Hi,

I only partially agree. IHC works on many tissues fixed and stored in 70% ethanol for short-medium time. Here for example is the validation study:

http://ajcp.oxfordjournals.org/content/137/3/429.long

For routine stainings it depends on the tissue. For example brain stored in 70% ethanol may show vacuolation and loss of myelinisation.

Generally 24-48h storage in 70% ethanol at 4 degrees should not have negative impact on most of the tissues in most of the staining routines.

Good luck,

Hi dear Yael,

If you want use of section for IHC, it is not possible while for routine staining such as H&E you can wait for 1-2 month (sample dependence). However, i suggest you to prepare paraffin blocks immediately. 

Good luck, 

Hi,

I only partially agree. IHC works on many tissues fixed and stored in 70% ethanol for short-medium time. Here for example is the validation study:

http://ajcp.oxfordjournals.org/content/137/3/429.long

For routine stainings it depends on the tissue. For example brain stored in 70% ethanol may show vacuolation and loss of myelinisation.

Generally 24-48h storage in 70% ethanol at 4 degrees should not have negative impact on most of the tissues in most of the staining routines.

Good luck,

Hi Yael,

after PFA/Formalin fixation you can store your tissue in 70% ETOH  over night. Then continue  with the dehydration over ascending alcohols. 70% ETOH is the best place to store tissue  during embedding.  Higher concentrated alcohls harden the tissue to much.  The quality of your embedding depends on the tissue size and of the fixation.

If you think twice about the right storage time in the different embedding steps try a piece of  tissue  for testing to optimize your embedding protocol,

Good luck!  Ute

I agree with all the suggestions. It is possible to keep the tissues in alcohol for a while. However this is dependent what what tissue analysis you are planning. For H & E, yes. But for any other it is not recommended

Hi Yael

I used to store ovarian samples in 70 ethanol either for immediate embedding in paraffin, or for delayed embedding (up to several months) after fixation in Bouin or AFA (alcohol-formol-acetic acid). Histology is excellent in both cases and storage in 70 ethanol does not affect IHC results (for Ab working well with Bouin fixation). I have exclusively worked with ovaries and I am unable to say that storage in 70 ethanol is good for other tissues.

Hi Yael

I also used to store organs (liver, kidneys, testis, uteries) in Eth70 for 24 hours  after fixation in Bouin or Fomaldehyde 10% and there after i proceed with de-hydratation procedure (95x2,100x3, Xylenex2, at least 1h each) before embedding in parrafin. HE staining and Masson trichrome did after that was good, but less in tissues that were fixed in formalin.

Dear yael

The duration of fixation for a given tissue is very variable, depending on the size of the sample. For example, the neonatal mouse ovary requires 1h fixation, the adult mouse or rat ovary, 3h, whereas the human ovary (slice around 1cm thick) or monkey ovary requires 24h fixation. After this time, you must stop fixation (to avoid overfixation) by immersion in tap water for 24h.

If the samples are fixed in the solution containing alcohol for 24 to 48 hours then we can safely preserve the samples in 70% alcohol for a longer duration. It is a  very common practice for plant samples fixed  in formalin-acetic acid-alcohol  to store in 70% alcohol for several months. After fixation I don't think alcohol to cause any damage to the tissues.   

For our paraffin work we routinely have stopping steps at 50% EtOH and 100% EtOH but those are normally over night. Better stopping points would be to fix and then let the tissue sit in fixative or even 1xPBS until you're ready to dehydrate, otherwise we get tissue to 100 Paraffin and have left samples in 100% hot wax for a month in the incubator and still had luck sectioning and staining after embedding. I would not park it in any of the TBA steps or the infiltration steps fro more than 24-48 hours. I work with plant tissue and we routinely infiltrate for 24 hours at each step as plant tissue can be difficult to infiltrate. I agree with Karumanchi if you're using FAA. You can also use 5$ paraformaldehyde in 1 x PBS. For the most part damage is more likely to occur pre-fixation vs post fixation.

(;-) Many myths found in some replies in this thread...

Seconding necessarily different approaches for "animal" and "plant" tissue (naturally depending on the special tissue substrate and especially the size of specimens to be fixed and dehydrated). Unfortunately, Yael Haimovich has not added much more information on the specimen she has / had to process. IMHO, generally it is now accepted standard to interrupt dehydration (e.g. when processing manually until evening instead of using automated devices overnight) to store fixed specimens over night... better in 70% than in 100% EtOH (there might be some adverse elution of non-Formalin-fixed substrata out of the tissue and - agreed - might harden tissue too much with regard to problems also during paraffin embedded specimens' sectioning.

Longer storage in 70% ethanol is necessary for special reasons (cf. after fixation with BOUIN's or 'TNP'-containing fixatives [NB: 'TNP'= 2,4,6-trinitrophenol (TNP) or picric acid] several washings for up to 1-2 hrs each, or even longer- also overnight, esp. as long as no NH4 has been added).

Agnieszka Piszczek in her answer has pointed to a special article, dealing with and validating 'No adverse effects' on IHC-specimens, stating a relationship with / dependance from the tissue to be examined / processed, embedded, stained.... but also I agree to: "Generally 24-48h storage in 70% ethanol at 4 degrees should not have negative impact on most of the tissues in most of the staining routines" (whereby one should expect increased hygroscopy of the 4°-8° C "cold" C 70% EtOH when exposed to RT without protection from adsorption of air moisture.)