Have been previously fixing tissue in 4% formalyin overnight and series of alcohols until 70% Etoh. We send out the samples for embedding (paraffin). This has always worked in the past but recently we see dissection of the tissue. Do anyone know a beter way of fixation?

Elly

Do you make the 4% formalin yourself? If so how? This can make a difference. The percentage seems off (do you mean paraformaldehyde powder made to 4%).
Do you fix at RT or 4 deg C?
I recommend fixing in 10% neutral buffered formalin (commercially available for reproducibility) and send out in 30% EtOH. 70% may cause extra drying in case it sits in it until processing (although isn't wrong at all).
What type of tissue are you processing? Human or mouse? What organ? If it is large you will need to fix longer. Is there necrosis in the tissue? Are you sure you are the one making the mistake?
Give me more info and I can help

Hi
I am working with mouse muscle tissue. And we are using two protocols. Currently we must also stain for presence of LacZ and therefore we fix the tissue in 4% paraformaldhyde (commercially bought, do not make) for 1 Hour at RT then leave in LacZ solution (x-gal buffer ) O/N at 37. The follwoing day it will be fixed in in 4% PFA and washed in Etoh 50 and 70%. From this point it is sent out. It has worked in the past and now the tissue is just horrible.
Elly

Well I am likely to think that 1 hour isn't long enough for fixation but if it has worked in the past for you I would suggest you call the company that embeds the samples ASAP and complain to them. It must be their problem.

Are your samples which didn't get B-Gal staining solution also as horrible as your B-Gal stained ones?