You can use poly-l-lysine slides but if this method is difficult or has not enough cost-benefit , you can enhancement your slides adhesion by  applying  blood serum/plasma to them ( an easy old method) with just 2 hours incubation time. in my experience i used these slides for Toluidine blue staining (mice Hippocampus) years ago.

Thank you Johan and Mehran. I use commercial positively charged slides. I also use same procedures with my previous -trouble-free- studies. The only thing that i changed this time is using 96% ethanol (commercially obtained from sugar cane) during paraffin embedding procedure. Could it be the reason?

I think ethanol is not a cause of the failure. I recommend a retry with a new charged slides and as Johan said heating the slides before xylene.

Good luck

I agree with Johan about the overnight incubation. We regularly incubate difficult tissues for 2 hours at 65°C and then overnight at 37°C. I wouldn't expect any artefacts from that, since you are not planning to use them for immunohistochemistry. Plus, the thinner the slides, the better the adhesion, of course. Good luck!

In addition to good answers provided, i would advice you to also check the strength of your clearing agents you are using. If has been used several times----i mean long time you may think of changing it. 

What is the temp of your water bath? Perhaps your sections are adhering to the slide properly.

SuperFrost slides are commercially available; they are - according to our own experience - useful für the application of classical paraffin section staining including HE.

Hi Melike!

I was looking the answers and I think to increase the adhesion by using new charged slides and heating your paraffin-embedded sections before the first xylene, are very nice advices. Did you try with SuperFrost slides? If your answer is yes, you can try charging uncharged slides with a solution of 3-aminopropyltriethoxysilane (silane solution, 2% is recommended).

Check the temperature of the water bath as Regina says. When I process voluminous samples (in my case, collagen-based hydrogels), i use the water bath at 50°C. Initially, when i didn't use adhesion techniques (only uncharged slides), after antigen retrieval for immunohistochemistry the samples were adhered to slides in a 50%. Using adhesion methods everything is ok.

Another thing that you have to check in water bath it has contaminations or not. The paraffin adhered to water bath walls when you start is an enemy of the adhesion. Additionally, the dust is capable to interact with charged slides and decrease the adhesion force.

By the way, what's the thickness of your sections? If the thickness is too high, the adhesion will decrease, but it depends on the tissue too.

Talking about Mehran's advice, the blood serum/plasma and the egg albumin are excellent methods "to fix" the sample to slides. However, this methods are very efficient for staining methods (with H&E work very well), but if you are planning to perform an immunohistochemistry/immunofluorescence you will have a lot of problems with antigen retrieval.

Hope it helps you. Good luck with your samples.

Kind Regards.

Juan Pablo.

I agree with Juan Pablo. I use egg white to adhere sections on slides. Put a tiny drop of egg white on a clean slide and spread the drop of egg white to whole slide. The sections stick on slides well for H&E stain.   

However, egg white is rich with proteins (albumin). So, if you put to much egg white on slides, the background of your H&E stain will be red.

Dear melike

We work all the time with rat brains .In histological routine, we use common  slides for H & E and do not use any type of coated or charges. Usually what happens is that when cuts are made are bubbles or folds in the tissue  that may cause  to fall off the slide. It is also important that you leave at least two or three hours or more in the oven to make them stick.

In addition to the previous answers, just mentioning that at hospital we use the slides provided by DAKO for paraffin embeded tissue and they work very well for us.

avoid wrinkles and bubbles between your coated slides and sections...

incubate in oven overnight 

and you can proceed for H & E staining....

Most glass slides today come acid washed and clean. Generally failure of sections to stick is caused by greasy slides, so ensure the slide is really clean. The trick I was taught as an apprentice was to coat the slide with egg albumin  rubbed on the slide just before the paraffin ribbon is floated on. The albumin acts as a glue and is stain neutral.

Hi - it may be a very simple problem. Although now over 30 years since I did human neuro histology as a job, somethings do not change that much I guess. What you need to do is to dry your sections fully at 37C, best done overnight. Once fully dry, only then melt at 56C for about 30 minutes. The previous comment about air bubbles and wrinkles are right, you need to move as many of them as possible.

By all means use charged slides.

The big issue is the 'shocking' of the sections when transferring from aqueous to alcoholic solutions.

Good luck...

Cheers

Hi Hedley, I put the slides at 58 C overnight. If I change it as you said maybe I can fix. Thank you.

Hi Melike - brain is not like other tissues and needs the 2 stage drying/melting process. Please let us know how you get on.

Cheers Hedley

The charged slide is the best for what you asked.

Good luck

After fixing sections to slide with egg albumen and incubation the sections can be coated with celloidin which on drying will fix the sections to slide .It does not interfere with staining

Albumen & celloidin will work, but these are really only 'last resorts'. Still best if you attach sections properly in the first place (as described above). Regarding charged slides, I found and still find that well cleaned plain glass slides are best. In the early days of ICC we used to store slides in alcohol or acid-alcohol and clean them as we went just before their use.

Cheers

Matthew is correct, but the question was about H&E staining. As always with histology, keep in mind what the final result will be used for.

As a pathologist I want a flat, well stained H&E section with no folds, bubbles or floaters.  If I'm doing laser microdissection using PET slides, or immunohistochemistry I have a completely different set of criteria for the sections I recieve.

It goes all the way through the process. The fixation used for the tissue and the temperature of the wax used for embedding affects the hardness of the tissue, likewise the melting point (and the plasticity) of the histology wax I use, and the ambient temperature of the knife will in turn affect the ribbon quality.  Its all interconnected.

Agree with the pathologist above. Review your whole procedure. Age of the animal makes a difference as early post natal dates the brain are non-mylinated and may require a different processing. Charged slides work usually well and is cleaner than covering the slide with albumin. It is important to dry the slide well after mounting the section. Clean slide is also important. Proper mounting of the section. Of course the whole procedure of fixation and embedding also impacts on the quality of the slides.  

Agree with the others above.Fixation and processing is important hence clean alcohols and clearing agents. The albumib/ egg white should work or else try poly-l-lysine coated slides if the superfrost slides are not helpful. The egg-white will be fine for staining as its only an H&E

As a said, the only thing that i changed this time is using 96% ethanol (commercially obtained from sugar cane) during paraffin embedding procedure. I compared the sections in which 96% ethanol or absolute ethanol was used in paraffin embedding procedure and realized that sections fall off slides during H&E staining if 96% ethanol was used in embedding procedure. If embedding procedure was performed with absolute ethanol, then staining was performed perfectly.

Absolute ethanol is needed for better dehydration and infiltration.  If the tissues are not properly infiltrated during the embedding process, you will have problems with section loss no matter what type of slide that you use. 

Hi - I see the discussion has moved on to fixation & processing now. My previous answers of course are dependent on correct fixation and processing of the tissue!