Dear Sarka,

I have had similar problems with sections when I was doing my PhD. I have tried a number of ways, and at the end I have decided that the best method was floating sections. Briefly the steps in fixed tissue is to cut sections either using a vibratome (40 micron) or cryostat (16 micron), 3 washes in PBS, block the non-specific binding (2 hours), incubate in primary antibodies (2-8 hours), 3 washes in PBS, incubate in secondary antibodies (2-8 hours), 3 washes in PBS, and mount with glycerol. This way you never need to dry your sections and because your sections are floating, the incubations and washings are shorter, effective and work better, as well as avoiding potential artefacts such as tissue shrinkage and cracking. For details and other relevant matters see my following publications.

best wishes, Refik

Thesis Molecular physiology of ATP-gated ion channels assembled fro...

Article The two-pore domain K+ channel TASK-1 is closely associated ...

Technical Report Research Methods and Technical Considerations: Answers to ov...

Hi,

I'm not sure if I mussed that point, but don't you have to fixate (usually with PFA) your sections prior to blocking?

Well, I have my cells already fixed with 4%PFA prior cryosectioning...

We always did the fixation step directly after the cutting.. Don't know if it helps to prevent the floating of the sections from your glass slides, but a least we never had problems regarding that point...

Good luck

Try Silane (APES) coated slides or SuperFrost Plus slides (note the 'Plus')

Dear Sarka,

I have had similar problems with sections when I was doing my PhD. I have tried a number of ways, and at the end I have decided that the best method was floating sections. Briefly the steps in fixed tissue is to cut sections either using a vibratome (40 micron) or cryostat (16 micron), 3 washes in PBS, block the non-specific binding (2 hours), incubate in primary antibodies (2-8 hours), 3 washes in PBS, incubate in secondary antibodies (2-8 hours), 3 washes in PBS, and mount with glycerol. This way you never need to dry your sections and because your sections are floating, the incubations and washings are shorter, effective and work better, as well as avoiding potential artefacts such as tissue shrinkage and cracking. For details and other relevant matters see my following publications.

best wishes, Refik

Thesis Molecular physiology of ATP-gated ion channels assembled fro...

Article The two-pore domain K+ channel TASK-1 is closely associated ...

Technical Report Research Methods and Technical Considerations: Answers to ov...

Well, thank you for your kind answers and hints. I will try to extend the drying part after cryosectionning. Nevertheless when I let the sections dry at RT during one hour, the autoflorescence occurs and I can´t use the Alexa 488 as a secondary antibody anymore.

Hallo Sarka, use either superfrost plus slides, as already mensionend,  or chorm-alum-gelatine coarted superfrost slides. The duration of airdrying depends on the thickness of your sections.

If you prepare thin sections around 5µm 1 h room temperature might be enough. If they are thicker it needs more time or you have to incraese the temperature, 37°C is ok.

Autofluorescence is a problem., I know. After the IHC mount your sections with mounting media which reduce the AF like Dinaova- Mountingmedium, Vectashield, Moviol, etc. Formalin/PFA increases the AF, so do not stay longer as necessary in the fixativ. Alcohol or acetone fixations are alternatives, but it depends on the primary antibody, some like it, some don't.  If it is to hard to distinguish the immuno signal from the background I recommend to do the detection with DAB and us brightfield microscope. Best wishes, Ute

Hi, I saw you questions too late< so already Ute has relised the right decision- You need to use Polylisine coated slides or gelatin coated (!) the later is as far as I know not commercially  available  and you need to prepare it from just  usual slides BUT! they will be the best  (!)  than any comercial covered with anythin slides  to stick your sections.  

Which of these you may use can dpend on your  further purposes or  what you are going to stain?

Natalia is perfectly correct. It's not the staining routine that causes your cells to come loose but rather the lack of proper adhesiveness of the section to the slide that results in its detactment from the slide during the staining protocol. You can use polylysine but better yet would be an albumin/glycerol mounting fixative. One is commercially available from AzerScientific.com and works very well using a rather thin coating on a slide. A little on the expensive side but well worth it if it allows your experiments to succeed.

 Thank all again for you sugestions. Natalia, I have pancreatic islets.

After sectioning, mount the cells with the OCT on a 18mm coverslip. Several cuts can fit on one coverslip. Place in a 12 well dish and proceed with your staining (wash the OTC using PBS). Mount on slide following staining procedure! (I life prolong gold or diamond). As others have mentioned, Superfrost Plus (that is positively charged slides) are also necessary to prevent samples from sloughing off if directly mounting to the slides.

Supergrass glass relies on electrostaticaly loaded surface. If the glasses lost their charge (grease,  fungal growth, sometimes even water damp are negating it) the sections will not stick. I had similar problem (my slides were too old) and sections kept falling off during antigen retrieval. I found after cleaning the surface with pure acetone all sections stuck through the entire process.