5/9/97
I am Resia Pretorius from South Africa and am currently working
on a PhD )the phylogeny of internal structures of the superfamily
Scarabaeoidea ((insects))
I would like to use a SEM to photograph one of the internal
chitinous structures (metendosternites) to which flight and leg muscles are
attached, and do calculations of different lengths for
morphometric analysis.
These muscles are attached firmly to the metendosternites and I
would like to remove them, but so far without luck. I have tried
Sodium perborate (different concentrations) they use this
chemical to clean muscles from snake skulls, but if only loosens the muscles
slightly, I still need to pull and tuck at the muscles to remove
them.
I have also tried tripsin a 0.5% without it working. Are there
any suggestions from anyone, I read in an 1890 article that one can
use nitric acid for the macercation of muscles but that was obviously
before the SEM was invented. Can one use an acid or would it
harm the actual structure of the metendosternites.
resia@mcd4330.medunsa.ac.za
Thanks,
Resia Pretorius
Lecturer (Biology)
Medunsa
South Africa
We've been doing this on Tingids for Dr Gerry Cassis here at the
Australian Museum. He's in a team which includes experts on Crustacea,
Arachnids, Trilobites, etc. who are looking at the evolutionary origins of
the arthropod groups.
The final protocol worked out by Sue Lindsay was;
Digest the muscle tissues in either KOH or NaOH (3 pellets in 3
or 4 mls water in watchglass at 70 degrees C). Leaving it in this
solution after the tissue has digested will also start to clear the exoskeleton.
Partially clear the exoskeleton for examination under transmitted
light microscopy by either leaving it in the caustic solution for much
longer or by BREIFLY boiling the solution. I'm told that Lactic Acid is
also useful for clearing exoskeletons. Light microscopy was useful for
detecting overlapping of sternites, relative thickness of sternites,
difference between sternite and arthrodial membrane, etc.
Clean in ultrasonic bath for extended time (say 5 minutes) to get
all the digested ooze out - obviously with some escape path such as the
holes where the head &/or abdomen were.
Dry the specimen. We got our best results with Critical Point
Drying, though this was because the tingids were a bit flimsy. Your
scarabs might me more robust, in which case HMDS (HexaMethyl DiSilazine) might
work well, or even just air drying from a volatile solvent (Acetone or
Ethanol?).
Dissect the specimen. We use eye surgery scissors, very small
blade length which worked on even the smallish specimens. Cut along the
dorsal and ventral midlines for two mirror-image samples; in our case, one
for SEM and the other for light microscopy.
Comments:
We found the order to be important, since cutting the specimen
first and then digesting the tissue lead to the skeleton just rolling up on
drying.
But this probably depends entirely on your particular specimen.
If given the choice I'd rather cut first and digest second, making
digestion quicker and allowing you to help it along by picking away at the muscle
with forceps. The ultrasound cleaning would also be easier. As
always in EM, you've just got to find what works with your particular beasts.
Geoff Avern
Manager
Microscopy Labs
Australian Museum
Sydney, Australia
geoffa@amsg.austmus.gov.au