Any particular reason why you use MQ water wash in your IHF protocol? I usually proceed to mount after finishing the washing and never use water.

I also would change FBS to a normal donkey or goat serum, depending on the host of your secondaries, for better blocking and less background.

Otherwise, the protocol is relatively standard. In your particular situation, I suggest shifting the fluorescent channels one step, skiing 488. I usually get worse results with the 488 channel. So you can try DAPI/595/647 (IR) to see if it helps to improve the image quality; keep in mind that 647 antibodies can only be viewed by the camera and not by the eye as it's emission in infrared.

Here is the recipe for the blocking and ab incubation buffer I have used for the last ten years, glycine is specifically added to reduce the 488 autofluorescence. I use donkey host secondaries, so change the NDS to NGS if you use goat host secondary.

If nothing else works, you can perform sequential imaging where you image your first signal, i.e., HA; remember location if your microscope has a motorized stand it might have a feature where you can return to the location of the previous image. Stain the specimens with mitochondria marker i.e. citrate synthase, and then align the nuclei in the two acquired images using software such as Photoshop or GIMP. It is a pretty annoying experiment to do and you might need to take multiple areas to find a few that match between the first and second imaging sessions, but may solve your problem.

Blocking buffer (500ml)

10% goat serum with 0.25% triton X, 0.25% tween -20, 0.1% sodium citrate, 0.3M Glycine (2.25g per 100ml) and 1% BSA in 1 xPBS

450 ml PBS

50 ML Normal donkey serum

1.25 ml Triton -X

1.25 ml Tween – 20

0.5g Sodium Citrate

11.25g Glycine

5g BSA

Allow it to dissolve and filter with fine filter paper.

Prepare 5 ml aliquots and freeze at -20°C.

Store thawed aliquots at 4°C and use within 2 weeks: discard in not clear.

Staining buffer (for incubation with ab) (500 ml)

0.25% triton X, 0.25% tween -20 and 1% BSA in 1xPBS

500 ml PBS

1.25 ml Triton -X

1.25 ml Tween – 20

5g BSA

Allow it to dissolve and filter with fine filter paper.

Prepare 5 ml aliquots and freeze at -20°C.

Store thawed aliquots at 4°C and use within 2 weeks: discard in not clear.

Thank you for your answer Anton Lennikov ! and these recipes.

No particular reason for water, it's the protocol that someone from the confocal core facility gave to my team so I started using this protocol. I have never done immunofluorescence before.

I will definitively change the serum for goat and shift the fluorescent channels.

Why do you have 2 blocking components in your blocking buffer, serum and BSA?

Also, from all the protocols I have read, some are dissolving antibodies in the blocking/permeabilizing buffer, some dissolve only in detergent buffer, is there a reason to choose one or another? Is it antibody related?

I also read that some people allow the mounted coverslip to harden overnight at room temperature (but dark) before nail polish sealing and cold storage. Is that change something? I noticed that applying my nail polish can be tricky, the coverslip moves a little, I have the feeling that my nail polish diffuses under the coverslip and is 'diluted' into the mounting medium; which could damage my staning?