Hi Anu,

Certain tissues have strong autofluorescence that can be decreased using proper agents. It would be good to give more information/details about the staining procedure and the type of tissue you are using. Otherwise, it is difficult to give a proper answer.

Hi Milica,

I used 5µm thick paraffin sections of mouse liver, colon and lung.

The procedure I used for staining as follows;

1. Heating slides @60°C for 1 hour

2. Rehydration in Xylene and grades of ethanol

3. 1x PBS wash (5 minutes)

4. 3%H202 for 10 minutes, then wash in PBS

5. Antigen retrieval by immersing slides in buffer containing EDTA and 0.05% Tween 20 (pH-9.0) for 40 min @90°C

6. 1%BSA in PBS for 2 hour in RT, then wash with PBS (3 times, 5 minute each)

7. Add Primary antibody (anti GFP mouse Mab) 1:1000 dilution in 1% BSA, overnight incubation@4°C.

8. Wash with PBS (3 times, 5 minute each)

9. Add Secondary antibody (goat antimouse IgG) with FITC (1:32 dilution) for 30 minutes @RT

10. Wash with PBS (3 times, 5 minute each)

11. Stain with Hoechst 33342 nuclei stain for 1 minute

12. Wash with PBS

13. Mount with Everbrite hardest antifade mounting medium

I want to know whether I need to dry the slides before mounting

Hi Anu,

To mount your tissue sections after the staining, first it is important to aspirate the excess of the washing buffer and than to proceed with the mounting. I would not leave the section to dry on the room temperature for a lot of time. Just aspirate the buffer and dispense approximately 10 - 20 - 50 uL of mounting medium on the tissue section. The volume of the mounting medium depends of the size of the tissue section and the coverslip you are about to use. Coverslips should be cleaned with a paper towel before use. Apply a continuous strip of medium along the center of a section. Do not put a drop of the medium in the center of the section. Gently and slowly place the coverslip on the section (at a 45 degree angle) and allow mounting medium to spread over the entire section. Applying a lot of mounting medium is not recommended. Gently press down the cover slip after mounted to expel the formed bubbles if they are present. Incubate your mounted sections overnight at room temperature on a flat surface protected from light, to allow the formation of a permanent hard seal. Regarding mounting Everbrite hardset mounting medium, before mounting your slides, warm the medium to room temperature and mix well by gently swirling or inverting bottle before use; avoid vortexing or rapid shaking, which may introduce air bubbles.

Regarding autofluorescence, as I mentioned before, this is a frequent problem and depends of many factors such as the fixation protocol, type of the tissue, preparation (frozen or paraffin-embedded tissue), etc. If you are experiencing autofluorescence, you should try to reduce it with a proper agent.

Here are some tips that could be useful in your specific case:

Evans Blue

A 50 ug/mL solution of the dye, Evans Blue, for 10 minutes, has been used to combat green autofluorescence contributed by catecholamine in paraffin sections, works by absorbance quenching. [De la Lande IS, Waterson JG. 1968. Modification of autofluorescence in the formaldehyde-treated rabbit ear artery by Evans blue. J Histochem Cytochem. Apr;16(4):281-2].

Pontamine Sky Blue

The dye Pontamine Sky Blue has been shown to be superior to Evans Blue in combating catecholamine autofluorescence in paraffin sections with minimal interference with FITC

labels. Use 0.05% Pontamine sky blue in PBS containing 1% DMSO. [T. Cowen, A.J. Haven, G. Burnstock. 1985. Pontamine sky blue: A counterstain for background autofluorescence in fluorescence and immunofluorescence histochemistry. Histochemistry 82: 205-208].

Photobleach Reduction of Autofluorescence

Autofluorescence of paraffin sectioned tissue can be reduced through photobleaching with mercury arc lamp or UV lamp, for up to 48 hours (M. Neumann & D. Gabel. Simple method for reduction of autofluorescence in fluorescence microscopy. J. Histochem & Cytochem 50(3): 437-439), though the authors did not examine recovery of autofluorescence. Reduction of autofluorescence in paraffin sections via photobleaching outperformed Sudan Black B, CuSO4 in ammonium acetate buffer, sodium borohydride, and Pontamine Sky Blue.

Best of luck!

Thank you Milica for sharing the detailed information.

Anu

You are welcome, Anu!

Paraffin sections are much worse than cryosections for autofluorescence, so you might want to go with cryosections if you aren't already. Sodium borohydride washing (3 x 10 minutes in VERY freshly-made 1 mg/ml sodium borohydride in PBS) after rehydration will significantly reduce autofluorescence, for both cryosections and paraffin sections. I have also noted that use of limonene solvent (such as Histoclear) instead of using xylene is another way to reduce autofluorescence, for paraffin sections.

I'm not familiar with that mounting medium, so I don't have advice for dismounting. If it is an aqueous-base mountant, you could put it in warm PBS for a half hour to redissolve it and have the coverslip slide off.

If the bubbles appeared over the tissue, only after the mountant cured, that's a sign that there was air trapped in your section prior to mounting, and when the mountant cured it compressed the tissue, causing the air to come out as bubbles. The way to prevent this from happening is to pull a vacuum on the sections while they are in a wash step (with the slides in a Coplin jar in buffer, you can put the jar in a vacuum chamber, perhaps for 15 minutes or so). This gently pulls out the air from the sections. THen mount.

Here is a trick to avoid bubbles. Put a drop of your mounting medium in the center of the coverslip (not the slide!). If you use a long coverslip, you can use a line of mounting medium instead of a drop. Put your slide upside-down and bring it down on the coverslip until it touches the drop of mounting medium and attaches to the coverslip. The medium will spread gradually to the edges of the coverslip without bubbles. Then you can turn the slide upside-up and press it gently with a paper wipe to absorb any extra medium from the edges of the coverslip.

We use Prolong Gold from Invitrogen-Molecular Probes as mounting medium. However, we use frozen sections, not paraffin.

You may want to switch from FITC to a more potent fluorophore like Alexa Fluor 488 (green) or Alexa Fluor 568 (orange-red), or one of the Atto dyes. They will outshine the autofluorescence.

Thank you Jason and Juan Carlos for clearing the doubts with detailed information.

Dear

You can try 70% glycerol or RIMS (Refracting index matching solution) or FocusClear (CelExplorer Labs) as a mounting medium immunofluorescence imaging

I usually use "VECTASHIELD Antifade Mounting Medium with or without DAPI to mount. Use half a drop on the sample. Hold the corner of the coverslip with a fine tweezer. Make sure that u don't press it too hard to break the cover slip. Lower the coverslip and u can see the edge touching the vectashield. Do it slow and gradual and therefore any bubbles if formed will move outward. Drop the cover slip only when u see no bubbles. Keep the mounted slide in 4oC for a few hours before imaging.