Question.
Here is my problem: tissue sections not adhering to the slides. Any hints on solving this problem?
Answers.
[Textbooks of microtechnique contain recipes for various adhesives: chrome alum-gelatin, Mayer's albumen and starch paste are traditional. More recent methods include giving the glass surface a positive charge by coating with polylysine or reaction with 3-aminopropyltriethoxysilane (APES or TES) to make "silanized" slides. See also the FAQ item on how to prepare silanized slides. There is also an FAQ item about polylysine. Here are some hints from individuals. No. 3 is pertinent to the use of any adhesive or none at all.]
1. We ran into the problem of tissues falling off the slides after about 5 hours of immunohistochemical processing. We seemed to have solved it with Super Frost Plus slides that have some sort of charge on them. Fisher/VWR I think carry them.
P. Emry
(emry[AT]u.washington.edu)
[Pre-prepared silanized slides are commercially available with a variety of trade names.]
2. We go to the expense of using charged slides for everything we do (Plus slides) and nothing really ever floats. If you don't want to go to that expense, we used to use chrom alum-gelatin with fairly good results and only an occasional problem. I personally don't like having chemicals in the waterbath. An exception would be immunos and some frozens for which I would recommend using "Plus" slides regardless.
Xylene in paraffin as a cause. I had an interesting thing happen to me once. I worked Saturdays for a while, training a girl at another lab in Histotechnique. The first Saturday we cut, most sections floated to varying degrees, even things like tonsil. The tonsil cut very nicely and seemed well processed. Well, I was supposed to be the person in the know and I was stumped. Took me a while but I finally figured out that they didn't change the processors very often and there was lots of xylene in the paraffin, and I mean lots. Apparently this was the problem, because after changing everything and rotating on a regular basis the problem went away. I just thought I would throw that story in - because this experienced histotech didn't realize that excess xylene in the paraffin could cause problems with adhesion of sections.
Marjorie A. Hagerty
(mhagerty[AT]emc.org)
3. Here is another possible contribution to the section loss. After picking up the ribbon on the waterbath do you purposefully pull out the water from between the section and the slide? ....you know, using a lap cloth (or whatever absorbant material you keep around) to touch the edge of the paraffin ribbon and soak up the water from under the ribbon. If the edges of the ribbon adhere to the slide but water remains between the section and the slide, when drying occurs, it is possible that not ALL of the water has evaporated from that space. Obviously, if a little water still separates the specimen from the slide (no matter what adhesive material is present), then the less than complete specimen attachment may not be strong enough to make it through the (even gentle) turbulence of the staining process.
This negative condition is most often seen when a ribbon is picked up and then the slide is immediately placed flat, horizontally, on th edge of the waterbath. It can also occur, though far less fequently, when the slide is immediately placed vertically against the waterbath or into a slide rack. The vertical positioning, however, does increase the draining of the water as long as the bottom of the ribbon has not fully attached to the slide creating a dam of sorts.
Anyway, that's just one more variable for you to consider before perhaps investing in something which may offer no greater adhesive advantage than what you are currently using.
Nancy Klemme
(nancy.klemme[AT]sakuraus.com)
4. Nancy is absolutely correct! Even with super adhesives or charged slides, you're liable to lose sections if the interface of the section and the microscope slide's glass is not water-free. This water is also a cause for "nuclear bubbling" artifact.
Ken Urban
Surgipath Medical Industries, Inc.
Richmond Illinois
(surgamy[AT]mc.net)
5. I bought 6 slide racks, the ones where slides stand on their ends, each holding 50 slides (Solmedia in the UK). These I keep for coating only. I've also got a couple of deep staining pots, again for coating only. I buy poly-L-lysine from Sigma or make my own gelatin-chrome-alum. Load the racks with slides, clean, I don't trust the manufacturers, wash thoroughly and coat with the coating of choice, dry and box. Couldn't be easier, I make enough in 2 days that will last months. Why be ripped off by the supply houses when for a few P.S./$ you can do it yourself.
Ian Montgomery
(I.Montgomery[AT]bio.gla.ac.uk)
6. Polylysine has free amine groups that form positively charged ions in water that's less alkaline than about pH 9. Slides are smeared with an aqueous solution of this basic amino acid polymer and then air-dried. This confers a positive charge to the slide's surface when immersed in water. Aminoacid anions(which Predominate in a section of a typical vertebrate animal tissue) are attracted to the polymer that covers the glass. It is a waste of money to buy poly-L-lysine rather than poly-D-lysine or poly-DL-lysine, because the stereochemical form of the amino acid does not affect its ionization. Buy the cheapest.
Positively charged slides can also be made in the reaction of an aminoalkylsilane with glass, in the presence of traces of water. It is easy to produce hundreds of "silanized slides in an hour. Alternatively, you can buy the silanized slides, which amounts to paying someone else's employer to do this simple job.
John A. Kiernan,
London, Canada
(kiernan[AT]uwo.ca)
7. I was satisfied with poly-L-lysine until I tried Superfrost Plus slides. I went from occasionally losing tissue to never losing it...so I vote for Superfrost Plus.
Mary Ross
(ross.8[AT]osu.edu)
Patricia Emry
(emry[AT]u.washington.edu)