Try mounting your sections on poly-l-lysine or TESPA treated slides. Also be very gentle with the water washing-put you slide rack at the bottom of a 5litre beaker that is already full of tap water, and then put it under a slowly running tap till the slides are sufficiently destained without any turbulence.

is the slide positive charged?

Is this FFPE section or frozen section.

possible, slides/tissue was not completely dry, was the room too damp?

I am not too familiar with 50 um section. Sounds to me whether for regular histology or IHC is too thick.

Hi i had the same problem with 5um cryosections on those slides that came off at first wash. i could save mine by a fixation step with aceton (-20°C) for 20 minutes prior to the staining procedure. Now i use + coated glass slides from which they don't float off.

Why do you dry your slides so long before the counterstaining and why not counterstaining immediately after your DAB staining?

We always use the Superfrost plus slides (although expensive) and place slides on slide warmer for several hours before commencing staining procedures. We have had problems with sections floating when we have placed slides with annealed sections in the freezer for staining in future.

Why was it left to air dry for longer? did it take longer to dry? Humidity may be a factor. try air drying in an dry oven at 37degC. Other possibilitites - your gelatine has degraded, your slides are not coated evenly. Using commercially available negatively charged slides are a great help in preventing sections floating away.... Hope this helps... all the best

hi i had the same problem with 5um cryosections on those slides: they also came off at first wash step but the next sections i could safe by a fixation step with acetone (-20°C) for 20 prior to the staining procedure. Now i use + coated slides and they don't come off anymore, even without the acetone step.

But why do you dry your sections for so long before counterstaining en not immediately after your DAB staining?

You could switch to a positive charged slide. It helps bond the tissue better through its negative charge and keeps it from floating off. Maybe you just grabbed the wrong box of slides...

It is not very unusual though. It happened to me on several occasions. For regular immunostaining I prefer free-floating sections, stain them in the wells as long as you can and transfer them on to the gelatin-coated slides, let them dry not for a long time, pass them through dehydration steps, clearing in xylene and mount them with permount.

But for radioactive/non-radioactive in situ hybridization experiments I have always used APES coated slides (you can do the coating yourself following the direction) for attachment of the sections firmly. I never tried them for regular histological/histochemical staining. You can try using them as well. Good luck.

I encountered same problem on last few runs. Use of Vectabond will help you.

Are these frozen sections?

We use the SuperFrost Plus slides from Fisher for all our staining. The positive charge on the slide really helps keep the tissue on.

Also, depending on how "gentle" you are when you stain/wash can dictate if the tissue begins to come off. Do you apply the water with a transfer/pasteur type of pipette? You might try this as well.

Maybe also try drying the tissue on a slide warmer...?

You may find that pre-baking slides at 65 for 10 may help.

Instead of gelatin coat the slides with Poly L Lysine!

The glycocalyx of cells is negatively charged so a positively charged surface will bind them better. If the tissue is aldehyde fixed it has a very strong negative charge from the aldehyde. Poly L Lysine is the substrate of choice!

I agree with Ramesh Nayak. Poly L Lysine coated slides are the best adhesives

When preparing your slides (days ahead of time), dip slides in gelatin, dry in a rack with a fan for hours; dip in gelatin again, dry again (= double-dipped gelatin-coated slides). This will help. Fisher superfrost slides are good, too but if you just opened a new box, try a new box -- I've had problems with individual boxes or batches of slides before (never have this problem if you coat your own slides).

When mounting your sections to the slides, you can add a bit of gelatin to your mounting solution; this has the added benefit of making them stick better immediately, so mounting is easier. When mounting, take care to ensure there are no wrinkles (this all assumes you have well-fixed tissue that was sliced smoothly on a microtome with a sharp blade -- 50 microns = thick sections, so this all has to be done well). Blot slides well on the edge; dry overnight on a flat tray. Dehydrate the sections before coverslipping (and/or in the process of counterstaining).

If you do all this, the sections should never come off the slide (for better or for worse). Good luck!

It can happen. Try to catch them back on the slide or on another, clean slide, immediately or after you finish washing, next incubation etc . Or use SuperFrost, super frost Plus slides or other tricks supplied by producers. They are really good

I do prefer superfrost slides (and I've seen really old batches perform less than adequately in comparison to a fresh box.. something to keep in mind). That said, if your gelatin solution you are treating the slides with is really old you can see a decrease in performance there.

Super plus or Poly L slides are very good but you can occasionally get a bad box (i.e. sections fall off). This has happen to me before (although very rarely). In my experience, slightly over drying doesn't seem to affect section adhesion.

In addition, to the good advice from Joel Geerling - Be sure that you do not place your sections so close together that the tissue from the current section rests on the mounting medium from the previous section. This will greatly increase your likelihood of losing sections as those areas will not adhere to the slide well.

I agree with Kelsey Potter to use superfrost slide from Fisher ,VWR or Mercedes medical. and wash gentle with tape water. or also try Poly l Lysin coated slide

i airdry the slides vertical after sectioning for 2 hours at room temp. to allow water trapped in between slides tto drain into towel. then dry at 50C oven 2 hours minimum or overnight.

If you are unable to use superfros plus continue wTransfer to buffer of choice. Continue with your proceduret slides, it will be helpful to post fix or incubate the deparaffinized slides in 4% neutal buffered formalin at room temperature for 1 hour to help adhesion of sections to slides. rinse in tap water. transfer to buffer of choice

sorry, correction on second paragraph, if unable to use superfrost plus slides, . it will be helpful to post fix or incubate the deparaffinized slides in 4% neutal buffered formalin at room temperature for 1 hour to help adhesion of sections to slides. rinse in tap water. transfer to buffer of choice. Continue with your procedure.

sorry again, the comment box seemed to mesh up on me today.

Good Luck

I agree with those who prefer SuperFrost slides!

Additionally be careful: during drying your should wape first water from the slide and then dry it vertically or at an angle position.

Clean the glass slides well and degrease with acetone. Then coat them with Poly-L or Poly-D lysine. Being Brain tissue your sections are hydrophobic. I would prefer to use collagen type I (3mg/ml) and allow to dry at RT. Gelatin is water and PBS soluble and collagen is not. You should apply sections at RT and allow to attach

Try to use Poly L slides or freshly prepared gelatin. it will give good result.

Dry section at 40C oven overnight and than put to hot plate on 50C on few second, use after cooling to room t

It seems to me the section size is too big for the slides you using which then might take too long to dry on the slides.But if you are sticking to that size then, I would live it overnight for which I too recommend using SuperFost positive slides and also I would dry the slides in an autoclave for 5-10 min at 20oC for better adhesion (making sure the sections don't start melting and this has worked with me good so far) before you dry them overnight. Hope this helps.

When you free float mount in gelatin it doesnt matter what slides are used. I use 0.5% in distilled water. Sections fall of when PB or PBS is used. Air dry, then stain. If they still fall off, then dip into fixative for 5 sec to bond the gelatin to the section, rinse then stain.

This may also happen if the sections are thicker than normal 4 or 5 microns.

Are you using a gelatin recipe that includes chrom alum? (chromium potassium sulfate). Also - 0.2% gelatin is very dilute. We used to use 0.5% and occasionally had problems with sections lifting from the glass - we increased to 1% and the problem went away. Another factor in subbing slides is how long the rack of slides is immersed in the gelatin solution. Finally, the choice of mounting buffer is also a factor - not so much the buffer itself, but the concentration. If you mount from 0.1 phosphate buffer, for example, the buffer carries significant salts that can leave a crystalline residue that can "physically" interfere with the tissue adhering to the glass. This tends to result in pockets of poor adhesion that lead to pooled counterstains rather than to total failure like you are describing.

First of all you should perform the good fixation of the tissue by formalin or any others....after the superfrost+ slide 2 hour 50 celsius.....

Hi, i agree with Seshadri that the sections might be too thick. I persenally have made the experience that slices thicker than 20µm might detach from the coated slides. I was using those superfrosted slides from Thermo Scientific, and if i remember correctly the recommand not to use sections thicker than 10 µm

Hi Rosie,

Slides will not detach if you use the following protocol:

http://www.rndsystems.com/ihc_detail_objectname_gelatin_slides_histological_tissue.aspx

Note: you can increase geltatin to 10 g/l, and keep the rest identically.

Good luck.

re: mounting buffer - I mentioned that 0.1M phosphate buffer is far too concentrated - as dilute a mounting buffer as you can use is preferable - I just used .01M PB - just enough buffer to keep the sections from osmotic shock - (distilled water will cause the sections to swell and pucker if left too long) - but mounting buffer would not cause the failure described - the gelatin solution (0.2%) is too dilute and a concentration between 0.5 and 1.0% (with 0.05% chrome alum) will work - even with sections as thick as 50µm.

The section is fine. Most of my sections for free floating is 50 um. One thing I noticed is the gelatine concentration on the slides you used. Try and increase the concentration to 0.5% gel. I believe you will be fine.

Thanks so much everyone! The protocol has been used for years in the lab without any of us experiencing problems like this, but maybe it's time to switch to something else!

Very interesting

David Finkelstein - when you say to dip the section in fix, what do you mean by this? PFA?! Or something else? Thanks!

Ramesh Nayak - the sections came from PFA fixed, and sucrose-cryoprotected brains - they were sectioned on a freezing microtome, so in that respect they are frozen sections... but they have been at RT since the IHC.

Hi Rosie,

I have used both PFA or neutral buffered formalin. The idea behind the dip is to cross link the section to the gelatin. I have performed enzmatic reations at 37degrees and the sections remain stuck. Good Luck.

Immersing sections in buffered formalin for 30 minutes to attach residual reactive sites on the section to the gelatin or PLL usually takes care of this problem, but I would wonder whether your sections were fully hydrated prior to mounting. We use a mounting medium containing up tp 20% ethanol and buffered with ammonium acetate and ammonium carbonate to the desired pH and find it important to stand sections for at least 30 minutes in the mounting medium before mounting. Also when transferring air dried sections through alcohols prior to re-hydrating or dehydrating it is important to use a 70% alcohol immersion as the first step otherwise thick sections occasionally become crazed.

In addition to my previous comment, vibrato me mounted sections of rat brains retain adherance when fixed with 4% neutral buffered formalin before start staining. This is if formalin is not issue per yor protocol. The formalin enhances morphology too. If you are not able to use formalin before your procedure, I suggest to do a quick fixation before DAb. Drain the liquid from the slides (vertical position)on top of absorbent paper for 5 min at room temp. Then develop with Dab.

It will be less complicated though, if you are able to mount sections into positive charge slides.

Caution: if using plus slides use distal led water to pick sections. Gelatin or any similar reagent will diminish adherance to plus slides. Sections will fall off if you combine positive charge slides and gelatin.

Good Luck!

Gelatin coated slides are not recomended for thick rat brain sections. You should use silanized slides instead. We have used great brands like Dako and the german W.Knitel. Other brands may eventually not work well. Mind not to air dry for more than 24 hours due section over shrinkage, specially if you are planning to use other techniques in the future like stereology or confocal images. Good luck.

It is with your adhesive used.Try with Poly-L-Lysine or AEPES coated slides.

We use 1% gelatine slides, also for 50% micron section.

Use positively charged slides

Try with Poly-L-Lysine coated slides.

As previously mentioned by a lot of people, I'd rather go for Poly-L Lysine rather than gelatin. In my hands it seems to work much better (combined with super frost slides).

Good luck

Have you tried using SuperFrost Plus GOLD slides instead of regular SuperFrost Plus slides? I've worked with 50um thick PFA fixed brain tissue slices and mount them on SuperFrost Plus GOLD slices when I perform I have to perform IHC (blocking, primary & secondary incubation, washing, etc) on the slides instead of performing IHC with free floating sections. In my experience, I haven't had an issue with the sections coming off the slides. Good luck!

We use gelatin

Rosie,

I am sorry, I may have missed this, but are you sectioning fresh tissue or tissue embedded in paraffin or OCT for crysectioning? We have found if any embedding medium is trapped under tissue upon mounting, even on Plus slides, the method of clearing the mounting medium will leave a portion of said section not attached and it is often soon lifted from the slide and floating tin the wash. Bubbles in the mounted tissue may also promote this problem, but the trapped embedding medium is usually the culprit, especially if you find your tissue curling more than usual during sectioning.

One more suggestion different to the others. I sometimes find paraffin processed brain falls off slides when the processor reagents have not been changed, especially if the wax has solvent in it. It seems to be one of the first indicators to change reagents.

Coated slide (Poly-L-Lysine slides) is the best way.

Thanks all! Really appreciate all the comments - I was using fresh tissue - stored in 40% sucrose w/azide, and they were generally frozen between cutting and staining/mounting.

I think it was just a strange batch in the end, as it was just one of those things that stopped as suddenly as it appeared, but really great to have all these suggestions :)