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
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