HI Vipra. To keep sections from washing off they should be mounted (after the cutting step) on coated slides such as Superfrost plus or Poly-L-Lysine coated slides, then they should be baked either in an incubator or on a slide warmer at 45-50 degrees for 1-2 days, to remove all water and allow binding to the coating. Antigen retrieval shouldn't be needed for phalloidin stains as the molecule is very small and it is a stain-not like an antibody.

Edgardo, I have used both DAPI and phalloidin in paraffin tissue sections. If you fail to get good staining with Phalloidin, you may have lost the cytoskeleton ultrastructure at the fixation step. Do some troubleshooting: First test your staining method on some cell culture cells to be sure your phalloidin method and reagents are good. Then try tissues with different fixation protocols and be sure the tissue pieces are very small so that the fix penetrates quickly and stabilizes things. Formalin and paraformaldhyde only travel about 0.16mm/hour, thus larger chunks of tissue lose quality in the middle due to autolysis before the fix gets in. This is why some people add 0.2% glutaraldehyde to their formalin or PFA fix -it absorbs faster and stabilizes things. Always use about a 20 fold (by volume) excess of fixative versus the tissue piece size. Be very aware of freshness of the tissue and PFA-it goes acidic quickly and should be made fresh every week or two and well buffered in phosphate or other buffer.

Hi Edgardo

1. Bring paraffin mounted sections to water

Staining with Phalloidin

When staining with any of the fluorescent phallotoxins, dilute 25 µl methanolic stock solution (freezer) into 1000 µl (1-ml) PBS. For paraffin sections, there is no need for permeabilization as it is already permeabilized by processing solutions.

2.Stain the cytoplasmic actin for 45-60 minutes at room temperature in the dark.

3.Wash the cells 4 times with PBS (each 2 minutes)

4.Stain the nuclei for 15 minutes in DAPI (1 µl in 1-ml PBS)

5.Wash in distilled water or PBS (3 X 2 minutes)

6.Remove as much buffer as possible with filter paper.

7.Put 50 microliters of slow fade buffer/or 50 microliter of aqueous mounting medium (1volume PBS + 1 volume of glycerol) on the cells.

8.Cover with the coverslip without trapping air bubbles

9.Gently blot the sides around the cover slip from excessive mounting medium.

10.Seal the sides of the cover slip with nail polish

11.Label the slides on the frosted edge

12.Examine under the fluorescence microscope

Now you have semi-permanent preparation

Good luck

Hi Edgardo

I forgot to add that you can use immunofluorescence antigen antibody reaction (ABC method) for actin and counterstaining with Propidium Iodide

Hello guys,

I want to do some paraffin preparations with honey bee brain. In my institute we haven't confocal microscope in the moment, and my whole mount preparations are too thick to visualize at wide field microscopy.

I had the idea to include the brains in paraffin, perform the slices (ranging from 10-20um) and then do phalloidin and DAPI staining above the slices, in the slides.

Do you think it works?

Hi Livia

I tried Eosin and DAPI on testis paraffin sections ranging between 5-7 um and i got good results by fluorescence microscope.

Thank you Gaber!!

Livia, You're welcome, best regards. Also, you can go through attachment which describe a protocol for staining paraffin sections with phalloidin and DAPI.

Amazing, Gaber :) Thank you so much! Tomorrow I'll try this protocol. Sounds very great!!

Good luck and best wishes

Thank you very, very much Gaber!

Edgardo, you are welcome, best regards and wishes.

Hi, I have cells grown in a hydrogel. I have fixed and paraffin embedded the hydrogel. After sectioning and deparaffinization, I put the slide in blocking solution (1% BSA). But at this step, the whole section washes off. Do I have to carry out antigen retrieval or something? Or some other step before blocking. I intend to do an actin and dapi stain subsequently.

Kindly advise.

HI Vipra. To keep sections from washing off they should be mounted (after the cutting step) on coated slides such as Superfrost plus or Poly-L-Lysine coated slides, then they should be baked either in an incubator or on a slide warmer at 45-50 degrees for 1-2 days, to remove all water and allow binding to the coating. Antigen retrieval shouldn't be needed for phalloidin stains as the molecule is very small and it is a stain-not like an antibody.

Edgardo, I have used both DAPI and phalloidin in paraffin tissue sections. If you fail to get good staining with Phalloidin, you may have lost the cytoskeleton ultrastructure at the fixation step. Do some troubleshooting: First test your staining method on some cell culture cells to be sure your phalloidin method and reagents are good. Then try tissues with different fixation protocols and be sure the tissue pieces are very small so that the fix penetrates quickly and stabilizes things. Formalin and paraformaldhyde only travel about 0.16mm/hour, thus larger chunks of tissue lose quality in the middle due to autolysis before the fix gets in. This is why some people add 0.2% glutaraldehyde to their formalin or PFA fix -it absorbs faster and stabilizes things. Always use about a 20 fold (by volume) excess of fixative versus the tissue piece size. Be very aware of freshness of the tissue and PFA-it goes acidic quickly and should be made fresh every week or two and well buffered in phosphate or other buffer.