Jan,

In a pinch, we have stored fixed whole mount embryos in methanol kept 
at -20C.  You must first dehydrate.  I think this yields high 
background, but it's better than nothing.

Also, we have stored fixed mouse organs at -20C in the following 
cryoprotectant:

500 mL 0.1 phosphate, pH 7.2
300 g sucrose
300 mL ethylene glycol

bring volume to 1 liter with distilled water.

I then wash them in several changes of 30% sucrose before 
cryosectioning.  It seems like it would work for tissues fated for 
paraffin sectioning after washing thoroughly with water.  The 
original reference on this technique:  Use of Cryoprotectant to 
Maintain Long-Term Peptide Immunoreactivity.." Peptides 7. pp 
155-159, 1986.



Good luck.

Karen in Oregon







>Usually the tissue samples I receive have been optimally fixed (time-wise)
>in formalin and promptly embedded into paraffin.  Today I received a call
>from a researcher out of state who wants to send me some samples, but cannot
>get his fixed tissue processed into paraffin within a 24-hour period after
>start of fixation.  He wants to know what storage alcohol he should switch
>the samples into until embedding, and I became brain-dead.  Can someone help
>me out here PDQ?  Is it 70% ethanol or some other percentage?  Is there a
>better storage medium than ethanol?  I have received some cases in PBS
>(post-formalin-fixation), but those staining results were less than optimal.
>
>Any help would be greatly appreciated.
>
>Jan Shivers
>Univ. of Minn. Vet. Diag. Lab