Carol,
            in my younger, and some would say more foolish days, I studied
dragonflies including adults and larval stages. The major problem is that the
exocuticle becomes hard during the paraffin processing. I used a variety of
fixatives including Bouins and  buffered formalin  neither of which caused any
excessive hardening. Most of the hardening occurred in the subsequent
processing. The best solution was an old softening technic for chitin using
equal parts of chloral hydrate: phenol (warmed until a fluid mixture) prior to
the clearing stage. Specimens were dehydrated and then soaked in this mixture
overnight. If chloral hydrate is unavailable  to you, the next consideration is
the use of n-butyl alcohol for dehydration. This is miscible  with paraffin wax
although slowly. After dehydration, use a mix of n-butyl and paraffin wax at
room temperature. The specimens will not become hardened in that and the time in
wax is reduced.
A third alternate is to use chloroform as the intermediary agent as tissues
appear not to be hardened by prolonged exposure as they would in xylene or
toluene.
It would help to know which structures you are going to examine. The most
difficult but beautiful are the eyes. The abdominal region is probably the
easiest to cut.
Have you asked your grasshopper man if whole mounts of parts will be useful as
this was used in the past to study the mouth parts of insects. A cleared
specimen especially if the insect has ingested a material containing opaque
particles might be useful?
Call me if you need any more details.
Barry



"McCollough, Carol" wrote:

> -----Original Message-----
> From: brbenson@wyoming.com
> Sent: Monday, June 14, 1999 9:53 PM
> To: Histonet@pathology.swmed.edu
> Subject: Grasshoppers
>
> Hello,
>    I am asking for any help I can get and give to an entomologist on
> campus here.  He is looking for any special handling, processing, etc.
> for grasshoppers???>>
>
> Kim:
>
> I don't have anything specific on grasshoppers, but we do crustaceans here,
> which may present some of the same problems.  We fix tissues in Davidson's
> fixative for at least 18 hours.
>
> 95 percent ethanol              330 ml
> 37-40 percent formaldehyde      220 ml
> glacial acetic acid             115 ml
> distilled water                 335 ml
>
> Other fixatives that have been used on crabs in the past are Bouin's and
> Helly's, both of which have disposal problems that we prefer to avoid.
>
> We then transfer to 70 percent ethanol for short term storage.  Our
> processing schedule is all under vacuum:
> 70 percent ethanol      10 min
> 80 percent ethanol      10 min
> 95 percent ethanol      10 min
> 95 percent ethanol      10 min
> 100 percent ethanol     10 min
> 100 percent ethanol     10 min
> xylene                  10 min
> xylene                  10 min
> Paraplast Plus          30 min
> Paraplast Plus          20 min
> (Obviously the pieces of tissue we dissect out are small.)
>
> We embed in regular Paraplast and section at 5 microns using poly-L-lysine
> coated slides.
>
> Two references you may find helpful are:
>
> Messick, G. (1995) Laboratory techniques to detect parasites and diseases in
> blue crabs, Callinectes sapidus.  Techniques in Fish Immunology 4:187-200.
>
> Johnson, P.T. (1980) Histology of the blue crab, Callinectes sapidus: a
> model for the Decapoda.  Praeger.  440pp.
>
> Regards -
> Carol
> *****************
> Carol B. McCollough, HT(ASCP)
> Diagnostics & Histology Laboratory Manager
> Maryland Department of Natural Resources
> Fisheries Service
> Cooperative Oxford Laboratory
> 904 S. Morris Street
> Oxford, MD 21654