[Histonet] RE: Dulbecco's PBS
TJJ <@t> Stowers-Institute.org
Fri Feb 23 08:14:57 CST 2007
1) Does it matter whether one uses PBS with or without Mg and Ca for immunos? I have always done it with PBS without Mg and Ca, and have never heard of anyone doing it with Mg and Ca, but have recently encountered a scenario in which this was the only PBS available. Wondering whether this will make a difference in my staining.
I don't know, I've never used Dulbecco's PBS. Gayle Callis uses this, perhaps she can address this point.
2) Can anyone explain the need for dehydrating and defatting slides (after using such substrates as DAB for color development) before coverslipping? Aside from the obvious that the sections look pretty crappy without these steps, can anyone explain the science behind this?
I don't think there is really anything scientific about this practice. At this point we are not using alcohol and xylene to defat the slides. I think it's more that after counterstaining, the slides are in water and water is incompatible with xylene or toluene based mounting media. Therefore, we dehydrate into graded alcohols. Xylene is used as the intermediary step when the mountant is incompatible with alcohol, and the slides are coverslipped wet from this reagent. Surgipath has an mounting medium, Clearium, that you can coverslip from alcohol.
While it is possible to air dry the slides after the water rinses after hematoxylin (or other) counterstain, and dry coverslip them using permanent mounting medium, we routinely dehydrate, clear and mount to coverslip H&Es and other histochemical stains. Why should we treat the immunostained slides differently? (Unless, of course, the chromogen is soluble in organic solvents.)
It is possible there are imaging issues that might crop up by coverslipping air-dried slides, but I'm personally not aware of any.
3) Because my tissue is perfused, I typically store the brains in 30% sucrose (with a little bit of sodium azide) in 4 degrees, until I am ready to cut the blocks. At this point, brains are usually frozen in the cryostat just before cutting 40um thick sections. I have obtained wonderful staining and morphology with this technique, however I have been recently advised to store the brains embedded in OCT (or similar) in a -20 degree =66reezer, rather than in 4 degrees. I am skeptical and concerned that this will damage the morphology and produce that "swiss cheese" freezer artifact (you know, where there are all these holes in the tissue?) Can anyone provide any advice, insight as to the benefits and drawbacks to either/both of these methods?
If you snap-freeze your samples and store them properly in a -20 (or -80) degree C freezer that is not self-defrosting, your samples will be fine. The freezer artifact would come from improper snap freezing, or repeated freeze/thaw cycles upon storage. I have found that samples stored long term in -80 may be more difficult to section. While it doesn't damage the sample, it does change it's sectioning characteristics as compared to samples stored at -20.
It would be my preference to store them long term frozen in OCT than in an aqueous solution. Although your tissue is perfusion fixed, it may still be possible to reverse the aldehyde cross-links with long term storage in aqueous solutions. I don't know for sure if the sucrose blocks that possibility or eventuality, and because I don't know that, I'd rather be safe than introduce another variable into a research hypothesis. Do you know for sure there is no difference if you store Sample A for 2 weeks in sucrose in the fridge, and when you repeat the experiment with Sample B (which was harvested at the same time as sample A), it has been stored in the fridge for an addition week (or two or three), etc.? How long do you store them in the refrigerator? You don't mention this.
I would worry about solubility of certain antigens or proteins as time in storage was increased. Again, I have no evidence to suggest it happens, as proving that for particular antigens or proteins is a research experiment in itself. Our responsibility to the researchers here is to preserve the integrity of their sample and not introduce variables from sample to sample as much as possible. The best way to accomplish that is to treat all samples the same, and our practice is to freeze them after storage in 30% sucrose after 24 hours.
My first thought about storage in 30% sucrose was microbial growth, but I see you use sodium azide in your solution. Have you seen any evidence sodium azide remains in your sample, possibly inhibiting HRP in your subsequent immunostaining?
Stowers Institute for Medical Research
Kansas City, MO 64110
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