Jenny,

I have worked with insects before and would like to pass along some tips
for working with them. We always used Bouins fixative for light
microspcopy and the glutaraldehyde buffered in a Phosphate buffer for
EM. Because you're doing IHC, I don't think the Bouin's would be a good
choice of fixative.  A word on cacodylate buffer:   My husband's lab
(the campus EM facility) has ceased using cacodylate buffer altogether
because of it's toxicity.  It is also probably more of a "fixative"
(notice the quotes) rather than a buffer.  It probably just simply
poisons the cells, rather than form linkages with proteins.  My husband
is the real expert on this, we just have these discussions over dinner
in our favorite restaurant where he conveys this info along to me. you
should see some of the looks we get!  *g*  I digress. 

Because you're doing IHC on paraffin embedded tissues, I would use 10%
Buffered Neutral Formalin with DMSO added to fix tissues in.  The reason
to put DMSO in the fixative is discussed below.  This has been a
critical ingredient in fixation of insects when I was working with them.
HOWEVER, I don't know how DMSO affects IHC.  If it turns out to be bad,
don't add it to the BNF formula.  Which case, I can't guarentee  how
well fixed your caterpillars will be.  Anyway, since you're a novice to
histology, I'll provide the formulation for this fixative (aren't I a
sweet heart?  *g*)

37% formaldehyde ---- 100 mL
DMSO ----------------   20 mL
Na2HPO4 -------------  6 g
NaH2PO4 -------------  4.5 g
DI water ------------ 880 mL

dissolve the phosphates in the DI water with the aid of stirring and
gentle heat (do not boil or let get so hot you cannot touch the flask).

Add the formaldehyde and DMSO in a fume hood, wearing goggles and
nitrile gloves.  Mix well.  Store this at room temperature in a tightly
sealed, unbreakable container.  Do not use if a white precipitate forms
or if it has been frozen.  Potassium phosphates may be used instead of
the sodium.  Some people I have worked with also add 2 -5% glacial
acetic acid, but i don't know how that will affect the IHC.  I would
avoid using acetic acid at this time.

DMSO is added to the fixative because all insects have a tough,
impermeable outer layer, the integument, that is designed to prevent
environmental things from entering the insect.  This includes larvae
(caterpillars are larvae).  If you're processing the entire caterpillar,
you will need to make slits in the sides with a very sharp razor blade
or scalpel blade.  These should be just deep enough to allow fixatives
and processing solutions to get inside.  The DMSO will help with the
infiltration.  Vacuum pressure is also very important to use in insect
histology to remove air bubbles trapped inside the body.  However, do
not use vacuum for long periods of time, just enough to remove the air
and let the solutions inside the body, then return to normal atmosphere.

Although I have never used microwave processing, my husband and others
tell me it's the greatest thing since hotcakes. If you have access to a
microwave, there are many people here who can tell you better how to use
it.

I hope this makes some sense... I've been running around doing things in
the lab, then coming back to this post, so it might sseem disconnected. 
hope it helps at any rate...

cheers!

Connie McManus


"J.L. Turnbull" wrote:
> 
> I am a biochemist by trade, but ahve wentured into some histology, rather
> unsuccessfully.  None of my biochemist contacts know anything about about this
> so i'd really appreaciate any assistance.
> 
> I have been trying to locate protein insecticides in sections of
> caterpillar gut with a  peroxidase-conjugated antibody.  I've done quite
> a few runs and changed many variables - still brown.
> 
> I based my protcol on a published paper using similar insects.
> I fixed in 2.5% gluteraldehyde 0.5M cacodylate for 24 hours.  Anatomy made
> sections in paraffin at 56 degrees.  I rehydrated by standard techniquwes
> and continued roughly as follows:
> 
> Incubate in iodine/K iodide
> 
> Equilibrate in Tris, Saline, Triton buffer.
> 
> Antigen revival in 1 mg/ml trypsin 10 min.
> 
> Quench endogenous peroxidase with 0.03% H2O2 in methanol 20 min.
> 
> primary monoclonal antibody in either milk. BSA, Haemoglobin or pre-immune
> serum.  Washed many times.
> 
> Either peroxidase-conjugated secondary antibody or VECTASTAIN ABC
> avidin-biotin kit.
> 
> Developed in either DAB or chloronaphthol in methanol,
> 
> Everything was brown, even antigen-free and antiserum-free controls.
> 
> Questions:
> 1) the protcol said that the iodine step was to remove the sublimate, what
> sublimate?
> 
> 2) How does the quenching step work?  I used the same recipies plus
> chromogen to develop and it didn't quench that?  Is it just oxidative
> damage?
> 
> 3)  I guessed that endogenous peroxidase was a problem (so haem. was bad
> choice then i realise).  I tried using sodium azide as an inhibitor (is it
> reversible or not?)  this didn't work either.
> 
> 4) i didn't know i had to filter the DAB - could this be the explanation?
> However. chloronapthol didn't work either.
> 
> 5) the other protocol used Hollandes fixative - would this make a
> difference.
> 
> 6) I tried doing this with whole guts (whilst i made more slides) just to
> see if the sectioning process had done something.  Most went brown but the
> 2 that i hadn't fixed gave a white control and brown sample.  Did i use
> the wrong fixative therefore.  I'm looking mainly at membranes, so i
> suppose methanol would have been better (I'm only just learning this
> histology business)
> 
> Please help (and sorry it's not more concise)
> 
> Jenny

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