From:Karen Larison <larisonk@uoneuro.uoregon.edu>


We use the 1% DMSO when staining embryonic zebrafish whole mounts. 
One lab here does embryonic chick and mouse whole mounts, and they 
don't use DMSO.  Apparently, critters that develop in utero or in ovo 
are different than those that undergo development in the environment.


>X-Sender: uvsgc@trex2.oscs.montana.edu
>Date: Mon, 23 Apr 2001 08:43:41 -0600
>To: Karen Larison <larisonk@uoneuro.uoregon.edu>, histonet@pathology.swmed.edu
>From: Gayle Callis <uvsgc@montana.edu>
>Subject: Re: DMSO/IHC
>What species?  Mouse embryos, zebra fish???
>At 02:43 PM 4/19/01 -0800, you wrote:
>>Connie and Jenny,
>>We routinely use 1% DMSO in our wash/dilution buffers when we do IHC.
>>We do a lot of IHC on whole mount embryos and the DMSO is required
>>for adequate penetration of the reagents.  I've also tested various
>>concentrations of DMSO.  1% DMSO works great, but greater
>>concentrations appear to destroy antibodies, etc.  In addition, we
>>have used 1% DMSO in our fixative with no deleterious effect on
>>antigenicity.  And we have used 10% DMSO to pretreat our fixed
>>embryos.  This is required to permeabilize the embryo if denaturing
>>fixatives have been used.  Antigenicity remains unaffected if the
>>specimen is washed thoroughly.
>>Use caution whenever you add DMSO to noxious chemicals like
>>fixatives.  DMSO is a carrier molecule and reportedly transports
>>molecules such as DAB and formalin through skin and gloves.  Wear
>>gloves, but also change them immediately if you have spills.
>>Karen in Oregon
>>>Date: Thu, 19 Apr 2001 14:38:32 -0600
>>>From: Connie McManus <conmac@cc.usu.edu>
>>>Subject: Insect processing tips --- long
>>>To: "J.L. Turnbull" <jlt25@hermes.cam.ac.uk>
>>>Cc: histonet@pathology.swmed.edu
>>>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
>>>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...
>>>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
>>>   > sections in paraffin at 56 degrees.  I rehydrated by standard
>>>>   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
>>>>   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,
>>>>   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
>>>>   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
>Gayle Callis
>Histopathology Supervisor
>Veterinary Molecular Biology
>Montana State University - Bozeman
>Bozeman MT 59717-3610
>406 994-6367
>404 994-4303 (FAX)

<< Previous Message | Next Message >>