The only special stain that I know that requires the use of a negative control is for the AFB. I understand to rule out false positives as the AFB bacteria might exist in tap water. Nevertheless, a good QA practice which we will implement now.
Other than Carson, does anyone know of a regulatory or accreditation agency is requiring this as well? Any suggestion on a good control tissue type? Carson recommends uterus. Also if there is a pick up on the negative slide (link to the tap water) will use of distilled water and a repeat procedure fix this? Any thoughts from fellow histonetters? Thanks Ian Bernard -----Original Message----- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Lee & Peggy Wenk Sent: Wednesday, March 13, 2013 9:21 PM To: Tighe,Sean T; histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Alizarin Red S Staining Protocol We always used a 2% Alizarin red S (never tried it with 1%. I'm guessing that you would just have to leave it in longer.) We pH it to 4.1-4.3 with 0.5% ammonium hydroxide. I don't remember what the pH is when we begin, but note, we're adding a BASE to bring the pH up to ~4.2. So alizarin red S must be fairly acidic when dissolved in water. I was told that this is a chelating solution for soft metals. It therefore is not a specific stain for calcium, as it will also stain magnesium, manganese, barium, and strontium. However, these metals are usually not in very high concentrations. But to make it more specific for calcium, a pH of 4.1-4.3 is recommended. I've never done this stain on cell cultures, only on formalin fixed, paraffin embedded, 5 um thick sections. And we find that staining between 1-2 minutes works the best. This is a very sensitive stain. The longer the slide is left in the solution, the more and more the alizarin red S stains the background, to the point that it becomes difficult to tell positive calcium from background staining. Every cell has calcium in it (think membrane transport), and eventually, all the cells are going to pick up background orange color, not just the lesions with calcium. You mention extraction with acetic acid. Extracting what? Are you trying to extract the background staining? I don't know if that will work. This is a chelating agent . . . the CLAW! Once chelating agents hook up with the metal, they don't let go. I just don't happen to know if acid will disrupt this chelating bond. So it would be better to cut the time way down (try 1-2 minutes, see how it looks), and don't have so much background staining to begin with. Let us know how it turns out. Peggy A. Wenk, HTL(ASCP)SLS Beaumont Health System Royal Oak, MI 48073 Opinions expressed are mine, and do not reflect upon my place of employment. -----Original Message----- From: Tighe,Sean T Sent: Wednesday, March 13, 2013 4:48 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Alizarin Red S Staining Protocol Good afternoon, In the past I have been using a 40mM Alizarin Red S Solution from Millipore to stain my cell cultures but I am now seeking an alternative. I plan on preparing fresh Alizarin Red S by adding the powder to 100ml of distilled water. In reviewing the literature, I have come across many labs using 1% ARS or 2% ARS and I am not sure if this concentration significantly matters. Furthermore, I understand some people use McGee-Russell's procedure using a pH of ~4.2 whereas others use Dahl's procedure using a pH of ~6.3. However using a pH of 4.2 seems illogical to me as this could remove some calcium from the monolayer. Currently I plan on fixing the cells with 4% Paraformaldehyde in PBS (pH 7) for 15 minutes, washing twice with water and then staining the cells with 2% ARS in water (pH 6.3). Do you see any problems with this? Also, should I stain for 5 minutes or roughly an hour with the ARS dye? I have had some problems with extracting this dye in the past. The 10% acetic acid does not seem to remove all of the dye and the results are variable. Note: before I extract the cells with acetic acid I wash the cells four times with water. Regards, Sean Tighe _______________________________________________ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet _______________________________________________ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
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