funnel and migrates free in the body cavity of the already dead host
(Mellini, 1990).
Larval development is usually completed in one to three weeks, except for species that
diapause in the host. Larvae develop either singly or gregariously and either pupate in the dead
host or outside the host remains. Quite a few tachinids overwinter in the pupal stage (Stireman
2006).
The survival strategies of tachinid larvae and their development in insect hosts have been
described in detail in several works (e.g. Belshaw 1994, Hafez 1953, Herting 1960, Ichiki &
Shima 2003, Mellini 1990, Stireman 2006) but only a few studies deal with their inner
morphology and anatomy (Gardenghi & Mellini 1990, 1995, Gardenghi et al. 1991).
12
1.1.3. Host range
Tachinids exploit a wide diversity of hosts belonging to many orders and families of
insects (Fig. 1). The most usual hosts are phytophagous insects, primarily Lepidoptera,
Coleoptera, Hymenoptera, Heteroptera, and Orthoptera. In addition, hosts from the other six
insect orders are attacked; Blattodea, Dermaptera, Diptera, Embioptera, Mantodea, and
Phasmida. Several genera of tachinids attack non-insect arthropods; centipedes, scorpions
(Williams et al. 1990) and also host record of spider is noted (Vincent 1985). The strictest
associations between tachinids and hosts can be find in Phasiinae related to heteropterous hosts,
Scarabaeidae as hosts of Dexiini, Lepidoptera as hosts for most Tachininae and Exoristinae, and
Orthoptera as hosts for Ormiini (Stireman et al. 2006). Many tachinid species are polyphagous,
and a number have been reared from dozens of hosts in multiple families (Belshaw 1994). The
most diverse clades of parasitoids attack primarily phytophagous insects, so the tri-trophic
interactions play an important role in the species diversification. Factors as habitat choice, host
searching strategy, oviposition strategy, methods how to deal with physiological defenses of the
host, and developmental synchrony, affect tachinid host range (Stireman 2006).
Since more than half of the tachinid species are without the host records, attention is
focused on solving this question and some partial national catalogues are published; Great Britain
- Ford et al. (2000), Germany - Tschorsnig et al. (1994a,b, 1999), Czech Republic - Vaňhara
(2009), Russia - Chicova (1987), Turkey - Kara et al. (2003), North America - Arnaud (1978).
13
Fig. 1: Host associations of Tachinidae subfamilies with tribes, according to Stireman (2006). Diversity of
subfamilies is given by box height. For comparison, the Czech tachinid fauna and its already known hosts
are red-marked ( ).
14
1.2. Host - parasitoid interactions
Endoparasitoids develop in the body cavities of their hosts and must to deal directly with
the host immune system for which specialized adaptations are required.
1.2.1. Insect immune system
The internal insect defensive systems consist of humoral and cellular responses; and
encapsulation is perhaps the most common cellular response to foreign organisms that are too
large to be phagocytosed by individual hemocytes (Salt 1970). During encapsulation, the foreign
material is surrounded by a multilayered envelope of hematocytes forming a capsule. Capsule
formation involves the initial recognition of foreignness by non-phagocytic granulocytes
inducing phagocytic cells, plasmatocytes, to adhere and to build-up the capsule (Ratner & Vinson
1983). This cooperation between one or more classes of hemocytes is likely mediated by
cytokines and adhesion molecules (Strand & Pech 1995). The structure of hemocyte capsule has
been considered to be consistent in all arthropods in which it has been described. Typical capsule
consists of 5-30 layers of hemocytes packed tightly together and flattened, occasionally with
desmosomes or septate junctions interconnecting the individual cells (Baerwald 1979). Very
often the inner cells of capsule formed around living organisms undergo melanization with
melanin deposited on the surface of foreign material.
Phenoloxidase, together with prophenoloxidase activating enzyme, is responsible for
initiating the biosyntesis of melanin and is associated with three physiologically important
biochemical processes in arthropods; i.e. sclerotization of insect cuticle (Sugumaran 1998),
encapsulation and melanization of foreign organisms (Söderhäll et al. 1990), and wound healing
(Lai-Fook 1966). The profenoloxidase-phenoloxidase system has been implicated in humoral
immunity (Söderhäll & Ajaxon 1982) and this cascade has been considered analogous to the
mammalian coagulation-fibrinolytic complement system (Seybold et al. 1975).
1.2.2. Parasitoid strategies to circumvent the host immune system
Parasitoid species have developed different strategies to circumvent the host immune
defense and these adaptations are varied including both passive (avoidence, evasion) and active
(destruction, suppresion, subversion) mechanisms (Vinson 1990). The encapsulation response to
a parasitoid may also be correlated to a number of biotic (e.g. development stage, host diet and
15
physiology, superparasitism, multiparasitism) and abiotic factors (e.g. temperature, humidity),
(Brodeur & Vet 1995, Salt 1970). Encapsulated parasitoids may die of suffocation, starvation or
physical prevention of development. Partially encapsulated parasitoids are able to survive and
may continue to develop normally (Van den Bosch 1964). Generally, parasites must to evade or
disrupt host immune system to survive (Rizki & Rizki 1990).
In Hymenoptera, encapsulation is typically prevented by the venom and associated
constituents injected during oviposition (Feener & Brown 1997). Many parasitoids from the
family Braconidae, Ichneumonidae and Cynipidae carry polyDNA viruses which avoid the
formation of the hemocyte capsule.
In Tachinidae, with the exception of a few groups with the piercing structures, female
flies must deposit eggs externally on or near the host, and the newly hatched larva must actively
penetrate into the host. This lack of an ovipositor inhibits the injection of paralytic poisons, used
by hymenopterans to immobilize the host immune system, and thus developing larvae must to
have other ways to overcome the host immune defense (Stireman 2006). They evolved two main
strategies of avoidance of the host immune system; larvae maintain contact with outside air by
attaching their posterior spiracles to the host tracheal system or to the respiratory funnel created
primarily from the host integument (Eggleton & Belshaw 1993) and turn the host immune system
to their own advantage by rebuilding this respiratory funnel from products of host immune
responses (Salt 1968). In some species of Tachinidae larvae move to a specific tissue and remain
hidden in these protected locations until they are ready to finish their development and consume
the host (Feener & Brown 1997). These strategies of how to circumvent host physiological
defenses with lack of host-specific adaptations could also explain the wide host ranges of many
tachinid species, which tend to be less host specific than hymenopteran parasitoids. Tachinids
adapted to the significantly narrower host ranges, usually remain free in the hemocoel without
forming a respiratory funnel (Belshaw 1994). In addition, tachinids may be relatively tolerant of
toxins actively or inadvertently ingested by their hosts and adapted to the toxic environments
(Gauld et al. 1992, Mallampalli et al. 1996).
As for immunological observations and host-parasitoid interactions, an overwhelming
majority of studies deals with parasitic hymenopterans, whereas there is a striking paucity of
knowledge regarding dipterous parasitoids. Despite of few studies describing the structure of the
16
hemocyte capsule and respiratory funnel in different tachinid species (Mellini & Cucchi 1965;
Baronio et al. 1974), there is still lack of sufficient data on tachinids.
17
2. Aims of the study
The general aims of this thesis were to investigate the host-parasitoid interactions based on:
1) biological adaptations of tachinid larvae
o to study the morphological aspects of larval developmental stages
o to study host immune defense reactions and parasitoid avoidance
2) ecological adaptations of tachinids
o to note life-history traits and obtain new host records
o to participate in preparing the annotated host catalogue for Tachinidae of the Czech
Republic
18
3. Material and methods
3.1. Insect rearing and experimental parasitization
Laboratory colonies of E. larvarum were established from puparia obtained from the
Laboratory of Entomology, DiSTA, University of Bologna, in 2006. The colonies were
maintained at 26°C, 60% RH and 16:8 hr (L:D) photoperiod, as described previously (Dindo et
al., 1999). Adult flies were held in Perspex cages (35 × 25 × 30 cm) with an air inlet and fed with
lump sugar and cotton balls soaked in 20% aqueous honey solution. For experimental
parasitization, last instar larvae of G. mellonella were exposed to fertile female tachinid flies kept
in the cage and generally removed after 3-4 parasitoid eggs had been laid on their body surface.
Three days after, the wax moth larvae were examined using a stereo binocular microscope to
confirm the successful penetration of the newly hatched tachinid larvae. Parasitized hosts were
collected and kept in plastic boxes until dissection, which was carried out on a daily basis
following parasitoid egg hatching. Their body parts with obvious parasitization were dissected
and fixed for subsequent tissue processing. In vivo observations of all three larval instars of the
parasitoid were documented by photomicrographs taken using a stereo binocular microscope
Olympus SZX12. Terminology of larval structures used in the present study follows those of
Chapman (1998) and Teskey (1981).
3.2. Histological procedure
Specimens were fixed in AFA (Alcohol-Formalin-Acetic Acid) fixative and processed
using standard histological methods. The material was embedded in Histoplast II and sectioned.
Sections 7 m thick were stained with hematoxylin-eosin, Masson's trichrome (blue and green)
or with Lillie's ferric-ferricyanide melanin staining method (Humason, 1967). Additional
specimens were fixed in fixatives containing paraformaldehyde (Karnovsky's fixative or Baker's
calcium formol) and processed for cryosectioning (frozen section procedure) in order to obtain
sections of better quality. Frozen sections were stained with azan or hematoxylin-eosin.
Histological sections were investigated using a light microscope Olympus BX51.
3.3. Scanning electron microscopy
Specimens were fixed overnight in freshly prepared 3% glutaraldehyde in cacodylate
buffer at 4°C, washed 3 times (5 min each) in cacodylate buffer, post-fixed in 1% osmium
19
tetroxide in cacodylate buffer for 2 hr at room temperature, and finally washed 3 times (5 min
each) in the same buffer. After dehydration in a graded series of acetone, the specimens were
critical point-dried using CO2, coated with gold, and examined using a JEOL JSM-7401F - Field
Emission Scanning Electron Microscope (FE SEM) or Vega\\XMH - Tescan Scanning Electron
Microscope (SEM).
3.4. Immunological experiments
3.4.1. Mouse immunization
Mice were immunized with salivary homogenate of E. larvarum, three times in two week
intervals. Every mouse was injected with 100 l of antigen and IFA solution (1:1). Two weeks
after the final injection, blood from the mice was collected and allowed to sediment overnight at
4°C. After the sedimentation, serum was asphyxiated and cleared by centrifugation at 1000g.
Antigen-binding capability of the sera was assessed by dot blot.
3.4.2. Injection of polybead microspheres
Polybead polystyrene 90 m microspheres (Polysciences) were used as encapsulation
targets. A total volume of 200 l of polybeads was coated with 3 l of antigens obtained from
tachinid salivary homogenate with 1% BSA and incubated during the night at 4°C, three times
centrifugated and washed in PBS, blocked in 1% BSA for 30 min, three times centrifugated and
washed in PBS and diluted in Ringer solution. The coated polybead microspheres in insect
physiological solution (1 l) were injected into the larvae of G. mellonella using the
micromanipulator. Control wax moth larvae were injected with polybead microspheres blocked
in 1% BSA alone without the antigen coat.
3.4.3. Indirect immunofluorescence
Deparaffined histological sections were incubated with anti-saliva immune serum diluted
in PBS (1:50) for 30 min at 37°C, three times washed with PBS for 5 min and incubated with
FITC-conjugated anti-mouse IgG (Sigma-Aldrich) diluted in PBS (1:300) for 30 min at 37°C.
Secondary antibody was flicked away and preparations were counterstained with Evans blue
(1:20000) for 3 min, two times washed with PBS for 5 min, mounted in anti-fade based on 2.5%
DABCO (Sigma-Aldrich) mixed with glycerol and 0.1 M PBS. Controls were performed with
20
FITC-conjugated anti-mouse IgG alone without primary antibody. Preparations were observed
and documented using an Olympus BX60 fluorescence microscope fitted with a WB filter cube,
having an excitation range of 450-480 nm.
3.5. Faunistic records and revisionary work
Host-parasitoid data were achieved from rearing of parasitized hosts collected in the field
and by searching for the host data in the private collections or museums. The tachinid species
were identified predominantly by the authors, hosts by the collectors.
The start of revisionary work concerning reared Tachinidae known from the area of
present-day Czech Republic was done by the late B. Herting, who studied the old tachinid
findings deposited in The Natural History Museum Vienna, Austria (Herting 1960) and collection
in the Forestry and Game Management Research Institute, Strnady near Prague. The large
collection of Wachtl from Vienna and Insbruck museums was completely revised by H.-P.
Tschorsnig and B. Herting (Tschorsnig & Herting 2005). Recent material sent by several Czech
entomologists (M. Barták, Z. Laštůvka, P. Mückstein etc.) was revised or identified by H.-P.
Tschorsnig and J. Vaňhara during the last fifteen years.
The arrangement of subfamilies, tribes and species of the tachinids mainly follows Herting
and Dely-Draskovits (1993), but updated nomenclature to the relevant catalogue is listed in
publication.
21
4. Summary of results
Presented thesis is composed of four partial studies, results of which were composed into
the original scientific papers. The publications are organized following the main topics and aims
of the thesis. First are listed the publications dealing with parasitoid morphology and hostparasitoid
interactions, followed by those related to the host-parasitoid records.
Paper A
Michalková V., Valigurová A., Dindo M.L. & Vaňhara J., 2009: Larval morphology and
anatomy of the parasitoid Exorista larvarum (Diptera: Tachinidae), with an emphasis on
cephalopharyngeal skeleton and digestive tract. Journal of Parasitology (accepted)
The endogenous development of the tachinid gregarious larval parasitoid Exorista
larvarum L. (Diptera: Tachinidae) has been analyzed in the last larval instar of a factitious host,
the wax moth Galleria mellonella L. (Lepidoptera: Pyralidae), with the use of histological
techniques and scanning electron microscopy. This study focuses on the parasitoid internal body
structures and their changes during the larval development. The first and second instars are
enveloped by a host-derived hemocyte capsule and attached to the respiratory funnel via a
prominent anal hook located between two anal lobes. The third instar abandons the respiratory
funnel and migrates free in the body cavity of the already dead host. Emphasis is given to the
prominent cephalopharyngeal skeleton, highlighting the morphological aspects of its sclerotized
as well as non-sclerotized components and to the replacement of the mouth hooks by a newer
pair, during which the new mouth hooks are formed above the old hooks. In addition to the
cephalopharyngeal skeleton, the anterior third of the larval parasitoid body is occupied by large
salivary glands, massive proventriculus, and cerebral ganglia. The extensive digestive tract,
which occupies the major part of the body, is differentiated into well-marked individual parts.
The three main regions of digestive tract are clearly distinguishable; stomodeum, mesenteron and
proctodeum. The stomodeum is differentiated into the pharynx, esophagus and proventriculus. As
the tachinid larvae exhibit a massive extraintestinal digestion enabled by prominent salivary
glands extending into the middle of the body cavity, the extremely thin esophagus together with
the prominent muscular proventriculus serve as a pump. The abdomen is predominantly filled
with the extremely long mesenteron that increases in size during the larval development,
22
outgrows the body size and reverses in the body cavity. The proctodeum is differentiated into an
ileum and rectum. The adipocytes possess a distinct plasma membrane and the adipose tissue
exhibit a high cellular density in later instars. The respiratory system is of amphipneustic type
and consists of two main longitudinal lateral trunks connected with transverse commissures, but
first instars possess a respiratory system of the metapneustic type. A well-developed musculature
firms the cephalopharyngeal skeleton and connects the body segments to provide appropriate
movement in the host tissue. The whole body is covered by an apparently thin integument, with
strong spines serving for the attachment to the host tissue and especially numerous in the anterior
and posterior body parts.
Paper B
Valigurová A., Michalková V., Koník P., Dindo M.L., Gelnar M. & Vaňhara J.: Penetration
and encapsulation of larval endoparasitoid, Exorista larvarum (Diptera: Tachinidae) in the
factitious host Galleria mellonella (Lepidoptera: Pyralidae). (in preparation)
The tachinid fly, Exorista larvarum is a polyphagous larval endoparasitoid with a direct
oviposition of un-incubated macrotype eggs on the host exoskeleton. The invasion process of the
larval parasitoid into host tissue and the formation of a hemocyte capsule, as a result of host
cellular response, were studied in the last larval instar of the wax moth, Galleria mellonella L.
(Lepidoptera: Pyralidae) in experimental conditions using advanced histological methods. The
newly-hatched tachinid larva penetrates the host integument, using the robust cephalopharyngeal
skeleton, leaving a dark spot at its entry point as a result of cuticle melanization. The parasitoid
profits from the host response leading to encapsulation, using it for building the primary
respiratory funnel and thus ensuring its direct contact with atmospheric oxygen. First instar larva
does not migrate within the host body cavity and develops through three larval instars being
attached to the respiratory funnel by an anal hook and numerous cuticular spines. The enveloping
hemocyte capsule consists of a multi-cellular sheath that later undergoes melanization, more
evident towards the abdominal part of the parasitoid. The third instar larva abandons the funnel,
migrates within already dead host to consume the remains and at the end of larval development
pupates outside the host carcass. Furthermore, additional immunohistochemical analysis and
23
simulated parasitization with polybead microspheres were used to study host-parasitoid
interactions.
Paper C
Mückstein P., Tschorsnig H.-P., Vaňhara J. & Michalková V., 2007: New host and country
records for European Tachinidae (Diptera). Entomologica Fennica 18:179-183.
The tachinid fauna of Europe is well known, but knowledge of tachinids biology,
including insect hosts is still rather poor. Many problems concerning the host specificity of this
large family have not been solved yet, so every host record is important. The paper presents host
records for 17 species of Tachinidae (of subfamilies Exoristinae and Tachininae) from the Czech
Republic, Slovakia, Austria, Croatia, Macedonia, Italy, Spain, Portugal, and Bulgaria. New
parasitoid-host couples are: Exorista larvarum - Melanchra pisi; Exorista segregata - Catocala
nymphaea; Sturmia bella - Hadena compta; Spallanzania multisetosa - Cycnia sordida (first host
record); Tachina praeceps - Cucullia bubaceki; Bithia modesta - Bembecia megillaeformis.
New country records of tachinid species: Rhacodinella apicata from the Czech Republic,
Masicera pavoniae from Macedonia and Bithia demotica from Portugal, are presented.
Paper D
Vaňhara J., Tschorsnig H.-P., Herting B., Mückstein P. & Michalková V., 2009: Annotated
host catalogue for the Tachinidae (Diptera) of the Czech Republic.
Entomologica Fennica 20: 22-48.
An annotated host catalogue for Tachinidae of the Czech Republic comprises 149 of 476
tachinid species which are currently known from this country. Totally 195 hosts are listed. The
bibliography for the host records consists of 116 papers of 55 researchers. The first host records
of Tachinidae date back to the second half of the 19th century. Several records of hitherto
unpublished material are included. Phryxe setifacies and Anthomyiopsis plagioderae are first
records for the Czech Republic. The majority of the Czech host records belongs to the subfamily
Exoristinae (103 tachinid species, 579 records of 332 host parasitoid couples represent 82 % of
all Czech host records) and a lesser extent to the subfamily Tachininae (33 tachinid species, 91
24
records of 54 host parasitoid couples represent 13 % of all Czech host records) and Dexiinae (13
tachinid species, 23 records of 34 host parasitoid couples represent 5 % of all Czech host
records). The average for the Palaearctic region is not much different (74 % of the host records
belong to Exoristinae, 13 % to Tachininae, 7 % to Dexiinae, and 6 % to Phasiinae). There are no
rearing records of Phasiinae for the Czech Republic because their hosts Hemiptera do not play a
significant role as pests in the area under the study. Also species of Orthoptera and Diptera are
usually not important as pests in the Czech Republic, which results in there not being a single
record for these host orders. Most hosts in the Czech Republic have the unspecialized
polyphagous parasitoids Exorista larvarum, Blondelia nigripes, Compsilura concinnata and
Phryxe vulgaris, but this is not different from what is already known from other European
countries. Investigations on tachinid-host relations in the Czech Republic were primarily focused
on pests in forestry, so it is not surprising that records for such typical hosts as Lymantria
monacha, Dendrolimus pini, Euproctis chrysorrhoea, Malacosoma neustria and Tortrix viridana
rank first. But also some common host species which are reared by practically every
lepidopterologist (e.g. Aglais urticae, Vanessa io, Saturnia pavonia) are presented by many
tachinid records. Parasitoids of unimpressive hosts or of those with hidden life strategies, or with
a nocturnal behavior are usually unknown and under-represented.
25
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27
Mallampalli N., Barbosa P. & Weinges K., 1996. Effects of condensed tannins and catalpol on
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parasitoids. Annual Review of Entomology 51: 525-555.
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28
Teskey H.J., 1981. Morphology and terminology - Larvae. In McAlpine J.F., Peterson B.V.,
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29
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30
6. Appendix: Papers presented in the thesis
Paper A
Michalková V., Valigurová A., Dindo M.L. & Vaňhara J., 2009: Larval morphology and
anatomy of the parasitoid Exorista larvarum (Diptera: Tachinidae), with an emphasis on
cephalopharyngeal skeleton and digestive tract. Journal of Parasitology (accepted).
Paper B
Valigurová A., Michalková V., Koník P., Dindo M.L., Gelnar M. & Vaňhara J.: Penetration and
encapsulation of larval endoparasitoid, Exorista larvarum (Diptera: Tachinidae) in the factitious
host Galleria mellonella (Lepidoptera: Pyralidae). (in preparation).
Paper C
Mückstein P., Tschorsnig H.-P., Vaňhara J. & Michalková V., 2007: New host and country
records for European Tachinidae (Diptera). Entomologica Fennica 18:179-183.
Paper D
Vaňhara J., Tschorsnig H.-P., Herting B., Mückstein P. & Michalková V., 2009: Annotated host
catalogue for the Tachinidae (Diptera) of the Czech Republic. Entomologica Fennica 20: 22-48.
Paper A
Larval morphology and anatomy of the parasitoid Exorista larvarum
(Diptera: Tachinidae),
with an emphasis on cephalopharyngeal skeleton and digestive tract.
MICHALKOVÁ V., VALIGUROVÁ A., DINDO M.L. & VAŇHARA J.
accepted in Journal of Parasitology (2009)
1
LARVAL MORPHOLOGY AND ANATOMY OF THE PARASITOID EXORISTA
LARVARUM (DIPTERA: TACHINIDAE), WITH AN EMPHASIS ON
CEPHALOPHARYNGEAL SKELETON AND DIGESTIVE TRACT
Veronika Michalková, Andrea Valigurová, Maria Luisa Dindo* and Jaromír Vaňhara
Department of Botany and Zoology, Faculty of Science, Masaryk University, Kotlářská 2, 611 37
Brno, Czech Republic. e-mail: vmichalkova@yahoo.com
*Dipartimento di Scienze e Tecnologie Agroambientali (DiSTA), Entomology Area, Universit degli
Studi di Bologna, Viale Fanin 42, 40127 Bologna, Italy
ABSTRACT: The endogenous development of the tachinid gregarious larval parasitoid
Exorista larvarum (L.) (Diptera: Tachinidae) has been analyzed in the last larval instar of a
factitious host, the wax moth Galleria mellonella (L.) (Lepidoptera: Pyralidae), with the use
of histological techniques and scanning electron microscopy. This study has focused on the
parasitoid internal body structures and their changes during the larval development. The first
and second instar are enveloped by a host-derived hemocyte capsule attached to the
respiratory funnel via a prominent anal hook located between 2 anal lobes. The third instar
abandons the respiratory funnel and migrates free in the body cavity of the already dead host.
Emphasis is given to the prominent cephalopharyngeal skeleton, highlighting the
morphological aspects of its sclerotized as well as non-sclerotized components. In addition to
the cephalopharyngeal skeleton, the anterior third of the larval parasitoid body is occupied by
large salivary glands, massive proventriculus, and cerebral ganglia. The extensive digestive
tract, which occupies the major part of the body, is differentiated into well-marked individual
parts. The abdomen is predominantly filled with the extremely long mesenteron that increases
in size during the larval development. The whole body is covered by an apparently thin
integument, with strong spines that are especially numerous in the anterior and posterior body
parts.
2
Tachinidae represent a very diverse and ecologically important family of dipterous
parasitoids, which are important natural enemies of insect pests in most terrestrial ecosystems
(Stireman et al., 2006) and thus useful in biological control (Grenier, 1988). Exorista
larvarum (L.) is a polyphagous gregarious larval endoparasitoid of lepidopterans and some
hymenopterans of the Tenthredinidae (Hafez, 1953a). Its parasitization strategy includes
direct oviposition of non-incubated macrotype eggs on the host body (Mellini, 1990). Females
deposit the eggs on the host body surface and the larvae hatch 3 days after oviposition under
laboratory conditions at 26 C (Dindo et al., 1999). Among most Tachinidae, the larvae exhibit
a remarkable way of assuring their respiration in which they form a respiratory funnel by
attaching their posterior body end, bearing a pair of spiracles, either to the host integument or
to the victim's tracheal system; a host hemocyte capsule developing around the parasitoid
larva contributes to the funnel formation (Mellini, 1990). The larvae of E. larvarum induce
the formation of the integumental funnel while penetrating into the host body (`primary
respiratory funnel' sensu Mellini, 1990) and thus from the outset of their endogenous
development they remain in direct contact with atmospheric oxygen. The larval development,
comprising of 3 endophagous instars, lasts 3 - 5 days at 26 C and pupation generally occurs
outside the host remains. The complete parasitoid development, from egg to adult, lasts about
15 days (Dindo et al., 1999). During the first 2 instars, all tachinid larvae, including those of
E. larvarum, feed mostly on host hemolymph and the fat body. The third instar larvae are
sarcophagous and rapidly destroy the victim's viscera (Mellini, 1990).
The survival strategies of tachinid larvae and their development in insect hosts have
been described in detail in several works (e.g. Herting, 1960; Mellini, 1990; Belshaw, 1994;
Ichiki and Shima, 2003). However, as for histological observations on the larval parasitoids,
an overwhelming majority of studies deals with parasitic hymenopterans, whereas there is a
striking paucity of knowledge regarding dipterous parasitoids. Only a few histological studies
illustrated also by diagrams concerning tachinids have been published and they deal either
with the morphology of their digestive tract (Gardenghi and Mellini, 1990; 1995; Gardenghi et
al., 1991) or reproductive system (Gardenghi and Mellini, 1980, 1992; Mellini et al., 1993b).
Furthermore, the structure of the hemocyte capsule and respiratory funnel in different tachinid
species has also been described (Mellini and Cucchi, 1965; Baronio et al., 1974).
The present study offers a complete histological analysis and detailed
photomicrograph documentation of the larval development of E. larvarum in its factitious
host, Galleria mellonella (L.) (Lepidoptera: Pyralidae). Contrary to previous works, the
present study describes the morphology of inner organs in all larval instars using advanced
3
histological techniques. The results presented herein focus mainly on the morphology of the
digestive tract and cephalopharyngeal skeleton, which, although described previously
(Ziegler, 1998), has not yet been investigated from a histological point of view. These
features are the most dominant in the larval development, overcome the extensive changes
during the development and their structure and function seem to be highly adapted for a
parasitic way of life.
MATERIALS AND METHODS
Rearings and experimental parasitization
Laboratory colonies of E. larvarum were maintained at 26 C, 60% RH and 16:8 hr
(L:D) photoperiod, as described previously (Dindo et al., 1999). The colonies were
established from puparia obtained from the Laboratory of Entomology, University of
Bologna, DiSTA, Italy in 2006. Adult flies were kept in Perspex cages (35 × 25 × 30 cm) with
an air inlet and fed with lump sugar and cotton balls soaked in 20% aqueous honey solution.
For experimental parasitization, last instar larvae of G. mellonella were exposed to fertile
female tachinid flies held in the cage and generally removed after 3-4 parasitoid eggs had
been laid on their body surface. Three days after, the wax moth larvae were examined using a
stereo binocular microscope to confirm the successful penetration of the newly hatched
tachinid larvae. Parasitized hosts were collected and kept in plastic boxes until dissection,
which was carried out on a daily basis following parasitoid egg hatching. Their body parts
with obvious parasitization were dissected and fixed for subsequent tissue processing. In vivo
observations of all three larval instars of the parasitoid were documented by
photomicrographs taken using a stereo binocular microscope Olympus SZX12.
Histological procedure
Specimens were fixed in AFA (Alcohol-Formalin-Acetic Acid) fixative and processed
using standard histological methods. The material was embedded in Histoplast II and
sectioned; sections 7 m thick were stained with hematoxylin-eosin, Masson's trichrome
(blue or green) or with Lillie's ferric-ferricyanide melanin staining method (Humason, 1967).
As the early developmental stages of E. larvarum within the host are difficult to cut, the
frozen section procedure (cryosectioning) and rapid microscopic analysis were used to obtain
sections of better quality. For this method, the material was fixed in fixatives containing
paraformaldehyde (Karnovsky's fixative or Baker's calcium formol). Frozen sections were
stained with azan or hematoxylin-eosin. Histological sections were investigated using a light
microscope Olympus BX51.
4
Scanning electron microscopy
The specimens were fixed overnight in freshly prepared 3% glutaraldehyde in
cacodylate buffer at 4 C, washed 3 times (5 min each) in cacodylate buffer, post-fixed in 1%
osmium tetroxide in cacodylate buffer for 2 hr at room temperature, and finally washed 3
times (5 min each) in the same buffer. After dehydration in a graded series of acetone, the
specimens were critical point-dried using CO2, coated with gold, and examined using a JEOL
JSM-7401F - Field Emission Scanning Electron Microscope (FE SEM) or Vega\\XMH Tescan
Scanning Electron Microscope (SEM).
Terminology of larval structures used in the present study follows those of Chapman
(1998) and Teskey (1981).
RESULTS
The female flies laid macrotype eggs on any part of the host body, including the head
and thoracic legs; however, most eggs were laid dorsally or latero-dorsally on the abdomen,
often on the membranous areas interconnecting the sclerites. The eggs usually hatched 3 to 4
days after oviposition and the newly hatched larvae actively penetrated the host integument
using the strong, well-developed cephalopharyngeal skeleton and musculature. Larvae pulled
the host tissue in while penetrating the host integument in order to build the primary
respiratory funnel. Parasitized hosts showed no obvious damage or abnormalities in this
phase; only a dark brown to black spot visible on the surface of their integument indicated the
area of tachinid larva penetration. Parasitization with multiple tachinid larvae of various body
size and stage of development, usually in number of 2-4 per host, was often observed. The
first instar larvae, characterized by a single mouth hook (Figs. 1, 2, 5, 6, 12), were easy to
recognize. In contrast, the 2 later instars possessing 2 mouth hooks (Figs. 3, 4, 7, 14) were
often difficult to distinguish from each other, especially during the late second and early third
instars. Thus, the longitudinal histological sections showing a majority of their internal
organs, especially the extremely long digestive tract with associated organs, turned out to be
very helpful for this purpose.
First instars
Histological sections confirmed that the first instar larvae do not migrate within the
host body cavity, but hang in the primary respiratory funnel (Fig. 20) attached to its wall by
means of the spines covering the last abdominal segment (Fig. 44). No anal hook was
observed in histological sections or specimens prepared for scanning electron microscopy.
5
Histology as well as examinations using a stereo binocular microscope during the dissection
of parasitized caterpillars constantly confirmed that the first instar of E. larvarum remained at
the point of entry with their posterior body part, bearing 2 spiracles, enclosed in the
respiratory funnel and thus being in direct contact with atmospheric oxygen. The invading
parasitoid induced formation of a hemocyte capsule as a result of the host cellular defense
(Fig. 20). The pseudocephalic segment bore a pair of short conical antennae and maxillary
sensory papillae (Fig. 5). The prominent cephalopharyngeal skeleton, protecting the anterior
part of digestive tract, terminated with a single curved, median mouth hook and possessed the
sharp short, already melanized cornua (Fig. 12). The mouth hook with an apparently sawtoothed
dorsal edge stuck out of the large mouth opening (Figs. 5, 6). The digestive tract
together with salivary glands and Malpighian tubules (Fig. 37) were already well-developed,
but not as prominent as in later instars, whereas the distinct cerebral ganglia already occupied
the major part of the anterior third of parasitoid body (Fig. 20). In our preparations, only the
posterior pair of spiracles was visible (Fig. 39).
Second instars
At this instar, the observed larvae were attached to the respiratory funnel (Figs. 3, 19,
21-24) via an anal hook located between 2 anal lobes (Fig. 10) and by sturdy spines of the
posterior segment (Fig. 11). Their integument was covered by narrow, sharp spines that were
especially numerous in the anterior body region (Fig. 7). The larvae had molted to the second
instar as shown by their morphology described below. The respiratory system was of
amphipneustic type and consisted of 2 main longitudinal lateral trunks, the same as in third
instar shown in Fig. 15, and connected with transverse commissures (Fig. 42). The
inconspicuous anterior spiracles were located latero-dorsally on the first thoracic segment
(Figs. 30, 41). Each spiracle was equipped with 4 papillae with circular openings (not shown).
The posterior spiracles (Figs. 19, 21) were located dorsally on the last abdominal segment and
each possessed a circular peritreme and 2 elongated spiracular slits strengthened by the
dentate chitinous processes (Fig. 40). The dark pigmented cephalopharyngeal skeleton
possessed 2 sickle-shaped mouth hooks, which were formed by mandibular sclerites and
extended from the oral cavity during locomotion and feeding (Figs. 3, 4, 7, 8). The bases of
the mandibular sclerites articulated with the hypopharyngeal sclerite (Fig. 25, 33). Its
posterior margins were in contact with the fused portion of the 2 large tentopharyngeal
sclerites , which were the most conspicuous parts of the cephalopharyngeal skeleton (Fig. 13).
Tentopharyngeal sclerite was composed of a pair of sclerites with 2 arms, dorsal and ventral
6
cornua, on either side of pharynx (Figs. 25, 26, 28-30). Both ventral cornua were joined,
supporting the pharynx, and the dorsal cornua were bridged anteriorly. At this instar, the
cornua were already intensively melanized (Fig. 13) and continued into non-sclerotized
connective tissue serving as the attachment point for musculature (Fig. 30). Both mouth hooks
(Fig. 27) and body spines were hollow inside (Fig. 31).
As the tachinid larvae exhibited a massive extraintestinal digestion enabled by
prominent salivary glands extending into the middle of the body cavity (Figs. 35), the
extremely thin esophagus together with the prominent muscular proventriculus served as a
pump. The stomodeum was lined by slightly flattened epithelial cells and differentiated into
the pharynx, esophagus and proventriculus (Figs. 30-36). The pharynx together with
pharyngeal dilator muscles was enclosed by pharyngeal sclerites of the cephalopharyngeal
skeleton, to which a well-developed musculature was deeply anchored (Figs. 25-31). The
conspicuous mesenteron outgrew the body size, reversed and occupied most of the body
cavity (Figs. 21-24). The mesenteron was lined by a monolayer of columnar cells bearing a
brush border on their luminal surface and an acellular peritrophic envelope formed a delicate
lining layer separating the luminal contents from the brush border of the mesenteric
epithelium (Fig. 38). Occasionally, host cells still not disintegrated by digestion could be seen
in the mesenteric part of the digestive tract (Figs. 36-38). The proctodeum was differentiated
into an ileum with apparently flattened cells and rectum (Fig. 36). The Malpighian tubules
arose from the ileum just behind the mesenteron (Fig. 23). The adipocytes possessing a
distinct plasma membrane increased in number (Fig. 42). A well-developed musculature
firmed the cephalopharyngeal skeleton and connected the body segments to provide
appropriate movement in the host tissue. Although the larvae were equipped with strong
tendons and apparently compact musculature (Fig. 43), the integument was very thin and
more extensive epidermal sheet covered only the areas nearby spiracles and mouth hooks
(Figs. 33, 40, 41). We did not observe a ventral nerve cord; only the extensive cerebral
ganglia could be seen in our observations (Figs. 17, 22-24, 35, 36).
At this stage of parasitization, the hosts ceased their movement. Apparent reduction
and disintegration of host tissue, especially the fat body, surrounding the encapsulated,
rapidly developing parasitoid could be seen. This disintegration was clearly evident especially
nearby the parasitoid anterior part with mouth hooks stuck into the hemocyte capsule. In
addition, the host internal organs were obviously compressed by parasitoid increasing in size
and occupying ever-greater space in host body.
7
Third instars
In the second instar larvae just before molting, we occasionally observed the
replacement of the mouth hooks by a newer pair, during which the new mouth hooks formed
above the old hooks (Fig. 14). After molting to the third instar, the chubby tachinid larva (Fig.
9) abandoned the respiratory funnel and migrated free within the body cavity of the already
killed host. The host tissues were completely destroyed; the internal organs were fully
disintegrated and exhibited a liquid appearance. As the internal organization of the second and
third instar larvae was very similar, only the photomicrographs documenting the internal
morphology of the second instar larva are shown here. The digestive tract of the tachinid larva
became even more extensive with several loops and extremely massive cephalopharyngeal
skeleton (Figs. 15, 16). Similarly to the second instar, the 3 main regions of digestive tract
were clearly distinguishable, i.e. stomodeum, mesenteron and proctodeum (Figs. 15-18). The
Malpighian tubules enlarged during later development. The adipose tissue exhibited a high
cellular density. The respiratory system was highly developed with numerous transverse
commissures. Cerebral ganglia were extensive as in previous instars, but still without a
definitive ventral nerve cord. At the end of larval development, all the internal contents of the
host were consumed and the third instar tachinid larvae abandoned the host carcass to pupate
outside the host remains.
DISCUSSION
As the insects are generally equipped with numerous hardened chitinous structures,
histological processing of insect tissue is usually very problematic. Possibly, also for this
reason, only a few detailed histological analyses can be found throughout the literature.
Although some of the previous works referred to the structure of tachinid cephalopharyngeal
skeleton (Ziegler, 1998) and illustrated diagrams were provided also to E. larvarum (Hafez,
1953b), recent possibilities of histological sectioning allowed us to document also the nonsclerotized
cavities of the mouth hooks and spines as well as the cornua continuing into the
non-sclerotized connective tissue with their attachment of musculature. Similar to the other
tachinids, the first instar of E. larvarum possess a simple cephalopharyngeal skeleton with
only a single mouth hook (Hafez, 1953b; Pape, 1992; Thompson, 1960). This unpaired
median hook is probably a fusion of 2 original hooks in the apical end of cephalopharyngeal
skeleton. Interestingly, we never found any signs of its rebuilding into a more complicated
cephalopharyngeal skeleton bearing 2 mouth hooks in the next 2 larval instars, but we may
suppose that the new mouth hooks are formed the same way as observed in the second instar.
8
As the first instar larvae possessed a flattened mouth hook with a saw-toothed upper edge, the
second and third instar larvae were already equipped with conical hooks with seemingly
multilayered surface. Individual layers were interlaced as shown in Figure 8. It is a question,
however, if this hook architecture is related to the development and/or replacement of hooks,
or is it only the result of the tissue processing for SEM?
Gardenghi and Mellini (1995) found a great quantity of host cells in the mesenteron of
E. larvarum larvae and noted that their extraintestinal digestion does not disintegrate the host
cells, but only the host tissues. However, we seldom noticed some particulate material,
generally only a few hemocytes, in the parasitoid mesenteron. The pharynx and
proventriculus, which possess well-developed musculature, seemingly allow the larvae to
rapidly engorge and homogenize the ingested tissues. The proventriculus, structurally the
most highly specialized part of the digestive tract serving as a pump for pre-digested liquid
food, did not bear any sclerotized pads or plates, but it was equipped with raised epithelium
and strong dense musculature. The extraordinarily long mesenteron, together with a unique
mechanism of tachinid larvae to block the defecation while feeding, is considered to be a form
of adaptation for a parasitic way of life (Gardenghi and Mellini, 1990; Gardenghi et al.,
1991). The digestive tract of E. larvarum larvae, however, lacks the retention structures
observed in other tachinids (Gardenghi and Mellini, 1995). Therefore, they defecate
throughout their development as observed by rearing them on artificial media (Dindo et al.,
1999); this fecal material is deposited and encrusted in the luminal side of wall of the primary
integumental respiratory funnel (Gardenghi and Mellini, 1995).
Although Hafez (1953b) provided detailed description of the external morphology of
E. larvarum larvae, he did not mention their attachment to the respiratory funnel via the anal
hook located between 2 anal lobes near the anal opening. Similarly, the tachinid Compsilura
concinnata (Meigen), laying the eggs directly into the host, uses 3 anal hooks surrounding the
posterior spiracles, 2 on the dorsal and 1 on the ventral side, to fix its posterior abdominal
segment to the wall of the host's organs (Ichiki and Shima, 2003). These structures are
considered to be a respiratory adaptation also suitable for holding the parasitoid body in a
fixed position within the host body. In contrast, the larva of E. larvarum is equipped only with
a single anal hook, but the attachment to the host tissue is supported also by sturdy spines
covering the posterior segment; thus, they contribute to the positioning of the parasitoid body.
As for the anal hook, the area of the anal lobes seemingly lacked the hook in first instar larvae
when observed using the scanning electron microscope. It is possible that the anal hook, if
present in the first instar larvae, is more fragile than in later instars and thus the putative lack
9
of this structure may also be due to mechanical damage caused by processing for scanning
electron microscopy. Unfortunately, we can not confirm its presence or absence as we did not
obtain complete histological sections showing this area.
Hafez (1953b) recorded only 2 commissures connecting the main tracheal trunks
leading to the posterior spiracles; however, we could see many of them in the histological
sections. It is known that first instar tachinid larvae inducing the formation of primary
respiratory funnel (including those of E. larvarum) possess a respiratory system of the
metapneustic type. As expected, in our preparations, only the posterior pair of spiracles was
visible.
The integument of the examined tachinid larvae was surprisingly very thin and we
could not detect the individual cuticle layers by light microscopy. We speculate that the
tachinid larvae take advantage of the hemocyte capsule enveloping their body within the host
tissues to protect themselves from the host internal environment and defense mechanisms, so
they do not need a protective thick cuticle. Extensive epidermis was seen only surrounding
the areas of spiracles and mouth hooks. It apparently serves as a mechanical support for these
openings. In comparison to their host, E. larvarum larvae were equipped with more prominent
and stronger tendons, and seemingly more compact musculature facilitating easier movement
in such constricted conditions.
In accordance with Mellini et al. (1993a), in the absence of suitable hosts or if unable
to reach them, the females oviposited even on inanimate objects or our hands, thus ensuring
the death of their offspring. The oviposition strategy was the same as observed by Hafez
(1953c), but it should be noted that females preferred oviposition on the surface of soft
membranous connections between body segments; in this way, the eggs were hidden between
the folds of cuticle and the penetration of the larvae into the host should be much easier. As
the parasitoid development was asynchronous, the larvae hatched and penetrated the host at
different times after oviposition and we often found the tachinid larvae of diverse
developmental stages within a single host. Even when we allowed females to deposit more
eggs (5-11), we could still find only 1 to 4 parasitoid larvae per host, with different body
sizes, depending on parasitoid number and intraspecific competition similar to the previous
findings (Mellini and Campadelli, 1996).
ACKNOWLEDGMENTS
The authors are gratefully indebted to Jana Benešová (Department of Experimental
Biology, Masaryk University) for the help with the rearing of Galleria mellonella and
10
Romana Šebestová (Department of Histology and Embryology, Masaryk University) for the
help with cryosectioning. We also wish to thank Elisa Marchetti and Laura Depalo (DiSTA,
University of Bologna) for their help with the rearing in Italy. This study was financially
supported by the research project MSM 0021622416, doctoral grant GAČR 524/05/H536 and
by the Italian Ministry of University and Research PRIN 2005.
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guaina attorno alle larve dei Ditteri Larvevoridi. Bollettino dell'Istituto di Entomologia della
Universit di Bologna 32: 91-103.
BELSHAW, R. 1994. Life history characteristics of Tachinidae (Diptera) and their effect on
polyphagy. In Parasitoid community ecology, B. A. Hawkins, and W. Sheehan, (eds.). Oxford
University Press, Oxford, U.K., p. 145-162.
CHAPMAN, R. F. 1998. The insects. Structure and function, 4th ed. Cambridge University
Press, Cambridge, U.K., 770 p.
DINDO, M. L., R. FARNETI, M. SCAPOLATEMPO, AND G. GARDENGHI. 1999. In vitro rearing of
the parasitoid Exorista larvarum (L.) (Diptera : Tachinidae) on meat homogenate-based diets.
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GARDENGHI, G., AND E. MELLINI. 1980. Sulla formazione dell'uovo microtipico e del relativo
apparato di fissazione in Gonia cinerascens Rond. (Diptera Larvaevoridae). Bollettino
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------, G. CAMPADELLI, AND E. MELLINI. 1991. Osservazioni anatomo-istologiche
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11
GRENIER, S. 1988. Applied biological control with tachinid flies (Diptera, Tachinidae): A
review. Anzeiger für Schädlingskunde, Pflanzenschutz, Umweltschutz 61: 49-56.
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------. 1953b. Studies on Tachina larvarum L. (Diptera, Tachinidae) II. Morphology of the
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------. 1953c. Studies on Tachina larvarum L. (Diptera, Tachinidae) III. Biology and lifehistory.
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Tachinidae). Annals of the Entomological Society of America 96: 161-167.
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Entomologia "Guido Grandi" dell'Universit di Bologna 45: 1-38.
------, AND G. CAMPADELLI. 1996. Analisi del superparassitoidismo di Exorista larvarum
(L.) nell'ospite di sostituzione Galleria mellonella L. Bollettino dell'Istituto di Entomologia
"Guido Grandi" University of Bologna 51: 1-11.
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Steiniella callida Meig. (Dipt. Larvaevoridae) nelle larve di Melasoma populi L. (Col.
Chrysomelidae). Bollettino dell'Istituto di Entomologia della Universit di Bologna 27: 215-
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------, G. CAMPADELLI, AND M. L. DINDO. 1993a. Artificial culture of the parasitoid
Exorista larvarum L. (Dipt. Tachinidae) on bovine serum-based diets. Bollettino dell'Istituto
di Entomologia "Guido Grandi" dell'Universit di Bologna 47: 223-231.
------, G. GARDENGHI, AND A. K. COULIBALY. 1993b. Caratteristiche anatomiche ed
istologiche dell'apparato genitale femminile di Exorista larvarum L., parassitoide deponente
uova macrotipiche sull'ospite. Bollettino dell'Istituto di Entomologia "Guido Grandi"
dell'Universit di Bologna 48: 45-58.
PAPE, T. 1992. Phylogeny of the Tachinidae family-group. Tijdschrift voor Entomologie 135:
43-86.
12
STIREMAN, J. O. III, J. E. O'HARA, AND D. M. WOOD. 2006. Behavior, ecology and evolution
of tachinid parasitoids. Annual Review of Entomology 51: 525-555.
TESKEY, H. J. 1981. Morphology and terminology ­ Larvae. In Manual of Nearctic Diptera,
Vol. 1, J. F. McAlpine, B. V. Peterson, G. E. Shewell, H. J. Teskey, J. R. Vockeroth and D.M.
Wood, (eds.). Biosystematics Research Institute, Monograph No. 27, Ottawa, Canada, p. 65-
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(Diptera). Transactions of the American Entomological Society 86: 207-224.
ZIEGLER, J. 1998. Die Morphologie der Puparien und der larvalen Cephalopharyngealskelette
der Raupenfliegen (Diptera, Tachinidae) und ihre phylogenetische Bewertung. Studia
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13
FIGURES 1-11. Larvae of Exorista larvarum and their superficial structures.
(1) The first instar displaying the projecting single mouth hook (arrow); in vivo observation
(dorsal view). Scale bar = 100 m.
(2) The first instar, displaying the single mouth hook (arrow), sitting in the respiratory funnel;
in vivo observation. Scale bar = 100 m.
(3) The second instar larva sitting in the respiratory funnel (rf), surrounded by remains of
hemocyte capsule (ca); in vivo observation. Arrow indicates the cephalopharyngeal skeleton
with 2 mouth hooks. Scale bar = 300 m.
(4) The second instar showing the cephalopharyngeal skeleton in dorsal (upper) and lateral
view (lower); in vivo observation. Scale bar = 150 m.
(5) Pseudocephalic segment of the first instar showing the antennae (arrowheads), maxillary
papillae (circled) and single mouth hook (arrow); SEM. Scale bar = 10 m.
(6) Detail of the single mouth hook of the first instar (arrow); SEM. Scale bar = 10 m.
(7) Pseudocephalic segment of the second instar showing 2 prominent mouth hooks (arrows)
and numerous spines (arrowhead); FE SEM. Scale bar = 20 m.
(8) Mouth hook of the second instar larva; SEM. Scale bar = 50 m.
(9) The third instar with stout body and prominent posterior spiracles (asterisk) in dorsal
portion of terminal end; in vivo observation. Scale bar = 500 m.
(10) Anal hook (arrowhead) between 2 anal lobes (al) in the second larval instar; FE SEM.
Scale bar = 10 m.
(11) Cuticular spines (head-bound) surrounding the posterior spiracles in the second instar
larva; FE SEM. Scale bar = 10 m.
14
15
FIGURES 12-18. Cephalopharyngeal skeleton of Exorista larvarum larvae in histological
sections stained for melanin (dark green) and in vivo observations.
(12) Cephalopharyngeal skeleton of the first instar larva with 2 cornua (arrows) and a single
mouth hook (arrowhead) stuck in the capsule (ca), cuticular spines (circled). Scale bar = 20
m.
(13) Cephalopharyngeal skeleton of the second instar larva with 2 mouth hooks (arrowheads)
and 2 tentopharyngeal sclerites (arrows). Scale bar = 20 m.
(14) Replacement of the mouth hooks of the second instar (arrowheads) by newer pair
(arrows) before molting to the third instar. Scale bar = 250 m.
(15) Dissection of the third instar showing the longitudinal tracheal trunks (t), proventriculus
(p), mesenteron (me), Malpighian tubules (mt) and salivary glands (s). Scale bar = 1 mm.
(16) Digestive tract of the third instar larva: proventriculus (p), mesenteron (me), Malpighian
tubules (mt), ileum (il), rectum (r), anus (a). Scale bar = 1 mm.
(17) In toto observation of cerebral ganglia (asterisk), esophagus (e), proventriculus (p) and
beginning of mesenteron (me); third instar larva. Scale bar = 500 m.
(18) Detail of the rectum (r) with anus (a) and Malpighian tubules (mt) in third instar. Scale
bar = 1 mm.
16
17
FIGURES 19-24. Histological sections of larvae of tachinid Exorista larvarum encapsulated
within the host Galleria mellonella.
(19) Cross section of the host body with 2 tachinid larvae in second larval instar (ls longitudinal
section and os - oblique section); hemocyte capsule (ca), tachinid cerebral
ganglia (asterisk), host air sac (has), host mesenteron (hme), host silk gland (hsg), host
tracheal system (hts), respiratory funnel (rf), tachinid spiracle (arrow). Scale bar = 1 mm.
(20) Longitudinal section of the first instar tachinid larva showing the prominent cerebral
ganglia (asterisk), salivary glands (s) in the anterior part of the body and tracheal system
(arrows) visible especially in the posterior part. Note the respiratory funnel (rf) that continues
into the capsule (ca) enveloping the parasitoid. Scale bar = 100 m.
(21) Longitudinal section of the second instar larva surrounded by hemocyte capsule (ca);
salivary glands (s), esophagus (e), folded mesenteron (me), Malpighian tubules (mt), spiracles
(sp) leading into the respiratory funnel (rf) and thin integument of tachinid larva
(arrowheads). Scale bar = 200 m.
(22) Oblique (os) and longitudinal section (ls) of 2 second instar larvae; hemocyte capsule
(ca) with respiratory funnel (rf), cerebral ganglia (asterisks), salivary gland (s), esophagus (e),
mesenteron (me), ileum (il). Scale bar = 250 m.
(23) Longitudinal section of the second instar larva showing the capsule (ca), part of the
cephalopharyngeal skeleton (arrowhead), cerebral ganglia (asterisk), ileum (il), Malpighian
tubules (mt), mesenteron (me), esophagus (e), proventriculus (p) and salivary gland (s). Scale
bar = 250 m.
(24) Longitudinal section of the second instar larva; hemocyte capsule (ca) with respiratory
funnel (rf), cerebral ganglia (asterisk), proventriculus (p) connected with the extremely
narrow esophagus, mesenteron (me), trachea (arrows). Scale bar = 250 m.
18
19
FIGURES 25-31. Histological sections of the second instar of Exorista larvarum showing the
cephalopharyngeal skeleton.
(25) Cephalopharyngeal skeleton's ventral cornua (arrows) articulated via hypopharyngeal
sclerites (asterisks) with 2 mouth hooks (arrowheads). Note the prominent salivary glands (s)
lining both sides of the cephalopharyngeal skeleton and muscles (mu) connected to the
integument. Scale bar = 50 m.
(26) Longitudinal section of the cephalopharyngeal skeleton (arrows) protecting the pharynx
(ph) and the muscles (mu) attached to its cornua. Scale bar = 100 m.
(27) Cross section of the mouth hook with central cavity (asterisk). Scale bar = 10 m.
(28) Oblique section of the pseudocephalic segment with prominent salivary glands (s) and 2
mouth hooks (arrowheads); adipocytes (a), pharynx (ph), muscles (mu) attached to the
integument. Scale bar = 100 m.
(29) Pharynx (ph) protected by the cephalopharyngeal skeleton (arrows) and salivary glands
(s). Scale bar = 100 m.
(30) Oblique section of the pseudocephalic segment; anterior spiracles (sp), salivary glands
(s), pharynx (ph) protected by cephalopharyngeal skeleton's dorsal cornua (arrows). Note the
non-sclerotized tissue of the cornua (arrowheads) connected with muscles (mu). Scale bar =
100 m.
(31) Mouth opening (m) lined with epithelium (ep); pharynx (ph); cuticular spines (circled).
Scale bar = 25 m.
20
21
FIGURES 32-38. Digestive tract of larval Exorista larvarum in histological sections.
(32) Dorsoventral longitudinal section of the second instar larva's cephalopharyngeal skeleton
(arrows) with pharyngeal dilator muscles (mu), cerebral ganglium (asterisk) and salivary
gland (s). Scale bar = 100 m.
(33) Cross section of the mouth hooks (arrowheads) and cephalopharyngeal skeleton in
hypopharyngeal sclerite (arrows) of the second instar; pharyngeal muscles (pm), muscle (mu)
attached to the epithelium (ep) lining the hook that probably facilitates its movement. Scale
bar = 50 m.
(34) Detailed view of the mouth opening lined with epithelium (ep), suggesting the
extraintestinal digestion (arrow), second instar larva. Scale bar = 25 m.
(35) Detail of esophagus (e) located between cerebral ganglia (asterisk), proventriculus (p)
and salivary glands (s), second instar. Scale bar = 100 m.
(36) Longitudinal section of the second instar larva showing the cerebral ganglia (asterisk),
salivary glands (s), proventriculus (p), ileum (il), rectum (r), Malpighian tubules (mt) and
trachea (t). Scale bar = 100 m.
(37) Cross section of the first instar larva showing reversed mesenteron (me), Malpighian
tubules (mt) and thin integument (arrowheads). Scale bar = 20 m.
(38) The luminal surface of mesenteron of second instar covered by microvilli (circled);
perithrophic membrane (pm) and ingested but still compact host hemocytes (arrow). Scale bar
= 20 m.
22
23
FIGURES 39-44. Respiratory system and integumental structures of Exorista larvarum larvae.
(39) Posterior spiracles of the first instar larva; epidermis (ep), closed peritreme (pe), thin
larval integument (arrowhead). Scale bar = 20 m.
(40) Posterior spiracles of the second instar larva; peritreme (arrowheads), spiracular slits
(circled) with dentate chitinous processes, apertures of spiracles enclosed with epidermis (ep)
and attached trachea (t). Scale bar = 100 m.
(41) Anterior spiracle of second instar larva with papillae (asterisk) underlain by epidermis
(ep); integument consisting of thin cuticle (cu) and epidermis (ep). Scale bar = 25 m.
(42) Longitudinal section of trachea with clearly visible taenidium (arrowheads) and covered
with epithelium (ep); adipocytes (a), second instar larva. Scale bar = 20 m.
(43) Detail of muscles (mu) connected to the larval integument via tendon (circled); epidermis
(ep), cuticle (cu) covered with spines (arrowhead), second instar larva. Scale bar = 20 m.
(44) Detail showing the attachment of tachinid larva to the respiratory funnel (rf) via cuticular
spines (circled), first instar larva. Scale bar = 20 m.
24
Paper B
Penetration and encapsulation of larval endoparasitoid, Exorista larvarum (Diptera:
Tachinidae) in the factitious host Galleria mellonella (Lepidoptera: Pyralidae).
VALIGUROVÁ A., MICHALKOVÁ V., KONÍK P., DINDO M.L., GELNAR M. & VAŇHARA J.
manuscript in preparation*
Penetration and encapsulation of larval endoparasitoid, Exorista larvarum (Diptera:
Tachinidae) in the factitious host Galleria mellonella (Lepidoptera: Pyralidae)
Andrea Valigurováa1
, Veronika Michalkováa1
, Peter Koníkb,c
, Maria Luisa Dindod
,
Milan Gelnara
and Jaromír Vaňharaa
a
Department of Botany and Zoology, Faculty of Science, Masaryk University, Kotlářská 2,
611 37 Brno, Czech Republic
b
Faculty of Science, University of South Bohemia, Branišovská 31, 370 05 České Budějovice,
Czech Republic
c
Biology Centre of the Academy of Sciences of the Czech Republic, Institute of Parasitology,
Branišovská 31, 370 05 České Budějovice, Czech Republic
d
Dipartimento di Scienze e Tecnologie Agroambientali, Entomology Area, Universit degli
Studi di Bologna, Viale Fanin 42, 40127 Bologna, Italy
1
These authors contributed equally to the work
*As the authors feel the need to complete and confirm these already obtained results with
additional advanced methods, data described herein are processed and presented as a
manuscript in preparation.
1
Abstract
The tachinid fly, E. larvarum belongs to the polyphagous larval endoparasitoids depositing
their eggs on the host exoskeleton. The attachment of larval E. larvarum and formation of
capsule were studied in the last larval instar of the wax moth, Galleria mellonella L.
(Lepidoptera: Pyralidae). The tachinid larvae burrow through the host integument when they
hatch, using the robust cephalopharyngeal skeleton, leaving a dark spot at the point of their
penetration as a result of host cuticle melanization. Using mouthhooks, they feed on host
tissue as well as on nutrients from the host hemolymph. The larva is equipped with a thick
anal hook and numerous cuticular spines supporting the larva by its attachment to the
respiratory funnel. Endoparasitoid penetration induces the host cellular defense resulting into
the formation of the hemocyte capsule consisting of multi-cellular sheaths. This enveloping
capsule later undergoes the melanization, which is mostly obvious towards the posterior part
of endoparasitoid. The endoparasitoid uses the encapsulation response to build a respiratory
funnel, built-up from the modified host integument, leading to the host surface. Encapsulated
larva remains attached to the respiratory funnel via an anal hook and cuticular spines until
fully developed. Furthermore, additional immunohistochemical analysis and simulated
parasitization with polybead microspheres were used to study host-parasitoid interactions.
Keywords: endoparasitoid, respiratory tunnel, capsule, melanin, cephalopharyngeal skeleton
2
Introduction
Encapsulation is an essential defense reaction in insects, in which the presence of
foreign material, too large for phagocytosis, induces its gradual envelopment by a
hematocytes, finally forming a multilayered capsule. This cascade event involves the initial
recognition of foreign target by non-phagocytic granulocytes, inducing migration of
phagocytically active plasmatocytes ready to adhere and build-up the capsule (Ratner &
Vinson 1983). It seems that the cooperation between one or more classes of hemocytes is
mediated by the cytokines and adhesion molecules (Strand & Pech 1995). The structure of
hemocyte capsule has been considered to be consistent in all studied arthropods. Typical
capsule consists of 5-30 layers of flattened hemocytes packed tightly together, occasionally
exhibiting desmosomes or septate junctions between the individual cells (Baerwald 1979).
The inner cellular layer of capsule surrounding the living organisms often undergoes a
process called the melanization, in which the melanin is deposited on the surface of foreign
material. Endoparasitoids develop within the body cavity of their host and thus face a risk to
be encapsulated. In Hymenoptera, the encapsulation is typically prevented by the venom and
associated constituents (e.g. polyDNA viruses) injected during oviposition (Feener & Brown
1997). In Tachinidae, with the exception of a few groups equipped with piercing structures,
the female flies lacking an ovipositor deposit their eggs externally on or near the host and the
newly hatched larva is enforced to actively penetrate into the host body. Without the injection
of paralytic poisons, used by hymenopterans to immobilize the host immune system, the
tachinid larvae had to find other ways to overcome the host immune defense (Stireman 2006).
They have evolved two main strategies how to avoid the host immune system; larvae maintain
the contact with the atmospheric oxygen by attaching their posterior spiracles to the host
tracheal system or to the respiratory funnel created primarily from the host integument
(Eggleton & Belshaw 1993), and turn the host immune system to their own advantage by
reorganizing the products of host responses into the respiratory funnel (Salt 1968). In some
tachinid species, larvae migrate to a specific host tissue to be protected and remain there
hidden until they are ready to consume the host (Feener & Brown 1997). These strategies
developed by tachinids to overcome host physiological defenses without any host-specific
adaptations could also explain their wide host range. Tachinids adapted to the significantly
narrower host ranges usually remain free in the hemocoel without forming a respiratory
funnel (Belshaw 1994).
Here we focus on some aspects accompanying the penetration and encapsulation of
polyphagous gregarious larval endoparasitoid Exorista larvarum within its factitious host
3
Galleria mellonella. The oviposition strategy and chronology of the development of E.
larvarum has been described previously (Dindo 1999, Hafez 1953a,b,c, Michalková et al.
2009). The larvae of E. larvarum offer an ideal model to study the host-parasitoid
interactions, in which they turn the encapsulation response into formation of the respiratory
funnel, built-up from the modified host integument, and leading to the host surface. Despite of
few studies describing the structure of the hemocyte capsule and respiratory funnel in
different tachinid species (Mellini & Cucchi 1965; Baronio et al. 1974), there is still lack of
sufficient data on tachinids. Present study offers a complete analysis of attachment stratedy
and encapsulation using advanced histological techniques along with the scanning electron
microscopy.
Material and methods
Insect rearing and experimental parasitization. Colonies of parasitoid fly, Exorista
larvarum L. were reared in laboratory conditions as described elsewhere (Michalková et al.
2009). For experimental parasitization, last instar larvae of G. mellonella were exposed to
fertile female tachinid flies held in the cage and generally removed after 3-4 parasitoid eggs
had been laid on their body surface. To confirm the successful penetration of the newly
hatched tachinid larvae, three days later, the wax moth larvae were examined under a
stereobinocular microscope. Successfully parasitized caterpillars were kept in plastic boxes
until dissection, carried out on a daily basis following parasitoid egg hatching. Body parts
with developing larvae were dissected and fixed for subsequent tissue processing.
Histological procedure. Specimens were fixed in AFA (Alcohol-Formalin-Acetic
Acid) fixative and processed using standard histological methods. The material was
embedded in Histoplast II and sectioned; 7 m thick sections were stained with hematoxylineosin,
Masson's trichrome (green) or with Lillie's ferric-ferricyanide melanin staining method
with or without acetocarmine counterstain (Humason 1967). Additional specimens were fixed
in fixatives containing paraformaldehyde (Karnovsky's fixative or Baker's calcium formol)
and processed for cryosectioning in order to obtain sections of better quality. Frozen sections
were stained with Heidenhain's azan or hematoxylin-eosin. Histological sections were
investigated using an Olympus BX51 light microscope.
Scanning electron microscopy. The specimens were fixed in absolute ethanol and then
processed as describe in Michalková et al. (2009). Examination was undertaken using a JEOL
JSM-6300 Scanning Electron Microscope (SEM).
4
Mouse immunization. Mice were immunized with salivary homogenate of E. larvarum,
three times in two week intervals. Every mouse was injected with 100 l of antigen and IFA
solution (1:1). Two weeks after the final injection, blood from the mice was collected and
allowed to sediment overnight at 4°C. After the sedimentation, serum was asphyxiated and
cleared by centrifugation at 1000g. Antigen-binding capability of the sera was assessed by dot
blot.
Injection of polybead microspheres. Polybead polystyrene 90 m microspheres
(Polysciences) were used as encapsulation targets. A total volume of 200 l of polybeads was
coated with 3 l of antigens obtained from tachinid salivary homogenate with 1% BSA and
incubated during the night at 4°C, three times centrifugated and washed in PBS, blocked in
1% BSA for 30 min, three times centrifugated and washed in PBS and diluted in Ringer
solution. The coated polybead microspheres in insect physiological solution (1 l) were
injected into the larvae of G. mellonella using a micromanipulator. Control wax moth larvae
were injected with polybead microspheres blocked in 1% BSA alone without the antigen coat.
Indirect immunofluorescence. Deparaffined histological sections were incubated with
anti-saliva immune serum diluted in PBS (1:50) for 30 min at 37°C, three times washed with
PBS for 5 min and incubated with FITC-conjugated anti-mouse IgG (Sigma-Aldrich) diluted
in PBS (1:300) for 30 min at 37°C. Secondary antibody was flicked away and preparations
were counterstained with Evans blue (1:20000) for 3 min, two times washed with PBS for 5
min, mounted in anti-fade based on 2.5% DABCO (Sigma-Aldrich) mixed with glycerol and
0.1 M PBS. Controls were performed with FITC-conjugated anti-mouse IgG alone without
primary antibody. Preparations were observed and documented using an Olympus BX60
fluorescence microscope fitted with a WB filter cube, having an excitation range of 450-480
nm.
Results
The first instar larva
After the egg hatching, the larvae actively penetrate the host integument using the
strong, well-developed cephalopharyngeal skeleton and prominent body musculature. The
first instar tachinid larva remains attached to the host tissue by spines located on posterior
abdominal segment; no anal hook is observed at this stage. A dark brown to black spot visible
on the surface of host integument, as a result of cuticle melanization, indicated the area of
tachinid larva penetration (Fig. 1, 2). Larvae pull the host tissue while penetrating the host
integument to build the primary respiratory funnel (Fig. 3, 4, 7). The invading parasitoid
5
induces the formation of a hemocyte capsule as a result of the host cellular defense. This
structure develops gradually by attachment of granulocytes to invading endoparasitoid and
their degranulation, followed by a directed migration of plasmatocytes - the phagocytically
active components of host cellular defense, their aggregation and final adhesion to tachinid
larva. Completely developed capsule consists of several (more than 10) layers of flattened
hemocytes packed tightly together (Fig. 6, 9, 10). The intense red staining with Heidenhain's
azan demonstrated the acidophilic nature of cellular components forming the capsule in the
area of the respiratory funnel and the cell lysis resulting in the presence of cellular debris and
isolated nuclei incorporated into the capsular sheaths (Fig. 3-5). Its architecture changes
towards the respiratory funnel; the place of formerly deposited eggs on the host surface
exhibits strong necrosis accompanied by melanization (Fig. 4, 7-9) and the integumental
entrance point connected with the basis of respiratory funnel are typical by a strong
melanization along with the tissue hardening (Fig. 3-10). Melanization of host tissue has been
confirmed using a specific melanin staining methods as described above (Fig. 6-10). In the
first tachinid instar, the respiratory funnel is fixed to the adjacent host tissue by two closely
unspecified appendages, symmetrically oriented from each other, deeply embedded in the
host tissue (Fig. 4, 9).
The second instar larva
The larvae are still attached to the respiratory funnel via an anal hook located between
two anal lobes, and supported by sturdy spines on the posterior segment (Fig. 11). The
hemocyte capsule appears being intact and comprising of numerous compressed layers (Fig.
15). At this stage of parasitization, the host ceases its movement and food intake. Apparent
reduction and disintegration of host tissue, especially the fat body, surrounding the
encapsulated, rapidly developing parasitoid can be seen (Fig. 13, 14, 16). This disintegration
is obvious especially nearby the parasitoid anterior part with the mouth hooks stuck into the
hemocyte capsule (Fig. 11). The internal host organs are obviously compressed by the
parasitoid increasing in size and occupying ever-greater space of the host body (Fig. 15).
Larval exuvium with typical integumental spines is seen to be incorporated in the respiratory
funnel (Fig. 12) and strengthen this structure, which is due to the rapidly parasitoid growing
under permanent mechanical stress. The initial part of respiratory funnel, formed by the
modified host epidermis pulled by the parasitoid while penetrating, undergoes a strong
cuticular melanization (Fig. 24-28, 30). The aperture region is strongly hardened (Fig. 13, 17,
18) and this avoids an involuntary compression of respiratory funnel by the host movements.
6
This way the parasitoid remains safe with a continuous oxygen intake. Hemocyte capsule
contains thick melanized layers with patchy pattern especially in the posterior part near to the
respiratory funnel, though some single-layered melanized areas can be seen also in its anterior
part (Fig. 11, 19-23). The granulocytes attached to the respiratory funnel forming the first
layers of capsule are clearly to be recognized (Fig. 25). In addition to melanized layer stained
green with ferric-ferricyanide, the luminal surface of respiratory funnel and posterior part of
capsule exhibits an always non-stained, homogenous compact layer (Fig. 21, 24).
The third instar larva
The host is lethargic without any movement. Later on, the larva abandons the capsule
and moves free in the body cavity of already dead host, feeding on the residual host tissue. At
the end, before leaving the host body, the host tissues were completely destroyed and the
internal organs were fully disintegrated exhibiting a liquid appearance. The tachinid larva
consumes remaining body fluids and tissues except the cuticle with appendages and head
capsule.
Injection of polybead microspheres
Simulated parasitization using antigen coated polybead microspheres gently injected
into the wax moth caterpillars demonstrated the acute host response to the presence of foreign
antigens, but not to the object alone as showed the control test with microspheres not coated
with tachinid antigens. The simulated parasitization was observed in three dial period, during
which we noted no changes in wax moth behavior or physiological response in the first day
after microspheres injection. Following day the insects with antigen-coated microspheres
exhibited a ceased activity and black spots visible through the tegument, as the first obvious
traces of subtegumental melanization, could be noticed. In the third day after microspheres
injection, all caterpillars were dead exhibiting a strong evidence of melanization surrounding
the microspheres coated with antigen (Fig. 35, 36), however, the caterpillars injected with
microspheres without antigen showed no reaction.
Indirect immunofluorescence
Labeling of paraffin sections with antibody cocktail raised against tachinid salivary
glands including saliva, showed mediate signal corresponding to the tachinid salivary glands,
however, no labeling has been observed in the host tissue surrounding capsule (Fig. 38-40).
An intense signal noticed in the luminal side of hemocyte capsule, especially thick towards
7
the respiratory funnel (Fig. 38-40), can be not considered a priory to be the specific labeling,
as we can not exclude the possibility of autofluorescence often described in melanin. The area
of the necrotic host tissue near the parasitoid entrance point remained unlabelled in all
sections (Fig. 37, 38).
Discussion
Endoparasitoid insects must to deal directly with the host immune system for which
specialized adaptations are required. Encapsulated parasitoids may die of suffocation,
starvation or physical prevention of development. Partially encapsulated parasitoids, however,
are able to survive and continue to develop normally (Van den Bosch 1964). Generally,
parasites must evade or disrupt the host immune system to survive (Rizki & Rizki 1990), but
dipteran parasitoids try to benefit on host immune responses by building the respiratory
funnel (Feener & Brown 1997). Vinson (1990) divided strategies of how the parasitoids
handle with the host insect immune system in 5 subcategories: avoidance, evasion,
destruction, suppression, and subversion characterized for tachinids building the respiratory
funnel, where the parasitoid development is aided by the host response.
Mellini (1990) previously described the initial phase of tachinid penetration through
the host tegument, enabled by the use of saliva to soften the host cuticle and by the
mechanical action of the mouth hook. This way, larvae are able to penetrate three different
hosts successively as long as they do not remain in individual victims longer than 15 minutes.
We have observed females ovipositing their eggs on the soft membranous areas
interconnecting the sclerites, what facilitates larval penetration into the host body. An active
defense of host insect, oriented towards the parasitoid eggs laid on their exoskeleton or
penetrating larvae, by lacerating or rubbing them away from the body is also known and can
be clearly observed in Spodoptera litoralis larvae exhibiting remarkable body flexibility. As
the larvae of G. mellonella do not exhibit such extensive active defense, they represent an
ideal model for tachinid factitious host. In addition, by parasitization with gregarious
parasitoids, the hosts are unable to discard all of laid eggs and thus the chance for parasitoid
to develop increases significantly.
The encapsulation has been already described in various studies dealing predominantly
with hymenopteran parasitoids. Detailed description of encapsulation in tachinid-host insect
system has been already published on Macquartia chalconota parasitizing the beetle
Chrysomela herbacea (Baronio et al. 1974). The initial phase of hematocyte sheath
development started with the formation of a thin membrane that began to differentiate
8
between 1.30 and 2.30 hours after the endoparasitoid penetration. Mentioned membrane
originated from mucopolysaccharides freed by the hemocytes involved in the healing process
in the area of parasitoid penetration opening. The next development of the multicellular
sheath was comparable to those already described in other studies; comprising of the
apposition of hemocytes, their aggregation and subsequent flattening, however, in contrast to
Baronio et al. (1974), the nuclei were still to be recognized in flattened hemocytes when
observed by higher magnification. The respiratory funnel and hemocyte capsule exhibiting
similar architecture have been described in Steiniella callida parasitizing the chrysomelid
beetle Melasoma populi (Mellini & Cucchi 1965).
Experimental study on typical two-phase encapsulation process in G. mellonella by
inoculation of foreign implants demonstrated that the initial lysis of the granular hemocytes
contacting foreign material starts within 5 min after implantation (Schmit & Ratcliffe 1977).
The second phase, the plasmatocyte adhesion to foreign target, was restricted to the sites of
granular cell lysis only. The melanization of capsule has been observed in areas exposed to
direct contact with implant surface, while the inner region of complete capsule was built up
from the flattened and necrotic cells. Similarly to our preparations, granular cells remnants
were also occasionally observed. Interestingly, in the cross sections of the the respiratory
funnel surrounding the first instar tachinid larvae, we repeatedly noticed two closely
unspecified appendages, deeply embedded in the host tissue, probably serving as a fixation to
the host tissue. According to Salt (1970), maturing cellular capsules tend to darken and
harden, especially in the area of strongly melanized respiratory funnel, representing the oldest
part of host immune response. This hardness prevents compression of surrounding host tissue
during host's movement and facilitates the supply of atmospheric oxygen towards the
endoparasitoid.
As shown in previous study, the body structure of E. larvarum indicates that this
tachinid is perfectly adapted for the parasitic way of life (Michalková et al. 2009). As the
larvae lack the intestinal retention structures, they defecate throughout their development as
observed by rearing them on artificial media (Dindo et al. 1999) and this fecal material is
deposited and encrusted in the luminal side of the respiratory funnel (Gardenghi & Mellini
1995), often seen as a compact luminal layer of the respiratory funnel. As their integument is
surprisingly thin and seemingly lacks the typical protective function, the tachinid larvae
probably take advantage of the protection in hemocyte capsule from the host internal
environment and defense mechanisms (Michalková et al. 2009).
9
The results of injection of polybead microspheres in this study are of particular
interest. The wax moth caterpillars died three days after injection of antigen coated polybeads
in agreement with the natural parasitization with tachinids, where the third instar tachinid
larva fully develops in 3 to 4 days after hatching the egg and finally kills the host. The
caterpillars injected with antigen coated polybeads behaved and exhibited the same responses
in the same time period as observed in real parasitization, contrary to control group injected
with poybeads without antigen and exhibiting no obvious response to injection. Indirect
immunofluorescence did not show any labeling of tachinid antigens in the surrounding host
tissue, although we previously expected some extracts of parasitoid salivary glands as the
result of the extra-intestinal digestion at least near the area of hemocyte capsule interrupted by
tachinid cephalopharyngeal skeleton enabling the food intake. As all the necrotic parts of the
host tissue remained unlabelled, the intense signal noticed in the luminal side of capsule may
be related to the autofluorescence of melanin distributed in the capsule. These results could
support the hypothesis of exploitation and avoidance strategy of tachinid larva hidden in the
cover consisting of host own immune products. Similar strategy to stay hidden in hemocyte
capsule, in the host products, can be find in endoparasitic Strepsiptera, which remain enclosed
in a host-derived epidermal bag not recognized by the host (Kathirithamby et al. 2003). This
strategy of how to avoid the host immune responses and camouflage in host own material
leads into the unusual polyphagy, attacking dozens of host species belonging to multiple
families (Belshaw 1994, Kathirithamby et al. 2003, Stireman 2006).
Acknowledgement
The authors are gratefully indebted to Jana Benešová (Dept. of Experimental Biology, MU)
for the help with the rearing of Galleria mellonella and Romana Šebestová (Dept. of
Histology and Embryology, MU) for her help with cryosectioning. We also thank Elisa
Marchetti and Laura Depalo (DiSTA, University of Bologna) for their help with the rearing in
Italy. This study was financially supported by the MSM 0021622416, by the GAČR
524/05/H536 and by the Italian Ministry of University and Research PRIN 2005.
10
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11
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12
Figures 1-2. Necrosis of Gallera mellonela integument. 1. The host integument exhibiting
obvious necrosis at the entrance point (arrow) of the first instar tachinid larva, with the empty
egg shell (arrowhead). Scale bar = 250 m. 2. The advanced necrosis of host integument
surrounding the egg remnants (arrowhead) at the stage of second instar tachinid larva. Scale
bar = 1000 m.
13
Figures 3-10. Histological sections of the first instar larva of Exorista larvarum encapsulated
within the host Galleria mellonella. 3. The longitudinal section of tachinid larva (*)
surrounded with hemocyte capsule (ca); host tissue (ht), respiratory funnel (rf). Scale bar =
200 m. 4. The cross section of tachinid larva (*) surrounded with prominent respiratory
funnel (rf); area of host integument exhibiting a strong cuticular melanization (arrowhead).
Note the two unknown appendages inserted into the host tissue (arrows). Scale bar = 100 m.
5. Detail of Fig. 4 showing the wall of respiratory funnel (rf); the tegument of tachinid larva
covered with numerous spines (arrowhead), unknown appendage (arrow). Scale bar = 20 m.
6. Section stained for melanin showing the distribution of melanin (intense green to dark
green) along the luminal side of hemocyte capsule (ca) and respiratory funnel (rf); tachinid
larva (*). Scale bar = 100 m. 7. Section of entrance point with cuticular melanization
(arrowheads) and respiratory funnel (rf) demonstrating the presence of melanin (dark green).
Scale bar = 200 m. 8. Detail of the cuticular melanization (dark green) in the host
integument (arrowhead); host epidermis (he). Scale bar = 20 m. 9. The cross section of
tachinid larva (*) surrounded with respiratory funnel (rf) with obvious melanization (green); a
strongly melanized host integument near the parasitoid entrance point (arrowhead); unknown
slightly melanized appendages (arrows). Scale bar = 200 m. 10. Detail of Fig. 9 showing the
melanized wall of respiratory funnel (rf) and the tegument of tachinid larva with numerous
spines (arrowheads). Scale bar = 20 m.
Heidenhain's azan (Fig. 1-3); Lillie's ferric-ferricyanide melanin staining with (Fig. 7-10) or
without (Fig. 4) acetocarmine counterstain.
14
15
Figures 11-18. Second instar larva of Exorista larvarum encapsulated within the host
Galleria mellonella. 11. Longitudinal section of encapsulated tachinid larva (*) stained for
melanin showing the distribution of melanin (intense green to dark green) along the luminal
side of hemocyte capsule (ca) and respiratory funnel (rf), and in the area of cephalopharyngeal
skeleton (arrowhead); numerous cuticular spines covering tachinid integument (arrows). Scale
bar = 200 m. 12. Detail of the wall of respiratory funnel stained for melanin; incorporated
tachinid exuvium with obvious cuticular spines (arrows). Scale bar = 20 m. 13. The
longitudinal section of tachinid larva surrounded with hemocyte capsule (ca); tachinid
digestive tract (dt), tachinid posterior spiracle (arrowhead), prominent respiratory funnel (rf)
with a thick layer of necrotic tissue. Scale bar = 500 m. 14. The tangential section of tachinid
larva surrounded with hemocyte capsule (ca); cephalopharyngeal skeleton (arrow), respiratory
funnel (rf). Scale bar = 100 m. 15. The tachinid larva surrounded with a thick hemocyte
capsule (ca); tachinid digestive tract (dt), tachinid integument (arrowheads). Scale bar = 250
m. 16. Section of two encapsulated tachinid larvae (*); hemocyte capsule (ca), respiratory
funnel (rf), tachinid digestive tract (dt). Scale bar = 250 m. 17. Section of encapsulated
tachinid larva (*); hemocyte capsule (ca), respiratory funnel (rf), tachinid integument
(arrowheads). Scale bar = 250 m. 18. Scanning electron micrograph showing E. larvarum
larva (*) sitting in the respiratory funnel (rf). Scale bar = 500 m.
Lillie's ferric-ferricyanide melanin staining (Fig. 11, 12); hematoxylin-eosin (Fig. 13-15);
Masson's trichrome (Fig. 16, 17).
16
17
Figures 19-26. Modifications of the host tissue surrounding the second instar larva of
Exorista larvarum. 19. Longitudinal section of encapsulated tachinid larva (*) stained for
melanin showing the distribution of melanin (intense green to dark green) along the luminal
side of hemocyte capsule (ca) and respiratory funnel (rf). Scale bar = 100 m. 20, 21. Detail
of the wall of respiratory funnel showing a strong melanization (in green) and deposited fecal
material (*) of tachinid larva; flattened hemocytes packed tightly together and forming the
sheath (arrows). Scale bars = 20 m. 22, 23. Detail of the hemocyte capsule exhibiting a thin
melanized layer (in green, arrowhead) and multicellular sheaths (arrows). Scale bars = 20 m.
24. The modification of host tissue in the parasitoid entrance point (arrows) and respiratory
funnel (rf); host cuticle (hc), host epidermis (he), host tissue (ht), tachinid larva (*). Scale bar
= 100 m. 25. Detail of Fg. 24 showing the strong cuticular melanization (cm) of the host
tissue; remnants of granulocytes (arrowheads), multicellular sheath consisting of
plasmatocytes (arrow), host modified epidermis (hme), respiratory funnel (rf). Scale bar = 20
m. 26. Longitudinal section of respiratory funnel (rf) and part of hemocyte capsule (ca).
Scale bar = 200 m.
Lillie's ferric-ferricyanide melanin staining with acetocarmine counterstain (Fig. 19-23);
hematoxylin-eosin (Fig. 24, 25); Heidenhain's azan (Fig. 26).
18
19
Figures 27-34. Modifications of the host tissue surrounding the second instar larva of
Exorista larvarum.. 27. Longitudinal section of the parasitoid's entrance point and respiratory
funnel (rf) built up from the modified host epidermis (arrowhead).and cuticle (arrow);
melanization obvious in the wall of respiratory funnel (double arrowhead), destroyed host
tissue (ht). Scale bar = 200 m. 28. Detail of the wall of respiratory funnel (rf) near the
entrance point; modified host epidermis (arrowhead).and cuticle (arrow). Scale bar = 50 m.
29. Tangential section of the hemocyte capsule (ca) passing into the respiratory funnel (rf).
Scale bar = 200 m. 30. Detail of Fig. 27 showing the wall of respiratory funnel (rf) with
tachinid faeces deposited at its luminal side; melanization (arrowhead), host modified
epidermis (hme) and host modified cuticle (arrow). Scale bar = 50 m. 31. Longitudinal
section of the hemocyte capsule (ca) and respiratory funnel (rf). Scale bar = 1000 m. 32.
Detail of the respiratory funnel (rf) and multicellular sheaths (arrow). Scale bar = 50 m. 33.
Wall of the respiratory funnel (rf) with remnants of granulocytes (arrowhead) and
multicellular sheath consisting of flattened plasmatocytes (arrow). Scale bar = 20 m. 34.
Detail of the area interconnecting the respiratory funnel and hemocyte capsule; melanized
layers (arrowheads), multicellular sheath (arrow). Scale bar = 50 m.
Masson's trichrome (Fig. 27, 28, 30-32); Heidenhain's azan (Fig. 29); hematoxylin-eosin
(Fig. 33, 34).
20
21
Figures 35-40. Immunological experiments on Galleria mellonella. 35, 36. Host response,
necrosis (arrows) to injected polybeads covered with tachinid antigens. Scale bare = 1000 m.
37, 38. Longitudinal sections of the second instar tachinid larva (*) encapsulated within the
host treated with anti-saliva immune serum; labeled tachinid salivary glands (arrowheads),
hemocyte capsule (ca), respiratory funnel (rf). Fig. 37 is a control LM micrograph. Scale bar
= 100 m. 39. Detail of Fig. 38, with labeled tachinid salivary gland (arrowhead) and
hemocyte capsule (ca). Scale bar = 50 m. 40. Labeled tachinid salivary glands with antisaliva
immune serum (arrowheads) and surrounding hemocyte capsule (ca). Scale bar = 100
m.
22
Paper C
New host and country records for European Tachinidae (Diptera).
MÜCKSTEIN P., TSCHORSNIG H.-P., VAŇHARA J. & MICHALKOVÁ V.
published in Entomologica Fennica 18:179-183 (2007)
New host and country records
for European Tachinidae (Diptera)
Petr Mückstein, Hans-Peter Tschorsnig, Jaromír Vahara* & Veronika Michalková
Mückstein, P., Tschorsnig, H.-P., Vahara, J. & Michalková, V. 2007: New host
and country recordsfor European Tachinidae(Diptera).  Entomol. Fennica18:
179183.
The paper presents host records for 17 species of Tachinidae (of subfamilies
Exoristinae and Tachininae) from the Czech Republic, Slovakia, Austria, Croatia,
Macedonia, Italy, Spain, Portugal, and Bulgaria. New parasitoid-host couples
are Exorista larvarum  Melanchra pisi; Exorista segregata  Catocala
nymphaea; Sturmia bella  Hadena compta; Spallanzania multisetosa  Cycnia
sordida (first host record); Tachina praeceps  Cucullia bubaceki; and Bithia
modesta  Bembecia megillaeformis. New country records of tachinid species
Rhacodinella apicata from the Czech Republic, Masicera pavoniae from Macedonia
and Bithia demotica from Portugal are presented.
P. Mückstein, Administration of the Protected Landscape Area árské vrchy,
Brnnská 39, CZ  591 01 ár nad Sázavou, Czech Republic; E-mail:
muckstein@email.cz
H.-P. Tschorsnig, Staatliches Museum für Naturkunde, Rosenstein 1, D-70 191
Stuttgart, Germany; E-mail: tschorsnig.smns@naturkundemuseum-bw.de
J. Vahara (*corresponding author), Masaryk University, Faculty of Science,
Kotláská 2, 611 37 Brno, Czech Republic; E-mail: vanhara@sci.muni.cz
V. Michalková, Masaryk University, Faculty of Science, Kotláská 2, 611 37
Brno, Czech Republic; E-mail: vmichalkova@yahoo.com
Received 20 October 2006, accepted 8 December 2006
1. Introduction
The tachinid fauna of Europe is well known, but
new records are done every year. Information on
distribution of the European Tachinidae is available
on-line in Fauna Europaea (Tschorsnig et al.
2005), for the Czech Republic see Vahara et al.
(2004). New on-line and CD versions of the
checklist for the Czech Republic and Slovakia
were completed by Vahara and Tschorsnig
(2006).
Alltachinidsareparasitoidsofinsectsorother
arthropods. Knowledge on their biology, including
host insect species is still rather poor. Many
problems concerning the host specificity of this
large family have not been solved yet, so every
host record is important. The short history concerning
the research on hosts. has been presented
in our previous paper (Mückstein et al. 2004), including
general references.
At present, the authors look for further tachinid
hosts by the systematic cooperation particularly
with lepidopterologists, who contribute
reared tachinids from their breeding of caterpil-
 Entomologica Fennica. 6 September 2007
lars from the Czech Republic. But findings from
this country yield only a small part of new results.
After 1989 our lepidopterologists started freely
move out for collection abroad, and so there are
new hosts especially from southern and western
Europe.
2. Material and methods
As far as not explicitly cited in the text, information
on hosts is based on a critical host catalogue
for the Czech Tachinidae which is currently compiled
by the same authors of the present paper and
on Tschorsnigs database on Palaearctic host records
(unpubl.).
The tachinid species were identified by the
authors of the present paper, hosts by the collectors.
Nomenclature and arrangement of the tachinids
follow Herting and Dely-Draskovits (1993),
the names of the lepidopteran hosts follow
Karsholt and Razowski (1996).
The material provided by various breeders is
storedinthecollectionsofP. Mückstein(cM) and
J. Vahara (cV).
3. Faunistic records
3.1. Exoristinae
3.1.1. Exorista larvarum (Linnaeus, 1758)
Material: 3 $$, reared from Melanchra pisi
(Linnaeus) (Lepidoptera, Noctuidae), Czech Republic,
Moravia, Peckov, 1984, leg. Z. Latvka
(cM, cV); 2 ##, reared from pupa of Euthrix
potatoria (Linnaeus) (Lepidoptera, Lasiocampidae),
Czech Republic, Bohemia, Týnit
nad Orlicí, 13.VII.1986, leg. J. Voda (cM).
Notes: A common tachinid which develops in
many lepidopteran families.
Melanchra pisi is a new host record.
3.1.2. Exorista segregata (Rondani, 1859)
Material: 1 $, reared from Catocala nymphaea
(Esper) (Lepidoptera, Noctuidae), Spain, Andalusia,
NW Sierra Nevada, El Molinillo near
Diezma, 1200 ma.s.l., VII.2004, leg. Z. Latvka
(cV).
Notes: Exorista segregata is an unspecialised
species (common in southern Europe), parasitising
many lepidopteran families.
Catocala nymphaea is a new host record.
3.1.3. Meigenia mutabilis (Fallén, 1810)
Material: 1 #, reared from larva of Gastrophysa
viridula (Degeer) (Coleoptera, Chrysomelidae),
Czech Republic, Praha, Ruzyn, 10.VI.2003, leg.
M. Honk (cM).
Notes: A common parasitoid of Coleoptera
larvae. Already known from the host Gastrophysa
viridula (many records).
Gastrophysa viridula is a new host record for
the Czech Republic.
3.1.4. Compsilura concinnata (Meigen 1824)
Material: 2 $$, reared from Aglaope infausta
(Linnaeus) (Lepidoptera, Zygaenidae), Portugal,
Bragança, VII.2004, leg. Z. Latvka (cV).
Notes: Compsilura concinnata is the tachinid
species with the highest number of known host
species (currently at least 230 in the Palaearctic).
Aglaopeinfaustawasalready knownashostfrom
Spain and France (Benlloch & Caizo 1934,
Herting 1960).
Aglaope infausta is a new host record for Por-
tugal.
3.1.5. Winthemia quadripustulata
(Fabricius, 1794)
Material: 1 $, reared from larva of Cucullia
lactucae (Denis & Schiffermüller) (Lepidoptera,
Noctuidae), Czech Republic, Rokytnice nad
Jizerou, 30.VII.2003, leg. L. Traxler (cM).
Notes: Hosts of this common species are
mainly Noctuidae. Herting (1960) records Cucullia
lactucae as a host from Germany.
Cucullia lactucae is a new host record for the
Czech Republic.
3.1.6. Phryxe prima
(Brauer & Bergenstamm, 1889)
Material: 1 $, reared from larva of Zygaena
carniolica (Scopoli) (Lepidoptera, Zygaenidae),
Czech Republic, Bohemia, Kvtná near Lito-
180 Mückstein et al.  ENTOMOL. FENNICA Vol. 18
myl, TCP Strán 6264C, 18.VII.1998, leg. B.
Mocek (cV).
Notes: A parasitoid of Zygaena spp. Already
known from the host Zygaena carniolica.
Zygaena carniolica is a new host record for
Phryxe prima from the Czech Republic.
3.1.7. Lydella stabulans (Meigen, 1824)
Material: 1 $, reared from Hydraecia micacea
(Esper) (Lepidoptera, Noctuidae), Czech Republic,
Bohemia, Píany near Louny, 10.VII.2002,
leg. edivý (cV).
Notes: A known parasitoid of noctuid larvae
which feed hidden in stems of herbaceous plants.
Hydraecia micacea is a common host of Lydella
stabulans.
Hydraecia micacea is a new host record for
the Czech Republic.
3.1.8. Huebneria affinis (Fallén, 1810)
Material:1#,rearedfromArctiacaja(Linnaeus)
(Lepidoptera, Arctiidae), Czech Republic, Bohemia,
Vracov, VII.1983, leg. Havel [59884]
(cV); 1 $, same host, Czech Republic, Moravia,
Karlova Studánka env., 29.V.2003, leg. L.
Traxler (cM); 1 #, same host, Slovakia,
Strácovské vrchy, iroká dolina, 18.VI.2004,
leg. J. Rybár (cV).
Notes: Usually a common parasitoid of
Arctiidae. There are numerous records from the
host Arctia caja.
ArctiacajaisanewhostrecordforHuebneria
affinis from the Czech Republic.
3.1.9. Carcelia rasa (Macquart, 1849)
Material: 1 #, 2 $$, reared from larva of Calliteara
pudibunda (Linnaeus) (Lepidoptera,
Lymantriidae), Czech Republic, Bohemia, Týnit
nad Orlicí, 16.XII.1986, leg. J. Voda (cM).
Notes: This species develops in Lymantriidae.
Known frommany countries as parasitoid
of Calliteara pudibunda.
Calliteara pudibunda is a new host record for
Carcelia rasa from the Czech Republic.
3.1.10. Rhacodinella apicata (Pandellé, 1896)
Material: 1 $ (in bad condition), reared from
pupa of Phyllodesma ilicifolia (Linnaeus) (Lepidoptera,
Lasiocampidae), Czech Republic, Píbram,
28.VI.2002, leg. L. Traxler (cM).
Notes: An unspecialised and normally rare
species. The host Phyllodesma ilicifolia is already
recorded by Herting (1960) from Switzer-
land.
The tachinid species Rhacodinella apicata is
recorded for the first time from the Czech Repub-
lic.
3.1.11. Sturmia bella (Meigen, 1824)
Material: 1 $, reared from Hadena compta
(Denis & Schiffermüller) (Lepidoptera, Noctuidae),
Austria, Dachstein-Gruppe, Ramsau a. D.,
46°54 N, 13°36 E, 26.31.VII.2003, leg. V.
ervenka (cV).
Notes: A common species which is predominantly
reared from Nymphalidae, but also members
of several other lepidopteran families are
known as hosts, including a few Noctuidae.
Hadena compta is a new host record.
3.1.12. Masicera pavoniae
(Robineau-Desvoidy, 1830)
Material: 4 ##, 2 $$, reared from Saturnia pyri
(Denis & Schiffermüller) (Lepidoptera, Saturniidae),
Croatia, Istria, Rabac, IV.2005, leg. Z.
Latvka (cV); 2 $$, same host, Bulgaria, Zlaté
Písky, 5.V.1963, leg. V. Zeman (cM); 2 ##, 5
$$, reared from Saturnia spini (Denis &
Schiffermüller), Macedonia, Tetovo, 15.V.1992,
leg. L. Traxler (cM.)
Notes: Masicera pavoniae is  as a main parasitoid
of Saturnia spp.  often reared from S. pyri
and S. spini. One of the four males of the series
from Croatia is aberrant because it shows two
proclinate orbital bristles (instead of one).
The tachinid species Masicera pavoniae is recorded
for the first time from Macedonia.
ENTOMOL. FENNICA Vol. 18  New host and country records for tachinids 181
3.1.13. Spallanzania multisetosa
(Rondani, 1859)
Material: 1 $, reared from Cycnia sordida
(Hübner) (Lepidoptera, Arctiidae), Italy, Vall
dAosta, VII.2003, (cV).
Notes: Spallanzania multisetosa is not rare in
southern Europe and the warmer parts of the
Alps. Other species of the genus Spallanzania,
S. hebes and S. rectistylum, are known as parasitoids
of Noctuidae. It would be of interest to revise
the old record of Cnephalia bucephala
reared from the arctiid Phragmatobia fuliginosa,
as it cannot be ruled out that this might have been
S. multisetosa. Thisrecord, however, isonly cited
in a footnote of Bezzi (1907) and given without
any reference.
Cycnia sordida is the first host record.
3.2. Tachininae
3.2.1. Tachina praeceps Meigen, 1824
Material: 1 #, reared from Cucullia bubaceki
Kitt (Lepidoptera, Noctuidae), Spain, Aragón,
Belchite, 41°2530 N, 00°4502 E, 22.X.2003,
leg. V. ervenka (cV).
Notes: A predominantly southern European
species which develops in various host families
(Lymantriidae, Lasiocampidae, Arctiidae, Noctuidae,
Sphingidae).
Cucullia bubaceki is a new host record.
3.2.2. Bithia demotica (Egger, 1861)
Material: 1 $, reared from Bembecia psoraleae
Bartsch & Bettag (Lepidoptera, Sesiidae), Portugal,
Algarve, Messines Baixo, 25.VII.2003, leg.
Z. Latvka (cV).
Notes: A parasitoid of Sesiidae (Bembecia,
Chamaesphecia, Synansphecia). Bembecia psoraleae
was recorded as host by Bartsch and
Bettag (1997) from Spain.
The tachinid species Bithia demotica is recorded
for the first time from Portugal.
3.2.3. Bithia glirina (Rondani, 1861)
Material: 1 #, 1 $, reared from larva of Chamaesphecia
empiformis (Esper) (Lepidoptera, Sesiidae),
Slovakia, Hrádok nad Váhom, 1987, leg.
Z. Latvka (cM).
Notes: A parasitoid of Sesiidae (Chamaesphecia,
Bembecia). Chamaesphecia empiformis
was already recorded as host by Herting (1960).
Chamaesphecia empiformis is the first host
record for Slovakia.
3.2.4. Bithia modesta (Meigen, 1824)
Material: 3 ##, 1 $, reared from larva of Bembecia
megillaeformis (Hübner) (Lepidoptera,
Sesiidae), Slovakia, Zádiel, 25.VII.1987, 1990
and 1993, leg. Z. Latvka (cM, cV); 2 $$ (in
bad condition), reared from Synansphecia triannuliformis
(Freyer) (Lepidoptera, Sesiidae),
Bulgaria, Miurin, VI.1982, leg. Z. Latvka
(cV).
Notes: A parasitoid of Sesiidae (Bembecia,
Synansphecia, Pyropteron).
Bembecia megillaeformis is a new host re-
cord.
Acknowledgements. The reared material was kindly provided
by L. Traxler (Pardubice, CZ), B. Mocek (Hradec
Králové, CZ), Z. Latvka (Brno, CZ) and P. Bituík
(Bánská tiavnica, SK). For financial supporting we are
indebted to the Ministry of Education/Masaryk University
(MSM 0021622416) and the Grant Agency CR (524/
05/H536).
References
Bartsch, D. & Bettag, E. 1997: Eine neue Art der Gattung
Bembecia Hübner, 1819 aus Südwesteuropa: Bembecia
psoraleae spec. nov. (Lepidoptera: Sesiidae). 
Nachr. ent. Ver. Apollo 18: 2940.
Benlloch, M. & Caizo, J. del 1934: Las plagas de Aglaope
infausta L.  Bol. Pat. veg. Ent. agric. 7: 115129.
Bezzi, M. 1907: Tachinidae.  In: Becker, T., Bezzi, M.,
Kertesz, K. & Stein, P. (eds.), Katalog der paläarktischen
Dipteren 3: 189597, Budapest.
Herting, B. 1960: Biologie der westpaläarktischen Raupenfliegen
(Dipt., Tachinidae).  Monographien zur
angewandten Entomologie 16. P. Parey, Berlin. 188
pp.
Herting, B. & Dely-Draskovits, A. 1993: Family Tachini-
182 Mückstein et al.  ENTOMOL. FENNICA Vol. 18
dae.  In: Soós, A. & Papp, L. (eds), Catalogue of Palaearctic
Diptera 13: 118624. Hung. Nat. Hist. Mus.,
Budapest.
Karsholt,, O. & Razowski, J. 1996: The Lepidoptera of Europe.
A distributional checklist.  Apollo Books,
Stenstrup. 380 pp.
Mückstein, P., Tschorsnig, H.-P. & Vahara., J. 2004: Some
new host records of West Palaearctic Tachinidae
(Diptera).  In: Bituík, P. (ed.), Dipterologica bohemoslovaca,
12. Acta Fac. Ecol. Zvolen 12 (Suppl.):
111113.
Tschorsnig, H.-P., Richter, V. A., Cerretti, P., Zeegers, T.,
Bergström, C., Vahara, J., Van de Weyer, G., Bystrowski,
C., Raper, C., Ziegler, J. & Hubenov, Z.
2005: Tachinidae.  In: Fauna Europaea Service, 1.2
[www document]. URL http://www.faunaeur.org (accessed
June 2006).
Vahara, J. & Tschorsnig, H.-P., 2006. Tachinidae Robineau-Desvoidy
1830.  In: Jedlika, L., Kúdela, M.,
Stloukalová, V. (eds), Checklist of Diptera of the
Czech Republic and Slovakia.  Electronic version 1,
CD ROM edition, Comenius Univ., Bratislava.
Vahara, J., Tschorsnig, H.-P. & Barták., M. 2004: New
records of Tachinidae (Diptera) from the Czech Republic
and Slovakia, with a revised check-list.  Stud.
Dipt. 10 (2003): 679701.
ENTOMOL. FENNICA Vol. 18  New host and country records for tachinids 183
Paper D
Annotated host catalogue for the Tachinidae (Diptera) of the Czech Republic.
VAŇHARA J., TSCHORSNIG H.-P., HERTING B., MÜCKSTEIN P. & MICHALKOVÁ V.
published in Entomologica Fennica 20: 22-48 (2009)
Annotated host catalogue for the Tachinidae (Diptera)
of the Czech Republic
Jaromír Vahara*, Hans-Peter Tschorsnig, Benno Herting, Petr Mückstein
& Veronika Michalková
Vahara, J., Tschorsnig, H.-P., Herting, B., Mückstein, P. & Michalková, V.
2009: Annotated host catalogue for the Tachinidae (Diptera) of the Czech Republic.
 Entomol. Fennica 20: 2248.
An annotated host catalogue is given for the Tachinidae of the Czech Republic. It
comprises149of476tachinidspecieswhicharecurrently knownfromthiscountry
(included the two new records cited below). 195 hosts are listed. The first host
records of Tachinidae date back to the second half of the 19th century. The bibliography
for the host records consists of 116 papers of 55 researchers. Several records
of hitherto unpublished material are included. Phryxe setifacies and
Anthomyiopsis plagioderae are first records for the Czech Republic.
J. Vahara (*corresponding author), Masaryk University, Faculty of Science,
Kotláská 2, CZ-611 37 Brno, Czech Republic, vanhara@sci.muni.cz
H.-P. Tschorsnig, Staatliches Museum für Naturkunde, Rosenstein 1, D-70 191
Stuttgart, Germany, tschorsnig.smns@naturkundemuseum-bw.de
P. Mückstein Administration of the Protected Landscape Area árské vrchy,
Brnnská 39, CZ-591 01 ár nad Sázavou, Czech Republic, muckstein
@email.cz
V. Michalková, Masaryk University, Faculty of Science, Kotláská 2, CZ-611 37
Brno, Czech Republic, vmichalkova@yahoo.com
Received 22 August 2007, accepted 21 January 2008
1. Introduction
Tachinidae are a very large and important dipteran
family of (mainly) insect parasitoids. This
study provides for the first time a critical host catalogue
for Tachinidae of the Czech Republic. It is
based on a compilation of literature records from
the Czech Republic as well as on revisions of material
(see below). There was a thorough search of
literature, but there is nevertheless no guarantee
that every record was found by the authors, because
host records of Czech Tachinidae are very
scattered in many journals, sometimes only appearing
as marginal footnotes.
The tachinid species are listed in their actual
valid nomenclature; probable misidentifications
are  if possible  tentatively corrected, but the
original name is always given in parenthesis, and
a comment is added in case of corrections. Wrong
or most probably wrong records that cannot be
corrected are set in square brackets. Possible reasons
for wrong records  which are not rare in literature
 are either misidentifications of the
tachinid or misassociated hosts or both. Misidentifications
of tachinids were frequent, especially
duringthe19thandearly 20thcentury,becauseof
the lack of true specialists of this difficult group,
but they still happen today when workers in ap-
 Entomologica Fennica. 30 March 2009
plied entomology hesitate to contact specialists.
Wrong host associations easily happen in soil
samples (forest litter) or in mass rearings when
various hosts can be hidden among the plant material.
Uncertainties in the tachinid identification
can be solved when the tachinid material is still
available for study, but even if a revision yields
the correct identification of a tachinid, the host
might nevertheless be wrong. Such cases cannot
be solved, but they can be at least commented on
or tentatively corrected in the light of the presentday
knowledge of the host-range of the tachinid
species (mainly based on revisions and compilations
of European host records done by Hans-Peter
Tschorsnig and the late Benno Herting).
References of the general parts (Introduction
etc.) quoted below are cited in full. They can be
foundintheReferencesofthispaper,orifthey include
factual host information, they are listed in
the section 5. Host record bibliography.
1.1. Short outline of the history
of host/tachinid investigations
in the Czech Republic
Tachinids from the present-day Czech Republic
have been studied since the first faunistic record
from 1791 (Preyssler et al. 1793), but the knowledge
of the diversity and especially of the hosts of
this family is as incomplete as in most other European
countries. The most recent information on
Czech Tachinidae is given by Vahara et al.
(2004), Tschorsnig et al. (2005), Vahara &
Tschorsnig (2006), and Mückstein et al. (2007).
The first data on tachinid hosts originate from
the second half of the 19th century when the area
of the present-day Czech Republic was part of the
old Austrian-Hungarian Empire (Kirchner 1861;
Wachtl 1882, 1886; Gold 1893, 1895; Mik &
Wachtl 1895). The rearings were first and foremost
intiated by studies on parasitoids of insect
pests which caused severe damage in coniferous
forests during that time. Wachtl reared also many
other lepidopterous hosts, but he published only a
small part of it.
Brauer and Bergenstamm, both experts on
Diptera, worked in the Museum at Vienna on
Tachinidae at the end of the 19th century. They
(Brauer & Bergenstamm 1891, 1894) also published
breeding records of Tachinidae, which
they periodically received from other colleagues
(e.g.R.vonStein).Thematerialofthelepidopterist
Richard Ritter von Stein (abbreviated
Rtt. v. Stein or R. v. Stein in Brauer & Bergenstamm
1891, 1894) most probably originates
from W Bohemia, namely Chodov (=Chodau),
nr.Karlovy Vary,wherevonSteinlived(Koleka
1993), but possibly also from other places in Bo-
hemiawherehecollected(e.g.umavaMts.etc.).
The Czech entomologist A. Vimmer was the
only one who worked on Tachinidae of lepidopterous
hosts at the beginning of the 20th century,
especially soon after Czechoslovakia was established
(see Vimmer 19061938).
Tachinids were intensively studied by forestry
researchers for pest control between World
Wars I and II (Kolubajiv, Komárek, Pfeffer,
Rika). One result of this period was the first
parasitoid catalogue on Lymantria monacha of
Kolubajiv & Pfeffer (1931) summarized also by
Kolubajiv (1937), however with no exact information
on localities. These older findings were
later also used by Kudler (1954). Kolubajiv
(1962) used them again, but he added additional
breeding records.
Attheendofthe1930s,someresearchwasorganized
by foreigners to study parasitoids of European
forest pests that had been introduced into
Canada. Two papers on hosts in their pupal stage,
which were massively exported from central European
countries for purposes of biological control,
were published later (Finlayson & Finlayson
1958a, 1958b). The agriculture research branch,
especially of the Czech sugar industry, searched
also for new ways to combat sugar-beet pests using
biological control methods (Rambousek
1928, 1929). Between 1933 and 1942, D. Jacentkovský
recorded several hosts of tachinids in
Moravia, though his papers were rather focused
on faunistics with a special emphasis on practical
forestry.
From the 1950s on, J. epelák systematically
studied hosts, particularly in cooperation with M.
apek, and some others. Although their papers
were predominantly focused on Slovakia, there
were also some records from the Czech Republic.
Applied forestry research also brought new
knowledge about the hosts of tachinids during
this period (Hochmut, Kalandra, Martinek, UrENTOMOL.
FENNICA Vol. 20  Host catalogue for Tachinidae 23
ban, etc.). Many records are scattered in various
journals (Kleteka, ezá, Samiáková, ámal,
Weiser, Zuska, etc.).
Recently there have been attempts to systematically
search for new hosts of tachinids (e.g.
Mückstein et al. 2004, 2007). Our future projects
will use molecular-genetic methods, which could
perhaps help to explain phylogeny and possible
coevolutionary interactions between Tachinidae
and their hosts.
1.2. Revisionary work
Most revisionary work concerning reared Czech
Tachinidae was done by the late B. Herting, who
studied the old tachinid collections in NHMW
during several visits in the late 1950s. The results
were integrated in his book on the Biologie der
westpaläarktischen Tachinidae from 1960. The
reared material stored in NHMW was  as far as it
could be found  studied again during a visit by
H.-P.Tschorsnigin1988,leadingtoafewcorrections
and additions to Hertings results. The
tachinid collection of the Forestry and Game
Management Research Institute of Strnady near
Prague (Výzkumný ústav lesního hospodáství a
myslivosti) was revised by B. Herting between
1995 and 1997, and his unpublished list could be
used for the present publication. The large collection
of Wachtl was completely revised by H.-P.
Tschorsnig and B. Herting during 1996 and 2002
(see Tschorsnig & Herting 2005). Hence, it is
clear that B. Herting  although deceased in 2004
 must be a coauthor of the present paper. Recent
material sent by several Czech dipterists (M.
Barták, J. Vahara, P. Mückstein etc.) was revised
or identified by H.-P. Tschorsnig during the
last fifteen years, and material (partly of the collection
epelák) was also revised or identified by
J. Vahara.
2. Explanation of the layout of the lists
Annotated parasitoid-host list (Section 3)
 Leftcolumn:Thearrangementofsubfamilies,
tribes and species of the tachinids mainly follows
Herting & Dely-Draskovits (1993), but
updated nomenclature to this relevant catalogue
is listed below. Names used in the original
papers are always cited in parentheses
when they were different (synonyms, different
combinations, emendations, misspellings
or misidentifications), with an explanatory
comment if necessary.
 Right column: Hosts are arranged for each
tachinid in the systematic order of the Hostparasitoid
list. The host names which were
used in the original papers are cited after -as
when they were different, followed by an explanatory
comment if necessary. Wrong or
presumably wrong hosts are set in square
brackets [ ].
Host-parasitoid list (Section 4)
 Left column: Host orders are arranged as follows:
Lepidoptera, Coleoptera, Hymenoptera,
Dermaptera. The arrangement and nomenclature
of Lepidoptera follows Latvka &
Lika (2005), the one for Coleoptera corresponds
with Jelínek (1993), and the arrangement
of the hymenopteran families, genera
and species is alphabetical, their valid scientific
names according to http://www.
faunaeur.org.
 Right column: The tachinids are arranged in
order of their importance for the host, the
most important or specific parasitoids are
mentioned first, doubtful or very uncommon
hosts have an asterisk *, wrong or presumably
wrong hosts are cited in square brackets
[ ].
Host record bibliography (Section 5)
 Only original records are cited, but in the case
of revisions or later confirmatory studies of
the same material, these papers are also mentioned.
Simple citations or replications in
some subsequent listings are cited only when
they are of special interest. Citations of several
workers fromBaer (1921) are omitted because
Baers records are not of Czech origin.
The reference numbers of the Host record
bibliography are used only in the Annotated
parasitoid-host list.
24 Vahara et al.  ENTOMOL. FENNICA Vol. 20
Abbreviations
CZ Czech Republic
HE Herting
ident. identification
misid. misidentified/misidentification
new rec. new record(s) of hitherto unpublished
material
prb. probable/probably
rev. revised/revision (= redetermined by a
specialist)
s.a. see also (= references based on the
same material, but cited because of
some interest)
TS Tschorsnig
VA Vahara
Acronyms of Depositories
NHMW Naturhistorisches Museum Wien,
Austria
VULH Výzkumný ústav lesního
hospodáství a myslivosti (Strnady)
Recent tachinid nomenclature [nomenclature of
Herting and Dely-Draskovits (1993) in square
brackets]:
 Actia resinellae Schrank [syn. nudibasis
Stein]  see Andersen (1996)
 Allophorocera [syn. Erycilla]  see
Tschorsnig & Richter (1998)
 Cleonice keteli Ziegler  not yet in Herting &
Dely-Draskovits, see Ziegler (2000)
 Hubneria [Huebneria]  valid spelling according
to ICZN Art. 33.3.
 Phenicellia [included in Thelaira]  see
Tschorsnig & Herting (1994)
 Phytomyptera [syn. Elfia]  see Andersen
(1988)
 Pseudopachystylum goniaeoides [gonioides]
validspellingaccordingtoICZNArt.33.3.
 Smidtia [syn. Timavia]  see Shima (1996)
ENTOMOL. FENNICA Vol. 20  Host catalogue for Tachinidae 25
3. Annotated parasitoid-host list
Subfamily Exoristinae
Exoristini
Exorista fasciata (Fallén, 1820) -Ennomos erosarius Denis & Schiff. (Lep. Geometridae) [6]
(misid. as Eutachina larvarum-see
2; misid. as Exorista larvarum-18) -Catocala fraxini Linnaeus (Lep. Noctuidae) [2-s.a. 18-rev. TS]
(misid. as Eutachina larvarum-
2; misid. as Exorista larvarum-18) -Calliteara fascelina Linnaeus (Lep. Lymantriidae) [2-s.a. 18-as Dasychira-rev.TS;
according to a note of HE labelled Ch =Chodov, i.e. material of R. v. Stein]
Exorista larvarum (Linnaeus, 1758) (as
Eutachina-100; Tachina-105) -Zygaena ephialtes Linnaeus (Lep. Zygaenidae) [100-s.a. 105]
(as Tachina) -Zygaena punctum Ochsenheimer (Lep. Zygaenidae) [22]
(misid. as Microtachina erucarum Rdi.) -Zygaena sp. (Lep. Zygaenidae) [2-rev. TS]
-Malacosoma neustria Linnaeus (Lep. Lasiocampidae) [8]
-Euthrix potatoria Linnaeus (Lep. Lasiocampidae) [63]
-Cosmotriche lobulina Denis & Schiff. (Lep. Lasiocampidae) [87]
(as Tachina) -Macroglossum stellatarum Linnaeus (Lep. Sphingidae) [105-as Macroglossa]
(as Eutachina-2) -Nymphalis antiopa Linnaeus (Lep. Nymphalidae) [2-as Vanessa-rev. 18; cited as
R. v. Stein in 18, but this is neither indicated on the label nor in 2]
(as Tachina) -Argynnis paphia Linnaeus (Lep. Nymphalidae) [105]
(as Tachina) -Simyra albovenosa Goeze (Lep. Noctuidae) [106-as Arsilonche]
(as Tachina) -Catocala fraxini Linnaeus (Lep. Noctuidae) [105]
(as Tachina; possibly misid. =Voria
ruralis) -Autographa gamma Linnaeus (Lep. Noctuidae) [69 as Plusia]
(as Eutachina-2) -Autographa jota Linnaeus (Lep. Noctuidae) [2-as Plusia-rev. 18]
(as Tachina) -Xylena exsoleta Linnaeus (Lep. Noctuidae) [68-as Calocampa exoleta]
(as Tachina) -Hadula trifolii Hufnagel (Lep. Noctuidae) [68-as Mamestra trifolii Rott.]
-Ceramica pisi Linnaeus (Lep. Noctuidae) [63-as Melanchra]
(as Tachina) -Agrotis segetum Denis & Schiff. (Lep. Noctuidae) [68]
-Euproctis chrysorrhoea Linnaeus (Lep. Lymantriidae) [10-as phaeorrhoea, 39]
(as Eutachina-2) -Leucoma salicis Linnaeus (Lep. Lymantriidae) [2-rev 18-as Stilpnotia 4-as
Stilpnotia]
(as Tachina) -Lymantria monacha Linnaeus (Lep. Lymantriidae) [37-rev. HE, 73-s.a. 74, 105]
(as Eutachina vidua-100; Tachina
vidua-105 ) -Phragmatobia fuliginosa Linnaeus (Lep. Arctiidae) [100-as Spilosoma-s.a. 105]
(as Tachina) -Arctia caja Linnaeus (Lep. Arctiidae) [68, 105]
(as Eutachina l. and E. vidua-100, both
with the same rearing data; Tachina l.and
T. vidua-105; possibly misid. E. fasciata) -Arctia festiva Hufnagel (Lep. Arctiidae) [100-as hebe-s.a. 105]
Exorista grandis (Zetterstedt, 1844) (as
Parasetigena segregata; misid. =prb.
E. grandis) -Saturnia pavonia Linnaeus (Lep. Saturniidae) [101-as carpini]
(as Tricholyga sorbillans) -Hyles euphorbiae Linnaeus (Lep. Sphingidae) [39-as Celerio]
(as sorbillans Wied.) -Deilephila elpenor Linnaeus (Lep. Sphingidae) [6]
(as Tricholyga sorbillans Wied. =prb.
E. grandis or misid.) -Lymantria monacha Linnaeus (Lep. Lymantriidae) [37]
Exorista deligata Pandellé, 1896 (as
Tricholyga abberans Strb.; prb. misid.,
species not known from CZ) -[Saturnia pavonia Linnaeus (Lep. Saturniidae)] [112]
Exorista mimula (Meigen, 1824) (as
Chaetotachina nigricans-100; =misid.,
identity unknown) -[Leucoma salicis Linnaeus (Lep. Lymantriidae)] [100]
(as Microtachina erucarum Rond.; prb.
misid.) -[Diprion pini Linnaeus (Hym. Diprionidae)] [39]
(as Microtachina erucarum Rdi.-2;
erucarum Rond.-18) -Cladius pectinicornis Geoffroy (Hym. Tenthredinidae) [2-as comari-rev. 18]
(misid. as Chaetotachina rustica-2 and
Exorista rustica-18) -Pristiphora pallidiventris Fallén (Hym. Tenthredinidae) [2-as Nematus flavicornis
(but flavicomus on label)-rev. TS]
Exorista rustica (Fallén, 1810) (as
Tachina rustica Mg. var.nigricans;
prb. misid.) -[Nymphalis polychloros Linnaeus (Lep. Nymphalidae)] [105-as Vanessa]
-[Cosmia trapezina Linnaeus (Lep. Noctuidae)] [39-as Calymnia; either host or
tachinid wrong]
(as Tachina-16, 75; prb. misid. =prb. E.
larvarum) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [16, 75]
(as Chaetotachina-2; ident. not
confirmed) -Rhogogaster viridis Linnaeus (Hym. Tenthredinidae) [2-as Tenthredo scalaris]
(as Chaetotachina-2) -Tenthredo arcuata Forster (Hym. Tenthredinidae) [2-as Allantus notus-rev. 18]
(as Chaetotachina-2) -Tenthredopsis coquebertii Klug (Hym. Tenthredinidae) [2-rev. 18]
(as Chaetotachina-2) -Tenthredopsis scutellaris Fabricius (Hym. Tenthredinidae) [2-rev. 18-as palmata]
Chetogena media Rondani, 1859 (as
Chaetogena; confirmed by TS) -Calophasia sp. (Lep. Noctuidae) [5]
Diplostichus janitrix (Hartig, 1838) -Diprion pini Linnaeus (Hym. Diprionidae) [2-as Lophyrus-rev. 18-as Diprion 14,
42, 87]
-Diprion similis Hartig (Hym. Diprionidae) [2-as Lophyrus-rev. 18]
-Gilpinia frutetorum Fabricius (Hym. Diprionidae) [2-as Lophyrus-rev. 18-as
Diprion]
-Gilpinia hercyniae Hartig (Hym. Diprionidae) [2-as Lophyrus]
-Gilpinia polytoma Hartig (Hym. Diprionidae) [15-as Diprion polytomum]
-Gilpinia variegata Hartig (Hym. Diprionidae) [2-as Lophyrus variegatus-rev. 18-as
Diprion]
-Gilpinia virens Klug (Hym. Diprionidae) [2-as Lophyrus-rev. 18-as Diprion]
Parasetigena silvestris (RobineauDesvoidy,
1863) (misid. as Masicera
silvatica-16-see figures, 17; Parasetigena
segregata or misid. Phorocera assimilis-37;
Tachinidae sp.-44- identified in 45;
segregata-27, 40, 45, 46, 47, 73, 74, -Lymantria monacha Linnaeus (Lep. Lymantriidae) [16, 17, 27-s.a. 30, 32-as
75, 76, 86, 104, 105) Liparis, 37-rev. HE 40, 44, 45, 46, 47, 73, 74, 75, 76, 86, 104-s.a. 105, new rec. -
26 Vahara et al.  ENTOMOL. FENNICA Vol. 20
CZ, Doksy Krupá + Halamky, leg. Kolubajiv?, 1948, VULH-rev. HE]
Phorocera assimilis (Fallén, 1810) -Malacosoma neustria Linnaeus (Lep. Lasiocampidae) [105]
-[Aglais urticae Linnaeus (Lep. Nymphalidae)] [105-as Vanessa; questionable host]
-[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [105; either host wrong or
tachinid misid. possibly Parasetigena silvestris]
(as Setigera -100; prb. misid., identity
unknown) -[Arctia festiva Hufnagel (Lep. Arctiidae)] [100  as hebe-s.a. 105]
Phorocera obscura (Fallén, 1810) (as
caesifrons) -[Malacosoma neustria Linnaeus (Lep. Lasiocampidae)] [105; questionable host]
(as Chaetogena vernalis-30, 32) -Operophtera brumata Linnaeus (Lep. Geometridae) [30-s.a. 32-as Cheimatobia,
61]
(as Chaetogena vernalis) -Operophtera fagata Scharfenberg (Lep. Geometridae) [30-as Cheimatobia boreata]
(as caesifrons) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [74, 105; prb. either host
or tachinid ident. wrong]
(as Setigera caesifrons-100; caesifrons-
105; prb. misid., identity unknown) -[Arctia festiva Hufnagel (Lep. Arctiidae)] [100-as hebe-s.a. 105]
Bessa parallela (Meigen, 1824) -Yponomeuta evonymella Linnaeus (Lep. Yponomeutidae) [87, new rec.-CZ, atec
(=Saaz), NHMW-rev. TS]
(as selecta; prb. misid. =prb. parallela) -Yponomeuta padella Linnaeus (Lep. Yponomeutidae) [30-as Hyponemeuta]
(misid. as selecta-39) -Tortrix viridana Linnaeus (Lep. Tortricidae) [39-rev. HE, new rec.-CZ, árka,
leg. Kolubajiv?, 1944, VULH-rev. HE]
(as Ptychomyia) -Acleris ferrugana Denis & Schiff. (Lep. Tortricidae) [83-as Acalla ferrugana Tr.]
Bessa selecta (Meigen, 1824) -Gilpinia polytoma Hartig (Hym. Diprionidae) [15-as Diprion polytomum-rev. HE
(1 sp.) in VULH]
(as Ptychomyia) -Cladius pallipes Serville (Hym. Tenthredinidae) [2-as Priophorus albipes-rev. 18as
Priophorus padi]
(misid. as Prosopodes fugax) -Hemichroa crocea Geoffroy (Hym. Tenthredinidae) [2-as rufa-rev. 18]
-Nematus melanaspis Hartig (Hym. Tenthredinidae) [88]
(as Ptychomyia) -Pristiphora erichsonii Hartig (Hym. Tenthredinidae) [80-as Nematus]
Blondeliini
Belida angelicae (Meigen, 1824) (as
Ceromasia-26; Aporotachina-30) -Arge sp. (Hym. Argidae) [26-as Arge berberidis Schrk.-s.a. 30-as Arge rosea; the
records have the same data, but the host is cited under different species names so the
exact identity remains unclear]
Meigenia incana (Fallén, 1810) (prb.
misid. =prb. Meigenia uncinata) -[Agelastica alni Linnaeus (Col. Chrysomelidae)] [92]
Meigenia mutabilis (Fallén, 1810) -Gastrophysa viridula De Geer (Col. Chrysomelidae) [63]
(as bisignata Meig.; might be misid.
because the typical parasitoid of
Agelastica alni is Meigenia uncinata) -Agelastica alni Linnaeus (Col. Chrysomelidae) [79]
Zaira cinerea (Fallén, 1810) (as Viviana) -Zabrus tenebrioides Goeze (Col. Carabidae) [77-only larvae]
Medina collaris (Fallén, 1820) -Galerucella lineola Fabricius (Col. Chrysomelidae) [96]
-Lochmaea capreae Linnaeus (Col. Chrysomelidae) [new rec.-CZ, Polnika. 1996
and Bílovice n.Sv., 2006, leg. Urban, det. epelák, VA]
Medina luctuosa (Meigen, 1824)
(prb. misid.) -[Plagiodera versicolora Laicharting (Col. Chrysomelidae)] [94; the true Medina
luctuosa is a parasitoid of Altica spp., whereas Plagiodera is a typical host of
Medina separata]
(prb. misid.) -[Linaeidea aenea Linnaeus (Col. Chrysomelidae)] [93]
(prb. misid.; erroneously given as
Meigenia mutabilis in the English -[Gonioctena quinquepunctata Fabricius (Col. Chrysomelidae)] [90; confirmed
summary) hosts of M. luctuosa are Altica spp.]
Medina melania (Meigen, 1824) (prb. -[Plagiodera versicolora Laicharting (Col. Chrysomelidae)] [94; hosts of Medina
misid.) melania are not yet confirmed, but Plagiodera is a typical host of Medina separata]
Staurochaeta albocingulata (Fallén,
1820) (as gracilis Egg.-2) -Monoctenus juniperi Linnaeus (Hym. Diprionidae) [2-as Lophyrus-rev. 18]
Leiophora innoxia (Meigen, 1824)
(as Hypostena-79; Arrhinomyia-11) -[Agelastica alni Linnaeus (Col. Chrysomelidae)] [79-s.a. 11; either host wrong or
tachinid misid. because L. innoxia is a parasitoid of Tetrigidae (Orthoptera)]
ENTOMOL. FENNICA Vol. 20  Host catalogue for Tachinidae 27
Admontia grandicornis (Zetterstedt,
1849) (misid. as Trichoparia seria Mg.) -[Epinotia tedella Clerck (Lep. Tortricidae)] [39-rev. HE; from soil samples-host
must be wrong because A. grandicornis is a parasitoid of Tipulidae]
Blondelia inclusa (Hartig, 1838)
(as Ceromasia-42) -Diprion pini Linnaeus (Hym. Diprionidae) [42]
(as Lophyromyia-39) -Gilpinia polytoma Hartig (Hym. Diprionidae) [15-as Diprion polytomum 39-rev.
HE]
-Neodiprion sertifer Geoffroy (Hym. Diprionidae) [57, 67, 98]
Blondelia nigripes (Fallén, 1810) -Yponomeuta evonymella Linnaeus (Lep. Yponomeutidae) [39-as Hyponomeuta,
new rec.-CZ, leg. Kudler, 1956, VULH-rev. HE]
(as Lydella) -Yponomeuta padella Linnaeus (Lep. Yponomeutidae) [105]
-[Paranthrene tabaniformis Rottemburg (Lep. Sesiidae)] [39; host must be wrong]
(as Ceromasia) -Cochylimorpha hilarana Herrich-Schäffer (Lep. Tortricidae) [110-as Conchylis]
-Tortrix viridana Linnaeus (Lep. Tortricidae) [39-rev. HE]
(as Lydella-37; according to an
unpubl. note of HE misid. by Fahringer
as Phorocera assimilis -81, 82 ) -Choristoneura murinana Hübner (Lep. Tortricidae) [37-as Cacoecia-rev. HE, 81s.a.
82  as Cacoecia]
-Pandemis cerasana Hübner (Lep. Tortricidae) [39-as ribeana-rev. HE]
(as Lydella) -Zeiraphera griseana Hübner (Lep. Tortricidae) [37-as Semasia diniana-rev. HE
65-as Enarmonia diniana-s.a. 66]
-Malacosoma neustria Linnaeus (Lep. Lasiocampidae) [8]
(as Lydella-41) -Dendrolimus pini Linnaeus (Lep. Lasiocampidae) [41-s.a. 39]
(as Ceromasia) -Inachis io Linnaeus (Lep. Nymphalidae) [28-as Vanessa]
-Aglais urticae Linnaeus (Lep. Nymphalidae) [25]
-Ennomos autumnarius Werneburg (Lep. Geometridae) [4]
-Operophtera brumata Linnaeus (Lep. Geometridae) [61]
-Xylena exsoleta Linnaeus (Lep. Noctuidae) [87]
(as Lydella) -Ceramica pisi Linnaeus (Lep. Noctuidae) [105-as Mamestra]
-Arctia caja Linnaeus (Lep. Arctiidae) [4]
-Gilpinia polytoma Hartig (Hym. Diprionidae) [15-as Diprion polytomum]
Blondelia piniariae (Hartig, 1838)
(as Lydella nigripes-37, 82-prb. misid. =
prb. piniariae; Tachina-58) -Bupalus piniarius Linnaeus (Lep. Geometridae) [37, 58-rev. 87, 82]
Compsilura concinnata (Meigen, 1824)
(as Chaetogena rufipalpis-105; s. note
under Acronicta aceris) -Malacosoma neustria Linnaeus (Lep. Lasiocampidae) [39-rev. HE 105]
(as Phorocera) -Bombyx mori Linnaeus (Lep. Bombycidae) [13-as Seidenspinner]
(as Phorocera rufipalpis Mg.-100;
Chaetogena rufipalpis-105; s.a. note
under Acronicta aceris) -Sphinx ligustri Linnaeus (Lep. Sphingidae) [100-s.a. 105]
(as Machaira serriventris Rd.-101) -Iphiclides podalirius Linnaeus (Lep. Papilionidae) [101-as Papilio-s.a. 104, 105]
-Pieris brassicae Linnaeus (Lep. Pieridae) [37-rev. HE 39-rev. HE 72, 105]
-Pieris rapae Linnaeus (Lep. Pieridae) [105]
-Pieris napi Linnaeus (Lep. Pieridae) [105]
-Nymphalis antiopa Linnaeus (Lep. Nymphalidae) [105-as Vanessa]
(as Machaira serriventris Rd.) -Inachis io Linnaeus (Lep. Nymphalidae) [2-as Vanessa jo]
(as Chaetogena rufipalpis; s. note
under Acronicta aceris) -Aglais urticae Linnaeus (Lep. Nymphalidae) [105-as Vanessa]
-Vanessa atalanta Linnaeus (Lep. Nymphalidae) [105-as Pyrameis]
(as Machaira serriventris Rd.) -Araschnia levana Linnaeus (Lep. Nymphalidae) [2-as Vanessa-rev. 18; R. v.
Stein on label according to HE, 4]
-Phalera bucephala Linnaeus (Lep. Notodontidae) [72]
-Acronicta psi Linnaeus (Lep. Noctuidae) [72]
(as Phorocera rufipalpis Mg.-100;
Chaetogena rufipalpis-105; =nomen
dubium, but according to unpubl. note
of HE prb. C. concinnata) -Acronicta aceris Linnaeus (Lep. Noctuidae) [100-as Acronycta-s.a. 105]
(as Machaira serriventris Rd-2) -Lacanobia oleracea Linnaeus (Lep. Noctuidae) [2-as Mamestra-rev. 18]
-Calliteara pudibunda Linnaeus (Lep. Lymantriidae) [87]
28 Vahara et al.  ENTOMOL. FENNICA Vol. 20
-Euproctis chrysorrhoea Linnaeus (Lep. Lymantriidae) [39-rev. HE]
(as Compsilura sp.-105) -Lymantria monacha Linnaeus (Lep. Lymantriidae) [37, 105]
(as Machaira serriventris Rd.-2) -Cimbex sp. (on Salix caprea) (Hym. Cimbicidae) [2-as Cimbex axillaris-rev. 18]
(as Machaira serriventris Rd-2) -Trichiocampus grandis Serville (Hym. Tenthredinidae) [2-as eucera Klg.-rev. 18-as
viminalis Fall.]
Acemyini
Acemya acuticornis (Meigen, 1824) -[Tortrix viridana Linnaeus (Lep. Tortricidae)] [39; either tachinid or host wrong]
-[Rhyacionia buoliana Denis & Schiff. (Lep. Tortricidae)] [39-as Evetria; either
tachinid or host wrong because A. acuticornis is a parasitoid of Orthoptera]
Winthemiini
Winthemia cruentata (Rondani, 1859)
(as ligustri Stein) -Sphinx ligustri Linnaeus (Lep. Sphingidae) [6, 31, 39]
(as ligustri Stein-6; ident. not confirmed,
might also belong to W. rufiventris Macq.) -Mimas tiliae Linnaeus (Lep. Sphingidae) [6]
Winthemia quadripustulata (Fabricius,
1794) -Deilephila elpenor Linnaeus (Lep. Sphingidae) [105-as Chaerocampa]
-Zerynthia polyxena Denis & Schiff. (Lep. Papilionidae) [105-as Thais]
-Inachis io Linnaeus (Lep. Nymphalidae) [39]
-Aglais urticae Linnaeus (Lep. Nymphalidae) [105-as Vanessa]
(as Chaetolyga-2) -Argynnis aglaja Linnaeus (Lep. Nymphalidae) [2-rev. 18, 105]
-Cucullia lactucae Denis & Schiff. (Lep. Noctuidae) [63]
(as Chaetolyga analis Mcq.-2) -Cucullia umbratica Linnaeus (Lep. Noctuidae) [2-rev. 18]
-Shargacucullia scrophulariae Denis & Schiff. (Lep. Noctuidae) [105-as Cucullia]
(as Chaetolyga erythrura-100;
quadripustulata var. erythrura-105; prb.
misid., W. erythrura not known from CZ) -Shargacucullia verbasci Linnaeus (Lep. Noctuidae) [100-as Cucullia-s.a. 105]
(as Chaetolyga-2) -Ceramica pisi Linnaeus (Lep. Noctuidae) [2-as Mamestra-rev. 18]
-Cerapteryx graminis Linnaeus (Lep. Noctuidae) [84-as Charaeas]
Winthemia rufiventris (Macquart, 1849)
(as xanthogastra; prb. misid., species not -[Dioryctria abietella Denis & Schiff. (Lep. Pyralidae)] [112; host impossible for a
known from CZ) large tachinid as Winthemia]
Nemorilla floralis (Fallén, 1810) -Tortrix viridana Linnaeus (Lep. Tortricidae) [39-rev. HE]
Nemorilla maculosa (Meigen, 1824) -Choristoneura murinana Hübner (Lep. Tortricidae) [37-rev. HE]
(prb. correct) -Zeiraphera griseana Hübner (Lep. Tortricidae) [37-as Semasia diniana, 65-as
Enarmonia dinianas.a. 66]
-Trachycera suavella Zincken (Lep. Pyralidae) [87]
Smidtia amoena (Meigen, 1824)
(as Winthemia) -Panolis flammea Denis & Schiff. (Lep. Noctuidae) [105-as griseovariegata Goeze]
Eryciini
Aplomya confinis (Fallén, 1820)
(as Exorista) -Aporia crataegi Linnaeus (Lep. Pieridae) [105-questionable host]
-Neozephyrus quercus Linnaeus (Lep. Lycaenidae) [4-as Zephyrus]
Phebellia clavellariae (Brauer &
Bergenstamm, 1891) (as Parexorista-1) -Pseudoclavellaria amerinae Linnaeus (Hym. Cimbicidae) [1-rev. 19; the material
ex Cimbex ariabilis is not from R.v.Stein, as it is erroneously indicated in 18,
corrected in 19]
Phebellia glauca (Meigen, 1824)
(misid. as Parexorista glirina Rdi.-2) -Acronicta tridens Denis & Schiff. (Lep. Noctuidae) [2-as Acronycta-rev. 18]
Phebellia glirina (Rondani, 1859)
(misid. as Parexorista grossa-2) -Abia sericea Linnaeus (Hym. Cimbicidae) [2-rev. 18]
Phebellia triseta (Pandellé, 1896)
(prb. misid., identity unknown) -[Choristoneura murinana Hübner (Lep. Tortricidae)] [37-as Cacoecia]
-Griposia aprilina Linnaeus (Lep. Noctuidae) [6-as Dichonia]
Nilea hortulana (Meigen, 1824)
(misid. as Parexorista polychaeta
Mcq.-2) -Acronicta tridens Denis & Schiff. (Lep. Noctuidae) [2-as Acronycta-rev. 18]
(as Exorista; prb. misid. = prb.
ENTOMOL. FENNICA Vol. 20  Host catalogue for Tachinidae 29
Myxexoristops sp.) -[Cephalcia abietis Linnaeus (Hym. Pamphiliidae)] [34]
Buquetia musca Robineau-Desvoidy,
1847 (as Eupogona setifacies-39;
Epicampocera setifacies-105) -Papilio machaon Linnaeus (Lep. Papilionidae) [39, 105]
Phryxe erythrostoma (Hartig, 1838) (as
Tachina-35; misid. as vulgaris-39) -Sphinx pinastri Linnaeus (Lep. Sphingidae) [35, 39-as Hyloicus rev. HE, new rec.CZ,
eský Rudolec, leg. Kolubajiv?, 1934 and Psovlky nr. Rakovník, leg. Martinek,
VULH-rev. HE]
Phryxe magnicornis (Zetterstedt, 1838) -Zygaena laeta Hübner (Lep. Zygaenidae) [23]
(misid. as vulgaris) -Tortrix viridana Linnaeus (Lep. Tortricidae) [39-rev. HE]
(misid. as vulgaris) -Erannis defoliaria Clerck (Lep. Geometridae) [37-as Hibernia-rev. HE]
Phryxe nemea (Meigen, 1824) -Choristoneura murinana Hübner (Lep. Tortricidae) [18-as Cacoecia 81-s.a. 82-rev.]
-Pieris brassicae Linnaeus (Lep. Pieridae) [18]
-Neozephyrus quercus Linnaeus (Lep. Lycaenidae) [4-as Zephyrus]
(as Blepharidopsis) -Abraxas grossulariata Linnaeus (Lep. Geometridae) [2-rev. 18]
Phryxe prima (Brauer & Bergenstamm,
1889) -Zygaena carniolica Scopoli (Lep. Zygaenidae) [63]
(as Ceratochaetops) -Zygaena punctum Ochsenheimer (Lep. Zygaenidae) [22]
Phryxe semicaudata Herting, 1959 -Thaumetopoea processionea Linnaeus (Lep. Thaumetopoeidae) [87]
Phryxe setifacies (Villeneuve, 1910)
(misid. as Blepharidopsis nemea) -Zygaena angelicae Ochsenheimer (Lep. Zygaenidae) [6-rev. VA; first record of
Phryxe setifacies for CZ]
Phryxe vulgaris (Fallén, 1810) (possibly
misid.) -Yponomeuta padella Linnaeus (Lep. Yponomeutidae) [105]
-Malacosoma neustria Linnaeus (Lep. Lasiocampidae) [8]
(=vulgaris or misid.) -Lasiocampa trifolii Denis & Schiff. (Lep. Lasiocampidae) [105-s.a. 108]
-Deilephila porcellus Linnaeus (Lep. Sphingidae) [105-as Metopsilus]
-Aporia crataegi Linnaeus (Lep. Pieridae) [39]
(as Blepharidea-2) -Pieris brassicae Linnaeus (Lep. Pieridae) [2-rev. 18, 105]
-Pieris napi Linnaeus (Lep. Pieridae) [105]
-Anthocharis cardamines Linnaeus (Lep. Pieridae) [105-as Euchloë]
-Satyrium pruni Linnaeus (Lep. Lycaenidae) [4-as Thecla]
-Nymphalis polychloros Linnaeus (Lep. Nymphalidae) [37-as Vanessa-rev. HE]
-Nymphalis xanthomelas Denis & Schiff. (Lep. Nymphalidae) [105-as Vanessa]
-Nymphalis antiopa Linnaeus (Lep. Nymphalidae) [18-as Vanessa]
(as Exorista-100) -Inachis io Linnaeus (Lep. Nymphalidae) [100-as Vanessa-s.a. 105, 108]
(as Exorista-100, Blepharidea-102) -Aglais urticae Linnaeus (Lep. Nymphalidae) [4-as Vanessa, 100-as Vanessa-s.a.
102, 104, 105, 108; new rec.-CZ, atec (=Saaz), NHMW-rev. TS]
-Araschnia levana Linnaeus (Lep. Nymphalidae) [18]
(as Blepharidea-2) -Issoria lathonia Linnaeus (Lep. Nymphalidae) [2-as Argynnis-rev. 18]
-[Bupalus piniarius Linnaeus (Lep. Geometridae)] [82; questionable host]
-Autographa gamma Linnaeus (Lep. Noctuidae) [87-as Plusia]
(as Blepharidea-2) -Tholera cespitis Denis & Schiff. (Lep. Noctuidae) [2-as Neuronia-rev. 18-as
Epineuronia]
-Agrotis segetum Denis & Schiff. (Lep. Noctuidae) [70-s.a. 105, 108]
-Orgyia antiqua Linnaeus (Lep. Lymantriidae) [105]
(prb. misid. =prb. Hubneria affinis) -[Parasemia plantaginis Linnaeus (Lep. Arctiidae)] [105]
(prb. misid. =prb. Hubneria affinis) -[Arctia caja Linnaeus (Lep. Arctiidae)] [108]
Periarchiclops scutellaris (Fallén, 1820)
(as Prosopaea abbreviata-2) -Conistra rubiginea Denis & Schiff. (Lep. Noctuidae) [2-as Acronycta rubiginosarev.
18-as Orrhodia]
Bactromyia aurulenta (Meigen, 1824) -[Pristiphora abietina Christ (Hym. Tenthredinidae)] [39; either parasitoid or host
must be wrong]
Pseudoperichaeta nigrolineata (Walker,
1853) (misid. as Exorista mitis in part-39) -Tortrix viridana Linnaeus (Lep. Tortricidae) [39-rev. HE, new rec.-CZ, Moravia,
Tvrdonice, leg. F. Gregor, 1954, VULH-rev. HE]
(as insidiosa R.-D.-4, Exorista roseanae
B.& B.-20) -Archips crataegana Hübner (Lep. Tortricidae) [4, 20-as Cacoecia]
(as P. major) -[Malacosoma neustria Linnaeus (Lep. Lasiocampidae)] [39; host prb. wrong]
(as P. major) -[Euproctis chrysorrhoea Linnaeus (Lep. Lymantriidae)] [39; host prb. wrong]
30 Vahara et al.  ENTOMOL. FENNICA Vol. 20
Lydella stabulans (Meigen, 1824) -Hydraecia micacea Esper (Lep. Noctuidae) [18-specimen in NHMW labelled
Saaz (=atec), according to an unpublished note of HE 63]
Lydella thompsoni Herting, 1959 -Ostrinia nubilalis Hübner (Lep. Pyralidae) [3]
Cadurciella tritaeniata (Rondani, 1859)
(as Parexorista-2, as Chaetina-18) -Callophrys rubi Linnaeus (Lep. Lycaenidae) [2-as Thecla-rev. 18]
Drino bohemica Mesnil, 1949 (as Sturmia
inconspicua-rev. HE, but ident. uncertain -Gilpinia hercyniae Hartig (Hym. Diprionidae) [37-as Lophyrus hercyniae Hrtg.
because of the bad shape of the material) (polytomus Hrtg.)]
-Neodiprion sertifer Geoffroy (Hym. Diprionidae) [48, 51]
Drino gilva (Hartig, 1838) (as
Argyrophylax gilva Htg.-2) -Gilpinia pallida Klug (Hym. Diprionidae) [2-as Lophyrus-rev. 18-as Diprion]
-Neodiprion sertifer Geoffroy (Hym. Diprionidae) [48, 51, 57, 67]
Drino inconspicua (Meigen, 1830) (as
Argyrophylax-39; Sturmia bimaculata
Htg.-41) -Dendrolimus pini Linnaeus (Lep. Lasiocampidae) [41-rev. HE-s.a. 39]
(as Argyrophylax-39; Sturmia bimaculata
Hart.-73, 74) -Lymantria monacha Linnaeus (Lep. Lymantriidae) [39-rev. HE 73-s.a. 74]
(misid. as Hemimasicera vicina-39) -Diprion pini Linnaeus (Hym. Diprionidae) [39-rev. HE 42, 87, new rec.-CZ,
Zbiroh, leg. I. Hájek, 1970, VULH-rev. HE]
(as Argyrophylax bimaculata Htg.-2) -Gilpinia pallida Klug (Hym. Diprionidae) [2-as Lophyrus-rev. 18-as Diprion]
(as Sturmia-59) -Gilpinia polytoma Hartig (Hym. Diprionidae) [15, 59-as Diprion polytomum]
(as Argyrophylax bimaculata Htg.-2) -Gilpinia variegata Hartig (Hym. Diprionidae) [2-as Lophyrus-rev. 18-as Diprion]
(as Argyrophylax bimaculata Htg.-2) -Gilpinia virens Klug (Hym. Diprionidae) [2-as Lophyrus-rev. 18-as Diprion]
(as Argyrophylax bimaculata Htg.-2;
Sturmia-37, 38) -Neodiprion sertifer Geoffroy (Hym. Diprionidae) [2-as Lophyrus rufus-rev. 18, 14,
37-s.a. 38-rev. HE 48, 51, 57, 67]
Drino lota (Meigen, 1824) (misid. as
Winthemia quadripustulata-39; misid.
Ernestia radicum-106, the figures clearly
indicate D. lota) -Deilephila elpenor Linnaeus (Lep. Sphingidae) [39-as Pergesa-rev. HE, 106-as
Chaerocampa]
(as Phorcida) -Naenia typica Linnaeus (Lep. Noctuidae) [6]
Hubneria affinis (Fallén, 1810) (as
Parexorista polycheta-100, Exorista
polychaeta and E. affinis-105, Exorista-
108) -[Malacosoma neustria Linnaeus (Lep. Lasiocampidae)] [100-as Bombyx-s.a. 105,
108; prb. misid.]
(as Exorista) -[Saturnia pavonia Linnaeus (Lep. Saturniidae)] [105-as carpini; host prb. wrong]
(as Exorista) -[Calliteara pudibunda Linnaeus (Lep. Lymantriidae)] [100-as Dasychira-s.a. 105,
108; prb. misid.]
(as Exorista) -[Lymantria dispar Linnaeus (Lep. Lymantriidae)] [105; questionable host]
(as Exorista) -Spilosoma luteum Hufnagel (Lep. Arctiidae) [39-as Spilarctia lubricipeda]
(as Exorista) -Spilosoma sp. (Lep. Arctiidae) [37]
(as Parexorista polycheta and Exorista
affinis-100; Exorista affinis and E.
polycheta-105; Exorista-108) -Phragmatobia fuliginosa Linnaeus (Lep. Arctiidae) [100-as Spilosoma-s.a. 105, 108]
(as Parexorista polycheta-100, 104;
Exorista affinis and E. polycheta-105;
Exorista-108) -Arctia caja Linnaeus (Lep. Arctiidae) [4, 39-rev. HE 63, 100-s.a. 104, 105, 108]
(as Parexorista polycheta Macq. and
Exorista affinis -101; Exorista polycheta-
105; Exorista-108) -Arctia villica Linnaeus (Lep. Arctiidae) [101-s.a. 105, 108]
(as Exorista-100; Parexorista-102;
Parexorista polycheta-104; Exorista-
105, 108) -Arctia festiva Hufnagel (Lep. Arctiidae) [100-as hebe-s.a. 102, 104, 105, 108]
Carcelia atricosta Herting, 1961 (misid.
as Sisyropa lucorum-2 and C.
puberula-18) -Orgyia antiqua Linnaeus (Lep. Lymantriidae) [2, 18-rev. 19]
(misid.as Parexorista cheloniae-2 and
C. puberula-18) -[Cimbex femoratus Linnaeus (Hym. Cimbicidae)] [2, 18-both as betulae Zdd.-rev.
ENTOMOL. FENNICA Vol. 20  Host catalogue for Tachinidae 31
19; host unusual for Carcelia and presumably wrong]
Carcelia gnava (Meigen, 1824) (as
Sisyropa excissa-100, S. excissa and
C. gnava-104, excissa and gnava-105) -Malacosoma neustria Linnaeus (Lep. Lasiocampidae) [100as Bombyx, 104as
Malacosoma and Bombyx, 105]
(as excissa; prb. misid.) -[Proserpinus proserpina Pallas (Lep. Sphingidae)] [105-as Pterogon, s.a. 108]
-[Aglais urticae Linnaeus (Lep. Nymphalidae)] [25-tachinid confirmed by TS, but
host unusual for Carcelia and prb. wrong]
(as Parexorista and Sisyropa excavata-
100; excissa and gnava-105, 108;
excissa is a wrong synonymization
of excavata, see 103) -Calliteara pudibunda Linnaeus (Lep. Lymantriidae) [4-as Dasychira, 87, 97, 100as
Dasychira-s.a. 104, 105, 108]
-Leucoma salicis Linnaeus (Lep. Lymantriidae) [new rec.-CZ, Kaplice env., leg.
Kolubajiv?, 1966, VULH-rev. HE]
(as Parexorista-100; prb. misid. =prb.
Carcelia lucorum) -[Arctia caja Linnaeus (Lep. Arctiidae)] [100-s.a. 104, 105, 108]
Carcelia iliaca (Ratzeburg, 1840) -Thaumetopoea processionea Linnaeus (Lep. Thaumetopoeidae) [87]
Carcelia laxifrons Villeneuve, 1912
(misid. as Parexorista cheloniae-2) -Leucoma salicis Linnaeus (Lep. Lymantriidae) [2-rev. 18-as Stilpnotia]
Carcelia rasa (Macquart, 1849) -Calliteara pudibunda Linnaeus (Lep. Lymantriidae) [63, 87]
Carcelia sp. -[Choristoneura murinana Hübner (Lep. Tortricidae)] [81-s.a. 82-as Cacoecia; prb.
wrong because Carcelia does not parasitize Tortricidae]
Senometopia excisa (Fallén, 1820)
(as Carcelia) -[Sphinx pinastri Linnaeus (Lep. Sphingidae)] [39-as Hyloicus; host prb. wrong]
(as Sisyropa excissa -102, 104; Carcelia-
105; might be some species of
Senometopia or Carcelia) -[Pheosia gnoma Fabricius (Lep. Notodontidae)] [102-as Notodonta dictoides Huf.s.a.
104, 105]
(as Carcelia excissa; prb. misid. =prb.
Carcelia puberula) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [108-as Limantria]
(as Carcelia excissa; prb. misid. =prb.
Carcelia bombylans) -[Phragmatobia fuliginosa Linnaeus (Lep. Arctiidae)] [108]
Senometopia pollinosa (Mesnil, 1941) (as
Carcelia obesa Zett.) -[Zygaena filipendulae Linnaeus (Lep. Zygaenidae)] [6; host prb. wrong]
(as Carcelia rutilla Meig.) -[Dendrolimus pini Linnaeus (Lep. Lasiocampidae)] [105; host prb. wrong]
(as Carcelia obesa Zett.-6; misid. as
Nemorea erythrura = prb. S. pollinosa-
17; Carcelia rutilla Meig.-37, 82; Exorista
flavicans-114) -Bupalus piniarius Linnaeus (Lep. Geometridae) [6, 17-as Fidonia piniaria 37, 82,
87, 114-rev.]
Senometopia susurrans (Rondani, 1859)
(as Carcelia; prb. misid., identity
unknown) -[Dioryctria abietella Denis & Schiff. (Lep. Pyralidae)] [111]
Erycia fatua (Meigen, 1824) -Melitaea athalia Rottemburg (Lep. Nymphalidae) [105]
(prb. misid. =prb. Allophorocera
ferruginea which is a parasitoid of
Tipulidae) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [74]
Erycia festinans (Meigen, 1824) (as
Hemimasicera; prb. misid. =prb.
Allophorocera ferruginea which is a
parasitoid of Tipulidae) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [74]
Xylotachina diluta (Meigen, 1824) (as
Lydella ambulans Rd.) -Cossus cossus Linnaeus (Lep. Cossidae) [105]
Townsendiellomyia nidicola (Townsend,
1908) -Euproctis chrysorrhoea Linnaeus (Lep. Lymantriidae) [new rec.-CZ, Damboice,
leg. Kolubajiv?, 1955, VULH-rev. HE]
32 Vahara et al.  ENTOMOL. FENNICA Vol. 20
Goniini
Eumea linearicornis (Zetterstedt, 1844)
(as Exorista westermanni and misid. in
part as Exorista mitis and Blondelia
nigripes) -Tortrix viridana Linnaeus (Lep. Tortricidae) [39-rev. HE]
(misid. as Phryxe vulgaris) -Archips rosana Linnaeus (Lep. Tortricidae) [39-as Cacoecia-rev. HE]
(as Exorista westermanni) -Pandemis cerasana Hübner (Lep. Tortricidae) [39-as ribeana-rev. HE]
(as spernanda Zett) -Cosmia affinis Linnaeus (Lep. Noctuidae) [5]
(as Exorista westermanni and misid. as E.
mitis in part) -Cosmia trapezina Linnaeus (Lep. Noctuidae) [39-as Calymnia-rev. HE]
Eumea mitis (Meigen, 1824) -Megalophanes viciella Denis & Schiff. (Lep. Psychidae) [6-as Psyche]
(as Exorista) -Archips crataegana Hübner (Lep. Tortricidae) [20-as Cacoecia-s.a. 21]
(as Exorista) -[Dendrolimus pini Linnaeus (Lep. Lasiocampidae)] [41-s.a. 39; questionable host]
(as Exorista) -Cosmia trapezina Linnaeus (Lep. Noctuidae) [39-rev. HE]
(as Platymyia) -Cerapteryx graminis Linnaeus (Lep. Noctuidae) [84-as Charaeas]
(misid. as Parexorista temera Mg.-2) -Nematus oligospilus Förster (Hym. Tenthredinidae) [2-rev. 18-as Pteronidea
oligospila]
Myxexoristops abietis Herting, 1964 (as
M. abietis and prb. misid. M. blondeli-49) -Cephalcia abietis Linnaeus (Hym. Pamphiliidae) [49-material reared from
Cephalcia sp., misid. by epelák as M. blondeli, was found by HE in coll. VULH,
52]
-Cephalcia sp. (Hym. Pamphiliidae) [52, 53]
Myxexoristops bicolor (Villeneuve, 1908)
(as M. bicolor, and prb. misid. M.
bonsdorffi and M. blondeli) -Cephalcia abietis Linnaeus (Hym. Pamphiliidae) [49; material reared from
Cephalcia sp., misid. by epelák as M. bonsdorffi and M. blondeli, was found by
HE in coll. VULH]
-Cephalcia alpina Klug (Hym. Pamphiliidae) [55-as falleni]
-Cephalcia arvensis Panzer (Hym. Pamphiliidae) [54, 56]
Myxexoristops bonsdorffi (Zetterstedt,
1859) (misid. as Parexorista cheloniae
Rd.-2) -Acantholyda posticalis Matsumura (Hym. Pamphiliidae) [2-as Lyda stellata-rev.
18-as A. stellata]
Zenillia dolosa (Meigen, 1824) (as
libatrix f. grisella; ident. not confirmed) -Tortrix viridana Linnaeus (Lep. Tortricidae) [39]
(as libatrix f. grisella; ident. not
confirmed) -Euproctis chrysorrhoea Linnaeus (Lep. Lymantriidae) [39]
Zenillia libatrix (Panzer, 1798) -Pandemis cerasana Hübner (Lep. Tortricidae) [39-as ribeana]
-Malacosoma neustria Linnaeus (Lep. Lasiocampidae) [8, 104-s.a 105, 108]
(as Exorista; =Z. libatrix or misid.) -Erannis defoliaria Clerck (Lep. Geometridae) [37-as Hibernia]
(prb. misid. =Senometopia pollinosa) -[Bupalus piniarius Linnaeus (Lep. Geometridae)] [104-s.a. 105]
-Thaumetopoea processionea Linnaeus (Lep. Thaumetopoeidae) [5, 87]
-Euproctis chrysorrhoea Linnaeus (Lep. Lymantriidae) [39-rev HE, 104-s.a. 105,
108]
-Lymantria dispar Linnaeus (Lep. Lymantriidae) [87]
Clemelis pullata (Meigen, 1824) -Pandemis heparana Denis & Schiff. (Lep. Tortricidae) [6]
Pales pavida (Meigen, 1824)
(misid. as Phryxe vulgaris) -Zeiraphera griseana Hübner (Lep. Tortricidae) [37-as Semasia diniana-rev. HE]
-Eriogaster lanestris Linnaeus (Lep. Lasiocampidae) [37-rev. HE]
(as Ctenophorocera-64) -Malacosoma neustria Linnaeus (Lep. Lasiocampidae) [64, 104-s.a. 105, new rec.CZ,
Praha-Hrdloezy, leg. Kolubajiv, 1937, VULH-rev. HE]
(as Nilea-39) -Dendrolimus pini Linnaeus (Lep. Lasiocampidae) [41-s.a. 39, 87]
(prb. misid. of Pales processioneae) -Thaumetopoea processionea Linnaeus (Lep. Thaumetopoeidae) [5]
-Panolis flammea Denis & Schiff. (Lep. Noctuidae) [37-rev. HE]
-Autographa gamma Linnaeus (Lep. Noctuidae) [68-as Plusia-s.a. 69]
(as Nilea-39; Ctenophorocera-116) -Euproctis chrysorrhoea Linnaeus (Lep. Lymantriidae) [4, 39, 116-as phaeorrhoea]
(as Nilea-39) -Leucoma salicis Linnaeus (Lep. Lymantriidae) [26-as Stilpnotia-s.a. 32, 39-rev.
HE, new rec.-CZ, Kaplice, leg. Kolubajiv, 1966, VULH-rev. HE]
-Lymantria monacha Linnaeus (Lep. Lymantriidae) [26-as Liparis-s.a. 27, 32]
(as Phorocera cilipeda-2) -[Allantus truncatus Klug (Hym. Tenthredinidae)] [2-as Emphytus cingillum-rev. 18;
ENTOMOL. FENNICA Vol. 20  Host catalogue for Tachinidae 33
tachinid confirmed by TS, but prb. wrong]
Pales pumicata (Meigen, 1824) (as
Nilea; must be misid., identity unknown) -[Paranthrene tabaniformis Rottemburg (Lep. Sesiidae)] [39]
(as Nilea; must be misid., identity
unknown) -[Saperda carcharias Linnaeus (Col., Cerambycidae)] [39]
(as Nilea; must be misid., identity
unknown) -[Diprion pini Linnaeus (Hym. Diprionidae)] [39]
Phryno vetula (Meigen, 1824) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [74-s.a. 108; questionable
host]
Cyzenis albicans (Fallén, 1810) -Operophtera brumata Linnaeus (Lep. Geometridae) [4, 61]
Bothria frontosa (Meigen, 1824) -[Aglais urticae Linnaeus (Lep. Nymphalidae)] [105-as Vanessa; questionable
because hosts of Bothria are Noctuidae]
Ceromasia rubrifrons (Macquart, 1834)
(as florum Macq.) -Zygaena carniolica Scopoli (Lep. Zygaenidae) [109]
(misid. as Masicera rutila Mg.-100;
Ceromasia (Masicera) ferruginea Mg.=
florum Rd.-104; florum Macq. and Erycia
fatua Mg.-105) -Zygaena sp. (Lep. Zygaenidae) [100-as fausta-s.a. 104, 105; Z. fausta is unknown
from CZ, so it was prb. a misid. of some other Zygaena]
(as florum Macq.; prb. misid. =prb.
Allophorocera ferruginea) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [37, 39, 105; hosts prb. larvae
of Tipulidae in soil samples]
Allophorocera ferruginea (Meigen,
1824) (misid. as Ceromasia rutilla Mg.;
= prb. A. ferruginea which is a parasitoid
of Tipulidae) -[Dendrolimus pini Linnaeus (Lep. Lasiocampidae)] [41-s.a. 39]
(as Ceromasia ferruginea Fall.) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [86; host prb. wrong]
Allophorocera rufipes (Brauer &
Bergenstamm, 1891) (as Tachina
vicinalis Pand.; possibly misid.) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [105; host prb. wrong]
Rhacodinella apicata (Pandellé, 1896) -Phyllodesma ilicifolia Linnaeus (Lep. Lasiocampidae) [63]
Eurysthaea scutellaris (RobineauDesvoidy,
1848) -Tortrix viridana Linnaeus (Lep. Tortricidae) [new rec.-CZ, árka, leg. Kolubajiv?,
1946, VULH-rev. HE]
Elodia morio (Fallén, 1820)(as Elodia
tragica and misid. Degeeria luctuosa) -Tortrix viridana Linnaeus (Lep. Tortricidae) [39-rev. HE]
Sturmia bella (Meigen, 1824) -Nymphalis polychloros Linnaeus (Lep. Nymphalidae) [37-as Vanessa]
-Inachis io Linnaeus (Lep. Nymphalidae) [4-as Vanessa jo 6-as Nymphalis-s.a. 116,
25]
-Aglais urticae Linnaeus (Lep. Nymphalidae) [4-as Vanessa, new rec.-CZ, Moravia,
Herálec, 2003, leg., det. and coll. P. Mückstein]
-Vanessa atalanta Linnaeus (Lep. Nymphalidae) [6, new rec.-CZ, Bohemia or.,
Rovný env., 2003, leg., det. and coll. P. Mückstein]
-[Phalera bucephala Linnaeus (Lep. Notodontidae)] [4; doubtful host]
-[Leucoma salicis Linnaeus (Lep. Lymantriidae)] [26-as Stilpnotia-s.a. 32; doubtful
host]
Blepharipa pratensis (Meigen, 1824) (as
Blepharipoda scutellata-39; Sturmia
scutellata-41) -Dendrolimus pini Linnaeus (Lep. Lasiocampidae) [41-rev. HE-s.a. 39]
(as Sturmia scutellata-104; S. scutellata
and Blepharipoda major-105; S.
scutellata-111; possibly misid. Drino
galii) -Hyles euphorbiae Linnaeus (Lep. Sphingidae) [104-as Deilephila-s.a. 105, 111]
(as Sturmia scutellata; prb. misid. =prb.
Sturmia bella) -[Aglais urticae Linnaeus (Lep. Nymphalidae)] [105-as Vanessa]
(as Sturmia scutellata) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [74, 105; questionable host]
Masicera pavoniae (Robineau-Desvoidy,
1830) (misid. as sylvatica-13; pratensis
Meig.-18, 104, 105; Sturmia scutellata-
107 =prb. misid.) -Saturnia pyri Denis & Schiff. (Lep. Saturniidae) [2-rev. 18; R. v. Stein on label
34 Vahara et al.  ENTOMOL. FENNICA Vol. 20
according to HE, 13, 104-s.a. 105, 107]
(as pratensis) -Saturnia spini Denis & Schiff. (Lep. Saturniidae) [104-s.a. 105]
(as pratensis Mg.) -Saturnia pavonia Linnaeus (Lep. Saturniidae) [99-as carpini 100-s.a. 104]
Masicera silvatica (Fallén, 1810)
(=silvatica or misid.) -Lasiocampa trifolii Denis & Schiff. (Lep. Lasiocampidae) [105]
-Macrothylacia rubi Linnaeus (Lep. Lasiocampidae) [4]
Masicera sphingivora (RobineauDesvoidy,
1830) (as pratensis-100, 104,
105; silvatica-103; prb. misid. = prb.
sphingivora) -Smerinthus ocellatus Linnaeus (Lep. Sphingidae) [100-s.a. 103, 104, 105]
(as pratensis; prb. misid. = prb.
sphingivora) -Laothoe populi Linnaeus (Lep. Sphingidae) [104-as Smerinthus-s.a. 105]
(as silvatia-105; prb. misid. = prb.
sphingivora) -Hyles euphorbiae Linnaeus (Lep. Sphingidae) [98, 105-as Deilephila]
Prosopea nigricans (Egger, 1861) (prb.
misid., identity unknown) -[Ceramica pisi Linnaeus (Lep. Noctuidae)] [105-as Mamestra]
Frontina laeta (Meigen, 1824) -Sphinx ligustri Linnaeus (Lep. Sphingidae) [105]
-Smerinthus ocellatus Linnaeus (Lep. Sphingidae) [105-as ocellata]
Thelymorpha marmorata (Fabricius,
1805) (as vertiginosa Fll.-2, as
Histochaeta-6, 18) -Arctia caja Linnaeus (Lep. Arctiidae) [2-rev. 18, 6]
Baumhaueria goniaeformis (Meigen,
1824) (possibly misid.) -Shargacucullia scrophulariae Denis & Schiff. (Lep. Noctuidae) [105-as Cucullia]
-Euproctis chrysorrhoea Linnaeus (Lep. Lymantriidae) [10-as phaeorrhoea]
Gonia capitata (De Geer, 1776) (as
capitata and divisa-105; divisa Mg.-
68, 70; prb. misid. =prb. capitata) -Agrotis segetum Denis & Schiff. (Lep. Noctuidae) [68-s.a. 70, 105]
Gonia picea (Robineau-Desvoidy, 1830)
(as Salmacia sicula R.-D.) -Cerapteryx graminis Linnaeus (Lep. Noctuidae) [84, 85-s.a. 116-as Chareas]
Spallanzania hebes (Fallén, 1820)
(as Pseudogonia-70; Cnephalia-105) -Agrotis segetum Denis & Schiff. (Lep. Noctuidae) [70-s.a. 105]
Subfamily Tachininae
Tachinini
Tachina grossa (Linnaeus, 1758)
(as Echinomyia) -Lasiocampa trifolii Denis & Schiff. (Lep. Lasiocampidae) [105]
Tachina fera (Linnaeus, 1761)(as
Echinomyia) -Lithomoia solidaginis Hübner (Lep. Noctuidae) [107-as Catocala]
(as Echinomyia-18) -Ceramica pisi Linnaeus (Lep. Noctuidae) [2-as Mamestra-rev. 18]
(as Echinomyia; =T. fera or misid. T.
magnicornis) -Lymantria monacha Linnaeus (Lep. Lymantriidae) [73-s.a. 74]
Peleteria ferina (Zetterstedt, 1844) (as
Tachina-100; Echinomya-105) -Parasemia plantaginis Linnaeus (Lep. Arctiidae) [100-as Nemeophila-s.a. 105]
Peleteria rubescens (Robineau-Desvoidy,
1830) (as Pallatiera prompta Mg.-74 =
misid. P. rubescens; nigricornis-105) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [74, 105; host prb. wrong]
Nemoraeini
Nemoraea pellucida (Meigen, 1824)
(as rubrica Mg.-106; rubica-107) -Colocasia coryli Linnaeus (Lep. Noctuidae) [106-as Demas-s.a. 107]
Linnaemyiini
Linnaemya comta (Fallén, 1810) (as
Micropalpus comptus-101, 102;
compta-104, 105) -[Saturnia pavonia Linnaeus (Lep. Saturniidae)] [101-as carpini-s.a. 102, 104, 105;
doubtful because Linnaemya is not known as a parasitoid of the very commonly
reared genus Saturnia]
Linnaemya impudica (Rondani, 1859)
(as Micropalpus impudicus) -Agrotis sp. (Lep. Noctuidae) [107]
ENTOMOL. FENNICA Vol. 20  Host catalogue for Tachinidae 35
(as Micropalpus impudicus) -[Euproctis chrysorrhoea Linnaeus (Lep. Lymantriidae)] [39; host prb. wrong]
Linnaemya haemorrhoidalis (Fallén,
1810) (as Micropalpus-102) -[Saturnia pavonia Linnaeus (Lep. Saturniidae)] [102-s.a. 104-as pavoniae and
carpini, 105-as carpini; doubtful, see note under Linnaemya comta]
Linnaemya picta (Meigen, 1824)
(as Micropalpus pictus Mg.; = picta or
Linnaemya sp.) -[Hyles euphorbiae Linnaeus (Lep. Sphingidae)] [105-as
Deilephila; questionable because hosts of L. picta are Noctuidae]
Lypha dubia (Fallén, 1810) -Retinia resinella Linnaeus (Lep. Tortricidae) [36]
-Rhyacionia buoliana Denis & Schiff. (Lep. Tortricidae) [50]
-Operophtera brumata Linnaeus (Lep. Geometridae) [61]
Ernestiini
Ernestia rudis (Fallén, 1810) (as
Panzeria strenua Mg.; = E. rudis or
misid.) -[Saturnia pyri Denis & Schiff. (Lep. Saturniidae)] [105; host prb. wrong]
(as Panzeria strenua Mg.; = E. rudis or
misid.) -[Saturnia pavonia Linnaeus (Lep. Saturniidae)] [105-as carpini; host prb. wrong]
-Panolis flammea Denis & Schiff. (Lep. Noctuidae) [37]
-[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [37, 74, 105; ident. in part
confirmed by HE, but host must be wrong because E. rudis failed to parasitize L.
monacha in rearing experiments]
Eurithia anthophila (RobineauDesvoidy,
1830) (as Erigone radicum
Fll.-2; Ernestia radicum F.-18) -Spilosoma lubricipeda Linnaeus (Lep. Arctiidae) [2-as menthastri-rev. 18]
Eurithia connivens (Zetterstedt, 1844)
(as Ernestia) -[Choristoneura murinana Hübner (Lep. Tortricidae)] [81-s.a. 82-as Cacoecia;
tachinid ident. correct (det. Mesnil), but host too small for Eurithia]
Eurithia consobrina (Meigen, 1824) (as
Ernestia; prb. misid., identity unknown) -[Eriogaster lanestris Linnaeus (Lep. Lasiocampidae)] [37]
(as Ernestia) -Cucullia artemisiae Hufnagel (Lep. Noctuidae) [105]
(as Ernestina) -[Lymantria dispar Linnaeus (Lep. Lymantriidae)] [105; host prb. wrong]
Hyalurgus tomostethi epelák, 1963 -Tomostethus nigritus Fabricius (Hym. Tenthredinidae) [9-s.a. 60]
Cleonice callida (Meigen, 1824) -Chrysomela populi Linnaeus (Col. Chrysomelidae) [95]
Cleonice keteli Ziegler, 2000 (misid. as
callida) -Chrysomela vigintipunctata Scopoli (Col. Chrysomelidae) [91-rev. TS]
Brachymerini
Pseudopachystylum goniaeoides
(Zetterstedt, 1838) -Cephalcia arvensis Panzer (Hym. Pamphiliidae) [54]
-Cephalcia sp. (Hym. Pamphiliidae) [49]
Pelatachinini
Pelatachina tibialis (Fallén, 1810) -Nymphalis antiopa Linnaeus (Lep. Nymphalidae) [26-as Vanessa-s.a. 30, 32]
-Inachis io Linnaeus (Lep. Nymphalidae) [26-as Vanessa-s.a. 30, 32]
(as Hyria) -Lacanobia oleracea Linnaeus (Lep. Noctuidae) [2-as Mamestra; untypical host]
Macquartiini
Anthomyiopsis plagioderae Mesnil, 1972
(misid. as A. nigrisquamata) -[Altica quercetorum Foudras (Col. Chrysomelidae)] [89-rev. VA, TS; the host is
unlikely because A. plagioderae is known from Chrysomelidae on Salix, primarily
Plagiodera versicolora; first record of A. plagioderae for CZ]
Triarthriini
Triarthria setipennis (Fallén, 1810) -Forficula auricularia Linnaeus (Derm. Forficulidae) [new rec.-CZ, atec (=Saaz),
NHMW-rev. TS]
(as Bigonichaeta) -[Gilpinia polytoma Hartig (Hym. Diprionidae)] [39; host must be wrong]
Neaerini
Phytomyptera cingulata (Robineau-
36 Vahara et al.  ENTOMOL. FENNICA Vol. 20
Desvoidy, 1830) -Tineidae sp. (ex Carpinus betulus) (Lep. Tineidae) [87]
Phytomyptera nigrina (Meigen, 1824) -[Anthonomus piri Kollar (Col. Curculionidae)] [39-as pyri; tachinid rev. by HE, but
host must be wrong]
Graphogaster brunnescens Villeneuve,
1907 -Acleris ferrugana Denis & Schiff. (Lep. Tortricidae) [18]
Siphonini
Ceromya bicolor (Meigen, 1824)
(as Actia) -Lasiocampa quercus Linnaeus (Lep. Lasiocampidae) [71, 72]
Ceromya flaviceps (Ratzeburg, 1844) (as
Actia-6, 39) -Dendrolimus pini Linnaeus (Lep. Lasiocampidae) [6-s.a. 39, 87]
-Cosmotriche lobulina Denis & Schiff. (Lep. Lasiocampidae) [87]
Actia crassicornis (Meigen, 1824) -Depressaria daucella Denis & Schiff. (Lep. Oecophoridae) [78-as nervosa]
Actia resinellae (Schrank, 1781) (as
nudibasis Stein-43) -Exoteleia dodecella Linnaeus (Lep. Gelechiidae) [43]
(as nudibasis Stein-12, 36; misid. as
pilipennis-39) -Retinia resinella Linnaeus (Lep. Tortricidae) [36-s.a. 12, 39-as Evetria-rev. HE,
new rec.-CZ, no locality, 1936, VULH-rev. HE]
(as nudibasis St.-29, 30, 32, 36, 50; misid.
as pilipennis-39; misid. as Tryptocera
crassicornis-113) -Rhyacionia buoliana Denis & Schiff. (Lep. Tortricidae) [29-as Evetria-s.a. 30-as
Grapholitha and 32-as Evetria, 36-s.a. 12, 39-rev. HE, 50, 113-as Retinia-rev. 87]
Actia pilipennis (Fallén, 1810) -Tortrix viridana Linnaeus (Lep. Tortricidae) [39-rev. HE]
-Archips xylosteana Linnaeus (Lep. Tortricidae) [39-as Cacoecia]
Siphona sp. (as Bucentes cristata;
cristata is prb. correct but not yet
confirmed) -Autographa gamma Linnaeus (Lep. Noctuidae) [69-as Plusia]
(as cristata incl. flavifrons-18 =prb.
cristata) -Ceramica pisi Linnaeus (Lep. Noctuidae) [2-as Mamestra-rev. 18]
(as Bucentes geniculata; prb. misid. =
prb. S. cristata) -Mamestra brassicae Linnaeus (Lep. Noctuidae) [105]
Leskiini
Demoticus plebejus (Fallén, 1810) (prb.
misid., identity unknown) -[Arctia festiva Hufnagel (Lep. Arctiidae)] [105-as hebe]
Leskia aurea (Fallén, 1820) -Synanthedon vespiformis Linnaeus (Lep. Sesiidae) [100-as Sesia asiliformis-s.a.
103-as Sesia asiliformis and cynipiformis, 105-as Sesia new rec.-CZ, Závist near
erná Hora, 1977 and Marovice near Moravský Krumlov, 1993, leg. and det.
Latvka, pers. comm.]
-Synanthedon myopaeformis Borkhausen (Lep. Sesiidae) [new rec.-CZ, Brno-Nový
Lískovec, 2006, leg. efrová, det. Latvka, pers. comm.]
Microphthalmini
Dexiosoma caninum (Fabricius, 1781)
(possibly misid. Dexia rustica) -Melolontha melolontha Linnaeus (Col. Scarabaeidae) [39]
Subfamily Dexiinae
Dexiini
Billaea adelpha (Loew, 1873) (ident.
not confirmed) -Isarthron fuscum Fabricius (Col. Cerambycidae) [82-as Tetropium]
Billaea irrorata (Meigen, 1826) -Oberea pupillata Gyllenhal (Col. Cerambycidae) [4, 33-s.a.]
(as Phorostoma parvula Portsch.-114) -Saperda populnea Linnaeus (Col. Cerambycidae) [4, 87, 114-rev.]
Billaea triangulifera (Zetterstedt, 1844) -Leiopus nebulosus Linnaeus (Col. Cerambycidae) [4-as Liopus]
-Oplosia fennica Paykull (Col. Cerambycidae) [24]
(as Gymnodexia) -Saperda octopunctata Scopoli (Col. Cerambycidae) [7]
(as Gymnodexia) -[Saperda populnea Linnaeus (Col. Cerambycidae)] [7, 103; doubtful because the
well-known parasitoid of this host is Billaea irrorata]
Dexia rustica (Fabricius, 1775) (as Dixia) -Melolontha melolontha Linnaeus (Col. Scarabaeidae) [115]
ENTOMOL. FENNICA Vol. 20  Host catalogue for Tachinidae 37
Voriini
Peteina erinaceus (Fabricius, 1794) -Autographa gamma Linnaeus (Lep. Noctuidae) [2-as Plusia-rev. 18]
Athrycia impressa (Wulp, 1869) (misid.
as Plagia ruralis-2) -Anarta myrtilli Linnaeus (Lep. Noctuidae) [2-rev. 18]
Voria ruralis (Fallén, 1810) -Autographa gamma Linnaeus (Lep. Noctuidae) [4-as Plusia, 105-as Plusia]
(as Plagia-2) -Arctia festiva Hufnagel (Lep. Arctiidae) [2-as hebe-rev. 18]
Hyleorus elatus (Meigen, 1838) (as
Athrycia elata) -Euproctis similis Fuessly (Lep. Lymantriidae) [6-as Porthesia]
Phenicellia haematodes (Meigen, 1824)
(as Phoenicella) -Arctia festiva Hufnagel (Lep. Arctiidae) [6-as hebe]
Thelaira nigripes (Fabricius, 1794) -Abrostola asclepiadis Denis & Schiff. (Lep. Noctuidae) [62]
Thelaira sp. (as leucozona Panz.; older
species identifications of Thelaira are
unreliable) -Laothoe populi Linnaeus (Lep. Sphingidae) [101-as Smerinthus-s.a. 105]
(as leucozona Panz.-101; nigripes F.-105) -Deilephila elpenor Linnaeus (Lep. Sphingidae) [101, s.a. 105]
(as leucozona Panz.) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [74, 105; questionable host]
(as leucozona Panz.; possibly correct in
this case because the true leucozona is a
parasitoid of Arctia caja) -Arctia caja Linnaeus (Lep. Arctiidae) [105]
Rondania dimidiata (Meigen, 1824) -[Zeiraphera rufimitrana Herrich-Schäffer (Lep. Tortricidae)] [87-tachinid
confirmed, but host must be wrong]
(misid. as Erynnia nitida-rev. HE) -[Lymantria monacha Linnaeus (Lep. Lymantriidae)] [37-host must be wrong]
-Liparus sp. (Col. Curculionidae) [39-as Molytes]
Dufouriini
Microsoma exiguum (Meigen, 1824) (as
Syntomogaster exigua) -[Tortrix viridana Linnaeus (Lep. Tortricidae)] [39-tachinid confirmed by HE, but
host must be wrong]
4. Host-parasitoid list
Lepidoptera
Tineidae -Tineidae g. sp. -Phytomyptera cingulata
Psychidae -Megalophanes viciella -Eumea mitis
Yponomeutidae -Yponomeuta evonymella -Bessa parallela, Blondelia nigripes
-Yponomeuta padella -Bessa parallela, Blondelia nigripes, *Phryxe vulgaris
Gelechiidae -Exoteleia dodecella -Actia resinellae
Oecophoridae -Depressaria daucella -Actia crassicornis
Zygaenidae -Zygaena carniolica -Phryxe prima, Ceromasia rubrifrons
-Zygaena ephialtes -Exorista larvarum
-Zygaena angelicae -Phryxe setifacies
-Zygaena filipendulae -[Senometopia pollinosa]
-Zygaena laeta -Phryxe magnicornis
-Zygaena punctum -Phryxe prima, Exorista larvarum
-Zygaena sp. -Ceromasia rubrifrons, Exorista larvarum
Sesiidae -Paranthrene tabaniformis -[Blondelia nigripes], [Pales pumicata]
-Synanthedon vespiformis -Leskia aurea
-Synanthedon myopaeformis -Leskia aurea
Cossidae -Cossus cossus -Xylotachina diluta
Tortricidae -Cochylimorpha hilarana -Blondelia nigripes
-Tortrix viridana -Actia pilipennis, Elodia morio, Bessa parallela, Nemorilla floralis,
Blondelia nigripes, Eumea linearicornis, Pseudoperichaeta
nigrolineata, Eurysthaea scutellaris, Zenillia dolosa, Phryxe
magnicornis, [Acemya acuticornis], [Microsoma exiguum]
-Acleris ferrugana -Bessa parallela, Graphogaster brunnescens
-Archips crataegana -Pseudoperichaeta nigrolineata, Eumea mitis
-Archips xylosteana -Actia pilipennis
-Archips rosana -Eumea linearicornis
38 Vahara et al.  ENTOMOL. FENNICA Vol. 20
-Choristoneura murinana -Nemorilla maculosa, Blondelia nigripes, Phryxe nemea, [Phebellia
triseta], [Carcelia sp.], [Eurithia connivens]
-Pandemis cerasana -Eumea linearicornis, Zenillia libatrix, Blondelia nigripes
-Pandemis heparana -Clemelis pullata
-Epinotia tedella -[Admontia grandicornis]
-Zeiraphera rufimitrana -[Rondania dimidiata]
-Zeiraphera griseana -Nemorilla maculosa, Blondelia nigripes, Pales pavida
-Retinia resinella -Actia resinellae, Lypha dubia
-Rhyacionia buoliana -Actia resinellae, Lypha dubia, [Acemya acuticornis]
Pyralidae -Dioryctria abietella -[Winthemia rufiventris], [Senometopia susurrans]
-Trachycera suavella -Nemorilla maculosa
-Ostrinia nubilalis -Lydella thompsoni
Lasiocampidae -Eriogaster lanestris -Pales pavida, [Eurithia consobrina]
-Malacosoma neustria -Carcelia gnava, Pales pavida, Compsilura concinnata, Exorista
larvarum, Blondelia nigripes, Zenillia libatrix, Phryxe vulgaris,
Phorocera assimilis, [Phorocera obscura], [Pseudoperichaeta
nigrolineata], [Hubneria affinis]
-Lasiocampa trifolii -Tachina grossa, Masicera silvatica, Phryxe vulgaris
-Lasiocampa quercus -Ceromya bicolor
-Macrothylacia rubi -Masicera silvatica
-Dendrolimus pini -Drino inconspicua, Blepharipa pratensis, Pales pavida, Blondelia
nigripes, Ceromya flaviceps, [Senometopia pollinosa], [Eumea mitis],
[Allophorocera ferruginea]
-Euthrix potatoria -Exorista larvarum
-Cosmotriche lobulina -Exorista larvarum, Ceromya flaviceps
-Phyllodesma ilicifolia -Rhacodinella apicata
Bombycidae -Bombyx mori -Compsilura concinnata
Saturniidae -Saturnia pyri -Masicera pavoniae, [Ernestia rudis]
-Saturnia spini -Masicera pavoniae
-Saturnia pavonia -Exorista grandis, Masicera pavoniae, [Exorista deligata], [Hubneria
affinis], [Linnaemya comta], [Linnaemya haemorrhoidalis],
[Ernestia rudis]
Sphingidae -Sphinx ligustri -Winthemia cruentata, Compsilura concinnata, Frontina laeta
-Sphinx pinastri -Phryxe erythrostoma, [Senometopia excisa]
-Mimas tiliae -Winthemia cruentata
-Smerinthus ocellatus -Frontina laeta, Masicera sphingivora
-Laothoe populi -Masicera sphingivora, Thelaira sp.
-Macroglossum stellatarum -Exorista larvarum
-Proserpinus proserpina -[Carcelia gnava]
-Hyles euphorbiae -Masicera sphingivora, Exorista grandis, *Blepharipa pratensis,
[Linnaemya picta]
-Deilephila elpenor -Drino lota, Winthemia quadripustulata, Exorista grandis,
Thelaira sp.
-Deilephila porcellus -Phryxe vulgaris
Papilionidae -Zerynthia polyxena -Winthemia quadripustulata
-Iphiclides podalirius -Compsilura concinnata
-Papilio machaon -Buquetia musca
Pieridae -Aporia crataegi -Phryxe vulgaris, *Aplomya confinis
-Pieris brassicae -Compsilura concinnata, Phryxe vulgaris, Phryxe nemea
-Pieris rapae -Compsilura concinnata
-Pieris napi -Compsilura concinnata, Phryxe vulgaris
-Anthocharis cardamines -Phryxe vulgaris
Lycaenidae -Neozephyrus quercus -Aplomya confinis, Phryxe nemea
-Satyrium pruni -Phryxe vulgaris
-Callophrys rubi -Cadurciella tritaeniata
Nymphalidae -Nymphalis polychloros -Sturmia bella, Phryxe vulgaris, [Exorista rustica]
-Nymphalis xanthomelas -Phryxe vulgaris
-Nymphalis antiopa -Pelatachina tibialis, Compsilura concinnata, Phryxe vulgaris,
Exorista larvarum
-Inachis io -Sturmia bella, Pelatachina tibialis, Compsilura concinnata, Phryxe
ENTOMOL. FENNICA Vol. 20  Host catalogue for Tachinidae 39
vulgaris, Winthemia quadripustulata, Blondelia nigripes
-Aglais urticae -Sturmia bella, Phryxe vulgaris, Compsilura concinnata, Blondelia
nigripes, Winthemia quadripustulata, [Phorocera assimilis],
[Carcelia gnava], [Bothria frontosa], [Blepharipa pratensis]
-Vanessa atalanta -Sturmia bella, Compsilura concinnata
-Araschnia levana -Compsilura concinnata, Phryxe vulgaris
-Melitaea athalia -Erycia fatua
-Argynnis paphia -Exorista larvarum
-Argynnis aglaja -Winthemia quadripustulata
-Issoria lathonia -Phryxe vulgaris
Geometridae -Abraxas grossulariata -Phryxe nemea
-Ennomos autumnarius -Blondelia nigripes
-Ennomos erosarius -Exorista fasciata
-Erannis defoliaria -Phryxe magnicornis, Zenillia libatrix
-Bupalus piniarius -Senometopia pollinosa, Blondelia piniariae, [Phryxe vulgaris],
[Zenillia libatrix]
-Operophtera brumata -Cyzenis albicans, Lypha dubia, Phorocera obscura, Blondelia
nigripes
-Operophtera fagata -Phorocera obscura
Thaumetopoeidae -Thaumetopoea processionea -Carcelia iliaca, Zenillia libatrix, Phryxe semicaudata, *Pales pavida
Notodontidae -Pheosia gnoma -[Senometopia excisa]
-Phalera bucephala -Compsilura concinnata, [Sturmia bella]
Noctuidae -Acronicta tridens -Nilea hortulana, Phebellia glauca
-Acronicta psi -Compsilura concinnata
-Acronicta aceris -Compsilura concinnata
-Simyra albovenosa -Exorista larvarum
-Catocala fraxini -Exorista fasciata, Exorista larvarum
-Autographa gamma -Voria ruralis, Phryxe vulgaris, Pales pavida, Peteina erinaceus,
Siphona sp., *Exorista larvarum
-Autographa jota -Exorista larvarum
-Abrostola asclepiadis -Thelaira nigripes
-Cucullia artemisiae -Eurithia consobrina
-Cucullia lactucae -Winthemia quadripustulata
-Cucullia umbratica -Winthemia quadripustulata
-Shargacucullia scrophulariae -Winthemia quadripustulata, *Baumhaueria goniaeformis
-Shargacucullia verbasci -Winthemia quadripustulata
-Calophasia sp. -Chetogena media
-Colocasia coryli -Nemoraea pellucida
-Cosmia affinis -Eumea linearicornis
-Cosmia trapezina -Eumea linearicornis, Eumea mitis, [Exorista rustica]
-Conistra rubiginea -Periarchiclops scutellaris
-Lithomoia solidaginis -Tachina fera
-Xylena exsoleta -Exorista larvarum, Blondelia nigripes
-Griposia aprilina -Phebellia triseta
-Hydraecia micacea -Lydella stabulans
-Hadula trifolii -Exorista larvarum
-Anarta myrtilli -Athrycia impressa
-Lacanobia oleracea -Pelatachina tibialis, Compsilura concinnata
-Ceramica pisi -Blondelia nigripes, Tachina fera, Exorista larvarum, Winthemia
quadripustulata, Siphona sp., [Prosopea nigricans]
-Mamestra brassicae -Siphona sp.
-Cerapteryx graminis -Gonia picea, Winthemia quadripustulata, Eumea mitis
-Tholera cespitis -Phryxe vulgaris
-Panolis flammea -Ernestia rudis, Pales pavida, Smidtia amoena
-Naenia typica -Drino lota
-Agrotis segetum -Spallanzania hebes, Exorista larvarum, Gonia capitata, Phryxe
vulgaris
-Agrotis sp. -Linnaemya impudica
Lymantriidae -Orgyia antiqua -Carcelia atricosta, Phryxe vulgaris
-Calliteara fascelina -Exorista fasciata
40 Vahara et al.  ENTOMOL. FENNICA Vol. 20
-Calliteara pudibunda -Carcelia gnava, Carcelia rasa, Compsilura concinnata,
[Hubneria affinis]
-Euproctis chrysorrhoea -Townsendiellomyia nidicola, Compsilura concinnata, Pales pavida,
Exorista larvarum, Zenillia libatrix, Zenillia dolosa, Baumhaueria
goniaeformis, [Pseudoperichaeta nigrolineata], [Linnaemya
impudica]
-Euproctis similis -Hyleorus elatus
-Leucoma salicis -Carcelia gnava, Exorista larvarum, Pales pavida, Carcelia
laxifrons, [Exorista mimula], [Sturmia bella]
-Lymantria monacha -Parasetigena silvestris, Pales pavida, Exorista larvarum, Drino
inconspicua, Compsilura concinnata, Tachina fera, Exorista grandis,
[Exorista rustica], [Phorocera assimilis], [Phorocera obscura],
[Senometopia excisa], [Erycia fatua], [Erycia festinans], [Phryno
vetula], [Ceromasia rubrifrons], [Allophorocera
ferruginea], [Allophorocera rufipes], [Blepharipa pratensis],
[Peleteria rubescens], [Ernestia rudis], [Thelaira sp.], [Rondania
dimidiata]
-Lymantria dispar -Zenillia libatrix, [Hubneria affinis], [Eurithia consobrina]
Arctiidae -Parasemia plantaginis -Peleteria ferina, [Phryxe vulgaris]
-Spilosoma lubricipeda -Eurithia anthophila
-Spilosoma luteum -Hubneria affinis
-Spilosoma sp. -Hubneria affinis
-Phragmatobia fuliginosa -Hubneria affinis, Exorista larvarum, [Senometopia excisa]
-Arctia caja -Hubneria affinis, Thelymorpha marmorata, Thelaira sp., Blondelia
nigripes, Exorista larvarum, [Phryxe vulgaris], [Carcelia gnava]
-Arctia villica -Hubneria affinis
-Arctia festiva -Phenicellia haematodes, Hubneria affinis, Exorista larvarum, Voria
ruralis, [Phorocera assimilis], [Phorocera obscura], [Demoticus
plebejus]
Coleoptera
Carabidae -Zabrus tenebrioides -Zaira cinerea
Scarabaeidae -Melolontha melolontha -Dexia rustica, *Dexiosoma caninum
Cerambycidae -Isarthron fuscum -Billaea adelpha
-Oplosia fennica -Billaea triangulifera
-Leiopus nebulosus -Billaea triangulifera
-Saperda octopunctata -Billaea triangulifera
-Saperda populnea -Billaea irrorata, [Billaea triangulifera]
-Saperda carcharias -[Pales pumicata]
-Oberea pupillata -Billaea irrorata
Chrysomelidae -Gastrophysa viridula -Meigenia mutabilis
-Plagiodera versicolora -[Medina luctuosa], [Medina melania]
-Chrysomela populi -Cleonice callida
-Chrysomela vigintipunctata -Cleonice keteli
-Linaeidea aenea -[Medina luctuosa]
-Gonioctena quinquepunctata -[Medina luctuosa]
-Galerucella lineola -Medina collaris
-Lochmaea capreae -Medina collaris
-Agelastica alni -*Meigenia mutabilis, [Meigenia incana], [Leiophora innoxia]
-Altica quercetorum -[Anthomyiopsis plagioderae]
Curculionidae -Anthonomus piri -[Phytomyptera nigrina]
-Liparus sp. -Rondania dimidiata
Hymenoptera
Argidae -Arge sp. -Belida angelicae
Cimbicidae -Abia sericea -Phebellia glirina
-Cimbex femoratus -[Carcelia atricosta]
-Cimbex sp. -Compsilura concinnata
-Pseudoclavellaria amerinae -Phebellia clavellariae
Diprionidae -Diprion pini -Drino inconspicua, Diplostichus janitrix, Blondelia inclusa,
ENTOMOL. FENNICA Vol. 20  Host catalogue for Tachinidae 41
[Exorista mimula], [Pales pumicata]
-Diprion similis -Diplostichus janitrix
-Gilpinia frutetorum -Diplostichus janitrix
-Gilpinia hercyniae -Diplostichus janitrix, Drino bohemica
-Gilpinia pallida -Drino gilva, Drino inconspicua
-Gilpinia polytoma -Drino inconspicua, Bessa selecta, Blondelia inclusa, Diplostichus
janitrix, Blondelia nigripes, [Triarthria setipennis]
-Gilpinia variegata -Diplostichus janitrix, Drino inconspicua
-Gilpinia virens -Drino inconspicua, Diplostichus janitrix
-Monoctenus juniperi -Staurochaeta albocingulata
-Neodiprion sertifer -Drino inconspicua, Blondelia inclusa, Drino gilva, Drino bohemica
Pamphiliidae -Acantholyda posticalis -Myxexoristops bonsdorffi
-Cephalcia abietis -Myxexoristops abietis, Myxexoristops bicolor, [Nilea hortulana]
-Cephalcia alpina -Myxexoristops bicolor
-Cephalcia arvensis -Myxexoristops bicolor, Pseudopachystylum goniaeoides
-Cephalcia sp. -Myxexoristops abietis, Pseudopachystylum goniaeoides
Tenthredinidae -Allantus truncatus -[Pales pavida]
-Cladius pallipes -Bessa selecta
-Cladius pectinicornis -Exorista mimula
-Hemichroa crocea -Bessa selecta
-Nematus melanaspis -Bessa selecta
-Nematus oligospilus -Eumea mitis
-Pristiphora abietina -[Bactromyia aurulenta]
-Pristiphora erichsonii -Bessa selecta
-Pristiphora pallidiventris -Exorista mimula
-Rhogogaster viridis -Exorista rustica
-Tenthredo arcuata -Exorista rustica
-Tenthredopsis coquebertii -Exorista rustica
-Tenthredopsis scutellaris -Exorista rustica
-Tomostethus nigritus -Hyalurgus tomostethi
-Trichiocampus grandis -Compsilura concinnata
Dermaptera -Forficula auricularia -Triarthria setipennis
5. Host record bibliography
[1] Brauer, F. & Bergenstamm, J. E. v. 1891: Die Zweiflügler des Kaiserlichen Museums zu Wien. V. Vorarbeiten
zu einer Monographie der Muscaria schizometopa (exclusive Anthomyidae), Pars. II.  Denkschriften der Akademie
der Wissenschaften in Wien, mathematisch-naturwissenschaftliche Klasse 58: 305446.
[2] Brauer, F. & Bergenstamm, J. E. v. 1894: Die Zweiflügler des Kaiserlichen Museums zu Wien. VII. Vorarbeiten
zu einer Monographie der Muscaria schizometopa (exclusive Anthomyidae), Pars. IV.  Denkschriften der Akademie
der Wissenschaften in Wien, mathematisch-naturwissenschaftliche Klasse, 61: 537624.
[3] Cagá, L., Turlings, T., Bokor, P. & Dorn, S. 1999. Lydella thompsoni Herting (Dipt., Tachinidae), a parasitoid
of the European corn borer, Ostrinia nubilalis Hbn. (Lep., Pyralidae) in Slovakia, Czech Republic and south-western
Poland.  Journal of Applied Entomology 123: 577583.
[4] apek, M. & epelák, J. 1970: Zoznam parazitov dochovaných z hmyzích kodcov. ás IV, Tachíny a mäsiarky
(Diptera) [List of parasites brought up from insectean pests. Part IV, Tachinidae and Sarcophagidae (Diptera)].
 Ponohospodárstvo 16: 254268. [In Slovak; Engl. & Russ. summ.].
[5] apek, M. & epelák, J. 1981: Zoznam parazitov dochovaných z hmyzích kodcov. VII. Tachíny a Mäsiarky
[List of parasites bred from insect pests. Part VII. Tachinidae and Sarcophagidae].  Ponohospodárstvo 27:
321332. [In Slovak; Engl. & Russ. summ.].
[6] epelák, J. 1952: II. Píspvek k poznání eských kuklic [IIe contribution  la connaissance des Tachinaires
tchques].  Acta Societatis entomologicae echosloveniae 49: 169180. [In Czech; French & Russ. summ.].
[7] epelák, J. 1955a: Píspvek k poznání slezských kuklic (Diptera-Larvevoridae) [Beitrag zur Kenntnis der schlesischen
Raupenfliegen].  Pírodovdecký sbornik Ostravského kraje 16: 222233. [In Czech; Russ. and Germ.
summ.].
[8] epelák, J. 1955b: Píspvek k poznání kuklic (Diptera-Larvevoridae) bourovce prsténivého (Malacosoma neustrium
L.) [Beitrag zur Kenntnis der Raupenfliegen des Ringelspinners (Malacosoma neustrium L.)].  Zoolo-
42 Vahara et al.  ENTOMOL. FENNICA Vol. 20
gické a entomologické listy [Folia zoologica et entomologica] 4: 167174.
[9] epelák, J. 1963a: Eschenblattwespe Tomostethus nigritus F. als Wirt einer neuen Raupenfliege Hyalurgus tomostethi,
n. sp. (Diptera, Larvaevoridae).  Biológia, Bratislava 18: 756759.
[10] epelák, J. 1963b: Píspevok k poznaniu kuklíc (Diptera-Larvevoridae) mniky zlatorítky (Euproctis phaeorrhoea
Dun.) [Contribution to the knowledge of brown-tail moths (Euproctis phaeorrhoea Dun.) tachinids (Diptera 
Larvaevoridae)].  Acta Universitatis Agriculturae (Nitra) 7: 125136. [In Slovak, Engl. & Russ. summ].
[11] epelák, J. 1963c: Príspevok k poznaniu kuklíc Slovenska (Larvaevoridae-Diptera) II [Contribution to the knowledge
of the tachinids of Slovakia (Larvaevoridae-Diptera) II].  Biologische Arbeiten 9/8, Entomologische Probleme
3: 5785. [In Slovak, Engl. & Russ. summ].
[12] epelák, J. 1997: Tachinidae: Part 1.  In: Vahara, J. & Rozkoný, R. (eds), Faunistic records from the Czech
and Slovak Republics: Diptera. Dipterologica bohemoslovaca Vol. 8.  Acta Facultatis scientiarum naturalium
Universitatis Masarykianae Brunensis, Biologia 95: 235.
[13] Cziek, K. 1906: Beiträge zu einer Dipterenfauna Mährens. 1. Die Umgebung von Brünn.  Zeitschrift des mährischen
Landesmuseums 6: 182234.
[14] Finlayson, L. R. & Finlayson, T. 1958a: Notes on parasites of Diprionidae in Europe and Japan and their establishment
in Canada on Diprion hercyniae (Htg.) (Hymenoptera: Diprionidae).  Canadian Entomologist 90:
557563.
[15] Finlayson, L. R. & Finlayson, T. 1958b: Notes on parasitism of a spruce sawfly, Diprion polytomum (Htg.) (Hymenoptera:
Diprionidae), in Czechoslovakia and Scandinavia.  Canadian Entomologist 90: 584589.
[16] Gold, J. 1893: Die Raupenfliegen.  Centralblatt für das gesamte Forstwesen 19: 300305.
[17] Gold, J. 1895: Entomologische Studien.  Naturalien-Cabinet 13: 196197; 14, 209210; 15, 225226; 16, 241
242.
[18] Herting, B. 1960: Biologie der westpaläarktischen Raupenfliegen (Dipt., Tachinidae).  Monografien zur angewandten
Entomologie Vol. 16. Parey, Hamburg, Berlin. 188 pp.
[19] Herting, B. 1961: Beiträge zur Kenntnis der europäischen Raupenfliegen (Dipt., Tachinidae), IIIVI.  Stuttgarter
Beiträge zur Naturkunde 65: 112.
[20] Hochmut, R. 1959: Píspvek k poznání morphologie, bionomie a populaní dynamiky obalee hlohového (Cacoecia
crataegana Hb.) [A contribution to the study of the morphology, bionomics and population dynamics of
Cacoecia crataegana Hb.].  Práce výzkumných ústav lesnických SR 16: 2358. [In Czech; Germ. & Russ.
summ.].
[21] Hochmut, R. 1964: Populaní dynamika obalee hlohového (Archips crataegana [Hb.]) v Dubinách SSR v letech
195761 [Population dynamics of Archips crataegana (Hb.) in the oak stands of Czechoslovakia in 1957
1961].  Práce výzkumných ústav lesnických SR 28: 3580. [In Czech; English summ.].
[22] Holik, O. 1937: Die Biologie von Zygaena punctum O.  Entomologische Rundschau 54: 3940.
[23] International Organisation for Biological and Integrated Control of Noxious Animals and Plants, 1989:  Determination
list of entomophagous insects Nr. 11. IOBC/WPRS Bulletin 12 (7), 63 pp.
[24] International Organisation for Biological and Integrated Control of Noxious Animals and Plants, 1993:  Determination
list of entomophagous insects Nr. 12. IOBC/WPRS Bulletin 16 (3), 56 pp.
[25] International Organisation for Biological and Integrated Control of Noxious Animals and Plants, 2005:  Determination
list of entomophagous insects Nr. 14. IOBC/WPRS Bulletin 28 (11), 71 pp.
[26] Jacentkovský, D. 1933a: Výskyt vzácných kuclic (Tachinidae) v SR [Occurrence of rare tachinids (Tachinidae)
in Czechoslovak Republic].  Sborník Vysoké koly zemdlské v Brn D, 20: 17. [In Czech].
[27] Jacentkovský, D. 1933b: Entomologické výzkumy v lesích Adamovských v letech [Entomological investigations
in forests near Adamov in 19301932].  Lesnická práce 12: 265289. [In Czech].
[28] Jacentkovský, D., 1934. Kuklice (Tachinidae) Masarykova lesa [Tachinidae from Masaryk forest].  Sborník
vysoké koly zemdlské v Brn D, 22, 138, 1 pl. [In Czech; French summ.].
[29] Jacentkovský, D. 1941a: K tachinologickému výzkumu kolního statku vysoké koly zemdlské v Brn [Zur tachinologischen
Durchforschung des Schulforstgutes der landwirtschaftlichen Hochschule in Brünn].  Sborník
entomologického oddlení Zemského Musea v Praze 19: 7680. [In Czech; Germ. summ.].
[30] Jacentkovský, D. 1941b: Kuklice (Tachinoidea, Diptera) Moravy a Slezska [Die Raupenfliegen (Tachinoidea)
Mährens und Schlesiens].  Acta Societatis scientiarum naturalium Moravicae 13 (4): 164. [In Czech; Germ.
summ.].
[31] Jacentkovský, D. 1942a: Tachinologická miscelanea [Tachinologische Miscelanea].  Sborník entomologického
oddlení Zemského Musea v Praze 20: 127131. [In Czech; Germ. summ.].
[32] Jacentkovský, D. 1942b: K tachinologickému výzkumu kolního statku vysoké koly zemdlské v Brn (Pokraování)
[Zur tachinologischen Durchforschung des Schulforstgutes der landwirtschaftlichen Hochschule in Brünn
(Fortsetzung)].  Sborník entomologického oddlení Zemského Musea v Praze 20: 172187. [In Czech; Germ.
summ.].
[33] Kadra, M. 1969: Oberea pupillata (Gyll.) (Col., Cerambycidae)  nový hostitel Billaea irrorata Meig. (Dip.,
Dexinae) [Oberea pupillata (Gyll.), (Col., Cerambycidae) ein neuer Wirt von Billaea irrorata Meig. (Dip., DexiENTOMOL.
FENNICA Vol. 20  Host catalogue for Tachinidae 43
nae)].  Zprávy eskoslovenské spolenosti entomologické pri eskoslovenské akademii vd 5: 50. [In Czech;
Germ. summ.].
[34] Kalandra, A. 1961: Verlauf der Massenvermehrung der gemeinen Gespinstblattwespe Cephaleia abietis L. im
Erzgebirge in den Jahren 19501960.  Communicationes Instituti forestalis echosloveniae 2: 173176.
[35] Kirchner, L. 1861: Zur Biologie der Sphinx pinastri.  Lotos 11: 103104.
[36] Kleteka, Z. 1996: Nálezy Dipter (Tipulidae, Tachinidae, Xylophagidae) zajímavé pro faunu ech [Remarkable
records of Diptera (Tipulidae, Tachinidae, Xylophagidae) from Bohemia].  Sborník Jihoeského muzea v
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44 Vahara et al.  ENTOMOL. FENNICA Vol. 20
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46 Vahara et al.  ENTOMOL. FENNICA Vol. 20
6. Discussion
The present catalogue lists 195 hosts for 149
tachinid species, but it must be taken into consideration
that some of the older tachinid identifications
or host associations are wrong or most probably
wrong. Thebibliography for thehostrecords
consists of 116 papers, which were based on the
work of 55 researchers.
The majority of the Czech host records concerns
the subfamily Exoristinae (103 tachinid
species, 579 records of 332 host-parasitoid couples,
making 82% of all Czech host records) and
to a lesser extent Tachininae (33 tachinid species,
91 records of 54 host-parasitoid couples, making
13% of all Czech host records) and Dexinae (13
tachinid species, 23 records of 34 host-parasitoid
couples, making 5% of all Czech host records).
The average for the Palaearctic region is not
much different (74% of the host records belong to
Exoristinae, 13% to Tachininae, 7% to Dexiinae,
and 6% to Phasiinae). There are no rearing records
of Phasiinae for the Czech Republic because
their hosts (Hemiptera) do not play a special
role as pests in this country. Members of
OrthopteraandDipteraareneither usually important
as pests in the Czech Republic. Accordingly,
there is not a single record for these host orders.
Most hosts in the Czech Republic have the unspecialized
parasitoids Exorista larvarum, Blondelia
nigripes, Compsilura concinnata and Phryxe
vulgaris, but this is not different from what is is
known from other European countries.
Investigations on tachinid-host relations in
the Czech Republic were primarily focused on
pestsinforestry,soitisnotsurprisingthatrecords
for such typical hosts as Lymantria monacha,
Dendrolimus pini, Euproctis chrysorrhoea, Ma-
lacosomaneustriaandTortrixviridanarankfirst.
But also some common host species which are
reared by practically every lepidopterologist (e.g.
Aglaisurticae,Vanessaio,Saturniapavoniaetc.)
are represented by many tachinid records. Larvae
of unimpressive hosts or of those which feed
ENTOMOL. FENNICA Vol. 20  Host catalogue for Tachinidae 47
ae 6: 6566. [In Czech; Germ. summ.].
[102] Vimmer, A. 1911: Píspvky k poznávání kukel hmyzu dvojkídlého. Diptera cyclorrhapha. ást prvá [Beiträge
zur Kenntnis der cyclorrhaphen Dipterenpuppen. I. Teil. Schizometopa).  Acta Societatis entomologicae bohemiae
8: 3443. [In Czech; Germ. summ.].
[103] Vimmer, A. 1913: Seznam eského hmyzu dvoukídlého [Catalogus dipterorum].  Entomologické píruky 8:
199. [In Czech].
[104] Vimmer, A. 1925: Larvy a kukly dvojkídlého hmyzu stedoevropského [Larvae and pupae of central European
Diptera].  eská grafická unie a.s., Praha. 348 pp. + 59 pls. [In Czech].
[105] Vimmer, A. 1928: Muí parasiti eských motýl. III. [Die Fliegenparasiten der böhmischen Falter. III.].  Acta
Societatis entomologicae echosloveniae 25: 4852. [In Czech; Germ. summ.].
[106] Vimmer, A. 1929: Dalí parasiti muí eských motýl (IV.) [Further parasites of Bohemian Lepidoptera].  Acta
Societatis entomologicae echosloveniae 26: 4647. [In Czech].
[107] Vimmer, A. 1930: Muí parasiti eských motýl. V. ást. [Fly parasites of Bohemian butterflies. Part V.].  Acta
Societatis entomologicae echosloveniae 27: 54. [In Czech].
[108] Vimmer, A. 1932: Druhy Tachin SR ze starého, velkého rodu Exorista Mg. [Die Tachinenarten d. SR der alten
grossen Exorista-Gattung].  Acta Societatis entomologicae echosloveniae 29: 126137. [In Czech].
[109] Vimmer, A. 1934: Nový parasit motýl [New parasite of butterflies].  Acta Societatis entomologicae echosloveniae
31: 189. [In Czech].
[110] Vimmer, A. 1935: Opt nový hostitel Tachin. [Again a new parasite of butterflies].  Acta Societatis entomologicae
echosloveniae 32: 46. [In Czech].
[111] Vimmer, A. 1938a: Parasiti eských motýl. VII. [Parasites of Bohemian butterflies. VII.].  Acta Societatis
entomologicae echosloveniae 35: 62. [In Czech].
[112] Vimmer, A. 1938b: Noví cizopasníci eských motýl. [New parasites of Bohemian butterflies].  Acta Societatis
entomologicae echosloveniae 35: 125126. [In Czech].
[113] Wachtl, F. A. 1882: Beiträge zur Kenntnis der Biologie, Systematik und Synonymie der Insekten.  Wiener entomologische
Zeitung 1: 275279.
[114] Wachtl, F. A. 1886: Einige Resultate meiner Zuchten.  Wiener entomologische Zeitung 5: 307.
[115] Weiser, J. 1960: Nemoci ponrav chroust v SR [Infections of Melolontha larvae in Czechoslovakia].  Acta
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[116] Zuska, J. 1963: The puparia of the European species of the family Larvaevoridae (Diptera) I (Subfamily Salmaciinae,
part 1).  Acta entomologica Musei nationalis Pragae 35: 333372.
hidden on roots or in stems, which live in or near
the soil, or which show a nocturnal behaviour are,
however, underrepresented  as usual. Future
search on interesting tachinid hosts should concentrate
on such animals.
Acknowledgements. For financial support we are indebted
to the Ministry of Education/Masaryk University for the
grants No. MSM 0021622416 and the Grant Agency CR
524/05/H536. We are grateful to I. Adamová (Brno, CZ)
for her help with finding tachinology literature. English
was kindly revised by P. J. Chandler (Melksham, Wiltshire,
GB).
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