Key to the World Genera of Eulophidae Parasitoids (Hymenoptera) of Leafmining Agromyzidae (Diptera)

Placido Reina^* and John La Salle*

^Dipartimento di Scienze e Tecnologie Fitosanitarie, Sezione Entomologia agraria - University of Catania, via S. Sofia, 98 - 95124 Catania, ITALY - [email protected]

*CSIRO Entomology, GPO Box 1700, Canberra, ACT 2601, AUSTRALIA - [email protected]

Introduction

About Eulophidae

The Key...

Tutorial

Collecting

Some links

Thanks to...

About Eulophidae

Eulophid wasps are the most common parasitoids recorded on leafminers worldwide, and the third most important family of Chalcidoidea in relation to all biological control successes, after Aphelinidae and Encyrtidae (La Salle & Schauff, 1995). Moreover, they are the most successful agents used within biological control programs against agromyzids (Minkenberg & van Lenteren, 1986; Waterhouse & Norris, 1987; Konishi, 1998; Murphy & La Salle, 1999). Diglyphus Walker, Chrysocharis Förster and Neochrysocharis Kordjumov, particularly, have been using successfully against Diptera leafminers, especially under greenhouse conditions. Other genera, e.g. Quadrastichus Girault, Cirrospilus Westwood, Citrostichus Boucek, Galeopsomyia Girault, have been used in the biological control of Lepidoptera leafminers under field condition (Noyes, 2003).
Higher classification of Eulophidae has recently been revised by Gauthier et al. (2000). It is currently divided into four subfamilies, Entedoninae, Euderinae, Eulophinae and Tetrastichinae.

FOR MORE INFORMATION ON EULOPHIDAE, GO TO THE EULOPHIDAE PAGE.

About the key

In this website, we provided a key of the world genera of eulophids recorded on agromyzid leafminers. We used 25 characters and recognized 29 genera, divided in the subfamilies as follows: 6 genera within Tetrastichinae, 10 within Entedoninae, and 13 among Eulophinae (7 Cirrospilini and 6 Eulophini).
For each genus, notes are given for diagnosis, classification, distribution, and biology. Taxonomic comments and numerous pictures have been also provided in order to facilitate their recognition. We however considered separately two taxa within Cirrospilus Westwood: the species C. ambiguus Hansson & La Salle and the “variegatus group” of species. Both taxa have in fact unique features within Cirrospilus, and warrant further investigation as to their exact generic limits. We were unable to include two other genera belonging to the subfamily Eulophinae and recorded as agromyzid parasitoids (Noyes, 2001): Ginsiella Erdös and Guptaiella Khan & Sushil (see comments under these genera).
Finally, some notes, supported by pictures, about the family Eulophidae and its subfamilies are given. Notes on Elasmini are also provided, as there are some Lepidoptera leafminers parasitoids within this tribe. We haven’t considered the tribes Gyrolasomini and Euderomphalini and the unplaced tribes Platytrecampini, Anselmellini and Ophemilini, as none of these have ever been recorded as parasitoids of leafminers.

AN INTRODUCTION TO HOW TO USE THIS KEY IS GIVEN ON THE TUTORIAL PAGE.

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Collecting leafminer parasitoids

The following is a brief summary of collecting protocol for rearing Agromyzidae leafminers parasitoids. Several further and more complete references are listed on collecting and preparing either insects in general (Martin, 1977; Walker & Crosby, 1988; Borror et al., 1989: 745-789; Huber, 1998), or Hymenoptera specifically (Prinsloo, 1980: 3-7; Noyes, 1982; Gauld & Bolton, 1988: 48-57; Grissell & Schauff, 1997: 54-60). In addition, Shaw (1997) specifically treats the subject of methods of rearing parasitic Hymenoptera.

1. COLLECTING

- Try to collect parasitoids when they are in the pupal (or late larval) stage.
There is generally a much lower success rate when trying to rear parasitoids which have been collected in the field as larvae.

- Prevent material from becoming moldy.
If you must place parasitoids in plastic bags, ensure that there is some tissue paper or paper toweling in the bag to absorb moisture (and this may have to be changed). Specifically, do not place material in sealed plastic bags and leave in a hot car! You can carry an ice chest or cool box for placing material.

- Gather all essential information when collecting material.
Information on the specimens should include (wherever possible):
Precise collecting locality – MALAYSIA, Selangor, Serdang, UPM Campus
Date Collected (with day in regular numbers, month in roman numerals, year) – collected 24.iv.2000
Date parasitoids emerged – emerged 25.iv-6.v.2000
Collector – coll. J. La Salle
Host – ex. Liriomyza huidobrensis
Host plant – on tomato

- Examples of labels
MALAYSIA, Selangor
Serdang, UPM Campus
coll. 24.iv.2000
J. LaSalle
ex. Liriomyza huidobrensis
on tomato
em. 25.iv.-6.v.2000

2. REARING

- Rear parasitoids in an environment that resembles field conditions.
If there is an outside shed or greenhouse which is close to natural conditions, this is good. Screen doors which allow air flow through a rearing area are good. We generally rear parasitoids on a shelf next to a window – BUT make sure that they are not getting overheated in direct sunlight.

- Isolate hosts as much as possible.
Try to isolate the Agromyzidae larvae and pupae. This means to remove as much excess plant material as possible, and check the remaining plant material for other potential host insects (such as aphids, mealybugs, etc.). This will prevent getting large numbers of unwanted parasitoids, which can waste time and expense in getting these specimens collected, curated, and shipped to specialists when they are not part of the project.

- Prevent mold and sweating.
There are two ways of trying to do this.
The first is to lower the humidity, and get an air flow through the sample. You can do this by holding the parasitized larvae and pupae with as little plant material as possible, keeping them in something with an air flow (such as a glass tube with cloth or cotton over one end). However, this may allow the specimens to get too dry, which can prevent the parasitoids developing.
Another method is to keep the parasitized larvae or pupae in sealed containers to retain humidity, but if you do this you will have to keep some tissue or paper towel in the container, and you will have to change this absorbent material regularly (probably daily).

- Prevent drying out.
This is difficult, particularly if you are trying to keep material dry to prevent mold (see above). If you have collected pupae, they are generally easier to handle, and by the time they pupate the host is usually fairly well consumed. You can keep them with a minimum of plant material in glass tubes. It might be easiest to keep them in glass tubes (where there is air flow), but place them in a sealed container which has a damp paper towel in it. If you are trying to rear larvae, it may be necessary to have the moribund host remains while the larvae finish their development. This may have to be kept on plant material, and it might be worth trying to keep them in larger containers (rearing cages), by taking whole leaves or stems and placing the bottoms in tubes with water. If you use a cork cut in half with a hole in it, you can generally place this around a stem and seal the bottom of the stem in a tube with water. However, in general this method can be quite difficult and time consuming, and may not be necessary for parasitoid survey work.

You will generally have to try a couple of different techniques to try and get the humidity balance right for rearing the parasitoids.

3. WHAT TO DO WITH THE ADULTS

- Do not kill the adults in the morning
In general parasitoids emerge early in the morning, and killing them at this time is bad because they have not had time to harden up and fully develop their colour. So, specimens killed early in the day tend to shrivel, and be more difficult to work with and identify. So, we always collect any emerged parasitoid adults at the end of the afternoon before we go home rather than first thing in the morning.

- Kill in 75-95% EtOH
There are several methods of killing and preserving specimens (see references). It is most expedient to kill specimens directly in ethyl alcohol – generally some range between 75 and 95%. This will facilitate sorting and shipping. Note: the above paragraph is true if you are going to send the parasitoids to specialist for identification. If material is going to be identified by project personnel or other taxonomists, then contact them to determine other methods of curation which might be more appropriate. There is no real need to keep every specimen in an individual tube – but do not mix parasitoids collected from different collecting localities or hosts. It is generally not important to have daily records of emergence, but weekly or monthly are nice to determine which species are present when.

- When putting specimens in a tube – ALWAYS MAKE SURE THAT COLLECTING DATA ARE PRESENT.
Do not trust your memory to remember which parasitoids came from which collection.
If you are putting a label in a vial containing alcohol, make sure the label is written with pencil, or waterproof ink. Otherwise it will run in the alcohol.

4. STORING AND HANDLING

- Keep alcohol specimens cold and dark.
After placing specimens in alcohol, they can deteriorate very quickly when exposed to light and heat. To ensure the best quality specimens, you should store specimens in alcohol in the freezer, or a refrigerator; but if this is not possible at least keep them in the dark in a cupboard and in as cool a place as possible.

- Keep dried material dry and away from insect pests.
If you have dry specimens (either mounted or unmounted) keep them away from areas of high humidity (where they may be attacked by fungi), and place some insect repellent (moth balls, naphthalene, etc.) with them to prevent attack by psocids, roaches, dermestid beetles, etc. DO NOT put any moth balls, crystals, etc inside the same box as the specimens, as they may come lose and damage specimens.

5. SHIPPING

- Keep tubes full
Make sure tubes are full of alcohol before shipping. If the alcohol can slop around in the tubes, this could damage the specimens.

- Make sure the tubes are sealed correctly.
We have often seen tubes where the alcohol has all spilled out (past a loose cap, or for whatever reason). This makes identification more difficult (or sometimes impossible). Make sure the caps are tight, and we generally place tape or parafilm around the caps.

- Make sure the tubes can not rattle together.
Mever let glass tubes come together in such a way that they can bounce around and break. We generally wrap each tube individually in paper towel or tissue, or place them in holes cut in a piece of old styrofoam. Other methods are possible – use your imagination, but do not let the tubes knock together.

- Make sure there is a protective layer around the package containing the tubes.
Place the container with the tubes in a larger box, and make sure that there is a protective layer around it (styrofoam chips, foam, styrofoam blocks, bubble wrap, etc.).

- Make sure each tube has a proper label inside it.

Some useful links

Universal Chalcidoidea Database (by John Noyes)

Glossary for Chalcid Wasps (by Gary Gibson)

Australian National Insect Collection (CSIRO Entomology)

Biological control of Liriomyza leafminers (Murphy & La Salle, 1999)
Dutch leafminers
Japanese key of leafminer parasitoids (by Konishi)
Research on Chalcidoid Systematics (by University of California)
Systematic Entomology Laboratory - Hymenoptera (USDA)
Insect and Spider Collections of the World
Parasitic Hymenoptera Research Laboratories (Department of Entomology, Texas A&M University)
International Society of Hymenopterists
Hymenoptera at OSUC (The Ohio State University)