Dear Laura, I agree with all  who recommmand  several coating techniques to keep your sections on the slides .  If you are using paraffin sections with a thickness around 5µm chrome- alum -gelatine coated slides or so called loaded slides (that means superfrost  plus slides) are suitiable. After mounting your sections on the slides let  them air dry or at 37°C over night. Then you can start any kind of staining. Referring  to your H&E stain protocol: After rehydration in Xylene and descending alkohols rinse short time in destilled water  before you put the sections in Haematoxyline for 5-10 min. Then rinse  again in destilled water and then blue  the section under running tap water for 10 min. Normally it is not necessary to differenciate the sections in HCl-Alcohol. A short check under the microscope would be helpful. Then continue with you Eosin staining as you have discribed it. Play  with the Eosin staining time to optimize the result.  In the attachment you will find my Cresylviolett protocol. Good luck and best wishes, Ute

When you say you lost your sample, do you mean it came off the slide? Are you using coated or charged slides? If you are, then it may be beneficial to pipette your stain (and washes) onto your sections instead of dipping the slide into the stain. You may need more washes if you do this, since you are using a smaller volume.

For the H&E, you may have left the sections in stain too long. The timing is something you need to determine empirically. I don't have much experience with H&E, but I know this can be an issue with other stains. Luckily, with cresyl violet, you can de-stain in 70% ethanol if you leave them in too long.

From my experience with floating sections, this seems like a good protocol to start with: http://web.qbi.uq.edu.au/microscopy/wp-content/uploads/2011/pdfs/Guide/Cresyl_Violet.pdf.

Again, the staining time will have to be determined empirically.

I generally find that CV is more likely to lead to sections coming off slides than H&E, especially when transferring slides from room temperature acid alcohol/room temperature water washes to boiling CV solution. Therefore, I'd be inclined to investigate why the sections are dropping off during H&E staining.

Are you drying the slides at low temperature (like 30oC) after mounting them on slides? Wet sections tend to drop off more easily.

How thick are your sections? Thick sections tend to come off more easily. I find that >10um are considerably more difficult to keep on.

Some cases do come off more easily, for a number of reasons. Are the sections that are coming off from the same case? If not, then I would suggest its a protocol issue. If it is just a particular case then I have found that being extra gentle when putting slide racks into solutions has been somewhat effective but not perfect.

I hope this helps in some way!

Daniel

Thank you guys for your answers.

Effectively, I mean that the sample comes off slides. Tim what do you mean by coated or charged slides? I use slides coated with paraffine, if you mean so. 

The protocol I follow starts with a deparaffinization at 100ºC, followed by a deshydration in xilol and ethanol (100%, 90%,70%, five minutes in each one). Then, I wash the sample in tap water and then I start with hematoxylin staining (4 min), again tap water, and then some dips in HCl-EtOH 96%. Again tap water and then eosyn staining (one and a half minutes). Finally, I dehydrate the samples again with xilol and ethanol but in the reverse way (70%, 90%, 100% and xilol). Then I use DPX mounting medium and I left the samples drying at room temperature (25ºC more or less).

The thickness of the samples is between 5 and 7 um (so they are not supposed to be dropped off the slide). I don't really know...I will test different minutes in eosin, as maybe 1'30'' is too much.

Thank you again.

Thank you Faiza

Why do you coat your slides with paraffin? Instead coat your slides with gelatin or Albumin. Then you dont need to keep the slides in 100ºC. Instead keep the slides in three changes of xylene which is a clearing agent. Since you coat the slides in paraffin the tissues detached from slide when it is treated with 100ºC and xylene. 

And for staining procedure follow the link

http://web.qbi.uq.edu.au/microscopy/cresyl-violet-staining-nissl-staining/

Dear Kathivaran,

thank you for your response. I coat mi slides with paraffin because is the usual procedure at the laboratory. My other labmates work with liver samples and  they always introduce the sample in formol for 24 hours and then it in ethanol, for finally including the sample in paraffine, and it always work for them. 

I certain agree with you about the treatment at 100ºC, as it maybe is too hard for the samples to stay in the slide.

Dear Laura Díaz Marugán,

We r using gelatin coated slides in our laboratory in which we never faced any such issues till date. It can even withstand heating upto100ºC for immunohistochemistry procedures. it is cheap and easy to coat as well.  

Good luck

Dear Laura,

for me personally it is inconceivable that

- as you are working in a Barcelona-Espana institution (and I know that people there are well educated and informed) -

EITHER

your laboratory (as you state in the previous post No 7) FAILS the 'state of the art ' mounting of "paraffin-"sections for either histological or also IHC-staining of "deparaffinized" sections in the end   [because use of "coated" or "charged" slides - be it poly-L-Lysin, APES, gelatin, albumin, egg white, chrome-alum gelatine, etc. etc. - in modern laboratories is quite frequent, usual and "state of the art / technology")

OR 

You  (and - as you write -  then also your lab mates ....

Citation: My other labmates work with liver samples and  they always introduce the sample in formol for 24 hours and then it in ethanol, for finally including the sample in paraffine, and it always work for them. End of citation )

are not quite informed or were not instructed correctly about the right / correct processing schedule of histological specimens in order to get finally well adhering, well stained and flat sections without wrinkling on slides      [there are a lot of good and well written Handbooks of Histological Practice and Techniques!)   -  see also section ii) below ].

Your statement  in post/reply No 3 reading: 

"The protocol I follow starts with a deparaffinization at 100ºC, followed by a deshydration in xilol and ethanol"...

is WRONG and 'scientifically/technically NONSENSE' because:

i) you never won't < melt >your paraffin sections at 100°C in an oven or a hot plate  to get rid of the paraffin in the sections - o. k., YES, you COULD DO THAT! -  but nobody professional does it ....    due to heat-induced antigen-denaturation   ||see  *) below  ||     [and this matters with regard to not only the proper IHC location by specific antibody binding, but also when staining classically with H&E, or other histological or histochemical staining methods)    

||   ')||  YES- after warming your fresh paraffin sections slightly in the waterbath (30 - 60°C for stretching sections, end temperature depending on the study task), then - after mounting your paraffin sections to the slide - you want to get rid of excess water from the slides (e. g. by wicking with filter paper) - then you'll put the slides into an oven or on a "hot plate" at 30-37(40)°C and let dry / adhere the paraffin sections (as others said) slowly (e.g. also over night). 

ii) HONESTLY spoken -  learn your lession:  

concerning: 

"followed by a deshydration in xilol and ethanol (100%, 90%,70%, five minutes in each one)." (ok- since you are not native English speaker or writer I am petient knowing that sometimes there are difficulties in keeping apart native tongue from foreign vocabulary or terms)...    Nevertheless ( and I know that all other posters know that for sure) let me tell:

It is known already that XILOL

(I know that the Spanish PC-or Laptop keybord/keypad often lacks a "Y" (XYLOL in german, XYLENE in English), so either you would have to use the ASCII keyboard or use the function "paste  (special) symbol" for the letter Y

definitely is immiscible with / insoluble in water, so by all means it CANNOT be a "deshydration" (dehydrating) agent     but instead,      chemically as a dimethylbenzene (derivative of benzene /benzole) xylene  is an aromatic hydrocarbon mixture consisting of a benzene ring with two methyl groups, and therefore an organic solvent (which solves e. g. also paraffin wax).

iii) in fact - when you deparaffinize your sections on the slide, you "DISSOLVE" the solid paraffin matter of / within the section / tissue with the organic solvent xylene (e. g. at least 2-3 times for 10-15min. each at RT for 3-7 µm sections)  then you change the organic (water-insoluble) solvent to a solution / fluid which is miscible with an organic as well as miscible with a hydrous/water soluble fluid (this is 100% EtOH / alcohol which should be WATERFREE! )...... and only but then you "RE-HYDRATE" (instead your telling us to deshydrate = dehydrate) using a descending alcohol series (96%), 90%, (80%), 70%  (perhaps then also 50%) to distilled or tap water or an appropriate buffer solution.

iv) As others have mentioned it might be that your sections had not been dried thoroughly after sectioning to the slide, but for me there also is possibility of

- poor initial fixation of the brains you process (it would be interesting to be informed about the species),

- poor dehydration and / or clearing. As you know: Brain tissue contains much lipid. If lipid isn't fixed well

OR

lipids ( at least a quite high proportion) due to short dehydration and clearing time schedule will not be eluted from the tissue properly

[as this usually is the case during processing for histology into paraffin (-/+ high% alcohols) and during CLEARING with / IN xylene]

 this eventually will hamper firm adherence of sections to the slides.

This -besides some technical difficulty (hot dye solutions etc.) - also could be a cause of mostly detaching or loss of sections you were faced with.

I really do hope that you don't think I am offending you or offend against good manners here...  

I had to add this my two cents honestly for the sake of

Dear Wolfgang,

Thank you for your answer and wasting you time in order to add your two cents for the sake of the scientific equality. 

I find all your advices very interesting. I will tell them to the rest of the lab in order to improve the technique so we can be at  the state of the art. 

I also find that you could use minuscule instead of using capital letters, as you probably know (as well informed man) that in written communication it represents like shouting. Related to how to write, It is also great to remind me some English language tips (as "xilol" and "deshydration") but we speak about a scientific technique, so it seems an out-of-place comment.  Anyway, thank you for your, as you have said, spelling "lession". 

Thank you again for the comment, I will apply your suggestions. 

Dear Ms./Mrs. cand PhD Laura Diaz Marugán,

you are welcome and I shall remind your words for my next post. 

Dear Laura, I agree with all  who recommmand  several coating techniques to keep your sections on the slides .  If you are using paraffin sections with a thickness around 5µm chrome- alum -gelatine coated slides or so called loaded slides (that means superfrost  plus slides) are suitiable. After mounting your sections on the slides let  them air dry or at 37°C over night. Then you can start any kind of staining. Referring  to your H&E stain protocol: After rehydration in Xylene and descending alkohols rinse short time in destilled water  before you put the sections in Haematoxyline for 5-10 min. Then rinse  again in destilled water and then blue  the section under running tap water for 10 min. Normally it is not necessary to differenciate the sections in HCl-Alcohol. A short check under the microscope would be helpful. Then continue with you Eosin staining as you have discribed it. Play  with the Eosin staining time to optimize the result.  In the attachment you will find my Cresylviolett protocol. Good luck and best wishes, Ute

Dear Ute,

thank you very much. I let the slides drying, but maybe wansn't enough time. 

thank you again =)