First I have to ask, are these fixed tissue or flash frozen? Fixation in PFA/Formalin give a lot of background in lower wavelengths and may be your culprit. Improper Mouse-On-Mouse technique could also be an issue, depending on your source antibodies.

The way I have previously done this is to use a previously selected zero/start point that is grossly visible while tissue is in the cryostat and take serial sections at 20/30/40 μm in individual wells that you can then number, giving you a deviation or depth from a defined point. There is at least one detailed brain atlas available online for mice that can help pick a 'start' point.

Clinically we frequently ink tissues for points of interest, and those inks are visible when sectioning. Check to see if the ink will react in your wavelength before using it to mark your samples. This lesson was learned the hard way.

Jason Rogers The tissue is fixed. I perform perfusion followed by PFA injections before obtaining the tissue.

Yes, I usually follow a similar sectioning criteria and I am looking to improve it so I will take that into consideration. Thank you!

Hi Muhammad,

I do a lot of swinging blade/vibratome sections of PFA fixed mouse tissues, that is probably also the case for you, right? I usually use them afterwards for confocal laser microscopy.

In a nutshell, what I usually do is:

-Embed fixed tissues in 4% low melting point agarose (in PBS) for sectioning.

-Cut sections with vibratome into 40-100 micron sections

-Take up and wash the sections in washing buffer (PBS containing 5% FCS, 0,2% Triton-X100 and something to keep it sterile, I use 0,01% Thimerosal w/v). If you are worried about high backgrounds, add some serum for blocking (e.g. 1-3% from the antibody-host-species, rabbit, goat, donkey etc) here. You can also store the sections in that buffer for a couple oif weeks without problem.

- Take a 24well plate and stain your sections for 4-12h in 500 microl. washing buffer containing your antibodies (typical antibody dilution range about 1:100 to 1:400). I often stain in the dark overnight for good results. If your sections are really big

- Wash sections in >5ml washing buffer for >4h, e.g. on a 6 well plate under constant agitation, I use a slow shaking shaker

- Optional: Repeat staining steps with secondary antibdodies etc.

- If I use DAPI , I usually add this for the last 5 min of washing.

- Mounting on glass slides using an antifade glycerol-based mounting medium, e.g. ProLong Gold.

My most important advice is:

Use a triton or tween-based washing and staining buffers for good results

Invest the long times for staining and washing, it´s worth is.

Hope that helps. Fingers crossed!

Urs

Urs Mörbe thank you! I usually block in FBS + PBST for one hour at RT only. Maybe I should increase the blocking time to help reduce background noise. I also feel that the integrity of the tissue doesn't give clear staining for some reason although I process brain tissue the same way you've indicated above. Confocal microscopy yields fine results but when it comes to taking really good images (Z stack for example, etc), certain axonal/dendritic stains don't give good enough images.

To reduce background noise, before to the immunotechnical, I store vibratome slice in 100% methanol (- 20 ºC at least 24 hours) and then, before to add the blocking, I use NaBH4 (0.02% in PBS; 30 minutes).

If you want to know more, let me know and I will try to help you :)

Anabel Alba González that's very interesting. I have never seen anyone store the tissue in methanol. Why is that? I'd love it if you could help me with this because I am trying to optimize the IF protocol and it's hard to pinpoint what steps to tackle.