Collecting and Preserving Parasitoids
Collecting
Rearing
What to do with the adults
Storing and Handling
Shipping
COLLECTING PROTOCOL FOR LIRIOMYZA
PARASITOIDS
The following is a brief summary of collecting protocol for rearing
Liriomyza parasitoids. Several further and more complete references
are listed at the end 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.
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 mouldy.
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
Example of a label
MALAYSIA, Selangor
Serdang, UPM Campus
coll. 24.iv.2000
J. LaSalle
ex. Liriomyza huidobrensis
on tomato
em. 25.iv.-6.v.2000
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. I 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 Liriomyza 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.
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, I always
collect any emerged parasitoid adults at the end of the afternoon before
I 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).
For the purposes of this project, 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 me for identification.
If material is going to be identified by project personnel or other
taxonomists, then contact either me or 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.
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.
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.
I have often received 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
I 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. I 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.
