Hello there Yuka

In my experience of staining brain sections, be it cryosections or thick sections, sample autofluorescence has always been a difficult reality. Neurons are autofluorescent mostly in green and some sporadically in red and rarely also far red.

Could you please tell me the thickness of your samples and whether they are fixed?

What are you trying to visualize: Fluorescent protein without antibody staining or immunolabelled antigens in the tissue. As for confocal imaging, unless your signal stands out super brightly it may be difficult to visualize over a high background fluorescence of the tissue.

We sometimes use a quick H2O2 bleaching step even prior to permeabilizing the tissue in order to get rid of some autofluorescence. However for good antibody reactions, optical settings in a confocal microscope are sufficient to give a clean image. If the signal is small relative to the background and the sample is quite thick, I would recommend switching to a 2-photon/multiphoton microscope where your illumination is planar and do not illuminate other planes during acquisition in the sample.

Good luck

Aparajita Lahree Thank you for your comment! I added more information to my description.

Oh. Could you tell me more detail about H2O2 step? Do you have protocol that you can share?

Best,

Yuka

Mohamed Mahdy Hello :) I added more information to my description. Thanks! Yuka

Dear Yuka

We normally remove the embedding gently if possible since, we use gelatin embedding, that exacerbates background fluorescence, but about agar I am not so sure.

We generally work without blocking, just permeabilization with 0,5% triton x-100 in PBS for 1h RT and the antibodies still work surprisingly well and specifically.

If you absolutely must block then you can try 1h blocking at RT, it works ok for us but there maybe increased autofluorescence depending upon what is the source of your primary antibodies. 3%BSA worked the best for us as a blocking agent with least grainy particles or random autofluorescent super bright particles.

While imaging for finer structures, we also perform some clearing of tissue with mild reagents that do not alter tissue volume (post immunolabelling). This helps with better light penetration and less scattering, which in turn makes the signal to noise ration better.

In case you plan to use the H2O2 bleaching prior to tissue processing : freshly prepared and cold: 5% H2O2 and 20% DMSO in PBS. Treatment time may need to be optimized since we never tried with brains from 5xFAD mice which I believe have more protein aggregates. For us 5-10 min works just fine. One precaution is to always ensure that all items for the experiment are cooled to 4 degrees celsius, since you do not want to massively damage the tissue lipids in the bleaching step.

Also for 50um sections, we use 1-2 days of primary depending upon antibody quality (at RT) and 1 day of secondary antibody (RT). All washes are with 0.3% triton x-100 in PBS for 5 times (15 min per wash) after each antibody round, followed by a final PBS wash 3 times. This cleans up particulate garbage fairly well.

Try to use fresh and filtered reagents to avoid particulate matter that shine as bright autofluorescent artifacts.

Good Luck

Hello Yuka, I think you had quite some input from Aparajita. I'll just try to give you some theoretical background...

As far as I can see you are using a primary Ab raised in mouse on mouse brain. A good perfusion will of course remove most of the blood from the vessels in the brain, still when using mouse primaries you will often have to deal with the natural reactivity of the secondary Abs with the residual mouse Abs in the brain, so unless your primary is very strongly binding to its epitope you'll have to cope with many stripes that light up all over your section.

Second issue is autofluorescence. As already mentioned it is highest in the green and often found also in the red. Still if you have a weak Ab it might not be wise to use a red or far red conjugation, since the photosensors arrays on the modern digital cameras are built dedicating two photodiodes to the green and only one to red and blue, so any emission richer in the green spectrum will result double the intensity than an equal emission in the red or the blue. And confocal microscopy loses so much signal in favor of image clarity and resolution, probably more than 80%, so you need good signal and good detection, as Aparajita already pointed out.

Autofluorescence is given by blood and many other unsaturated or aromatic substances found in the cells, as well as from free(unreacted) aldehydic groups from the fixation, so you do have some chemical treatments, typically via red-ox, to quench them before you start blocking. One possibility is, as described, H2O2. This should reduce at least blood fluorescence. Other possibilities, more specific for reducing aldehydic groups from formaldehyde (bur also sugars and other substances) are 100 mM NH4Cl or 100 mM glycine, 30 min to 1 hr at room temperature. Finally if you need to be really harsh you can use NaBH4, a strong reducing agent for aldehydic, chetonic, carboxylic, and ester groups, that you have to prepare fresh at 1 mg/ml and apply immediately to your sections, 3x20 min (it will fizz evidently, producing gas).

Other issue might be the penetration of your antibody in the thick tissue. For this we use to complement the blocking and the antibody dilution solutions with 1-4% DMSO (depending on the thickness and coarseness of the tissue, in your case 1-2% should be enough). DMSO is technically non-denaturing for proteins, but in substituting water molecules alters slightly their structures due to its greater steric hindrance, producing holes in the thick extracellular matrix and fixed cytoplasm, allowing your Abs to penetrate up to 2-3 layers of cells into the slice. Just a note on the use of 20% DMSO mentioned by Aparajita, seems high, we tried it without particular success on entire embryonic diaphragms and we went back to our standard 2% provided also with the Abs, but I suppose the idea is the same.

Concerning the blocking, you tried already a few things, from very stringent (10% serum) to mild (4% BSA)... if you didn't observe any change, it means that's not your issue: if you lose signal with stringent blocking it means your Ab needs less blocking, if you have too much background with low blocking it's the opposite... sometimes you'll need to look for the compromise. But if you didn't observe differences, then blocking is not an issue for your specific Ab, and you can stay low, or have no blocking.

Last issue are the conditions at which you incubate your Ab. The kinetics and thermodynamics of Abs binding are pretty complex and unpredictable. Three factors can play an important role: pH, temperature and time. Most Abs will work optimally at any pH, but some will prefer pH 8, while others will prefer pH 7.4... if you have problems with a staining, you should try if pH makes a difference. Secondly temperature: typically one incubates Abs O/N at 4 °C. If all works well, fine, if not, try O/N at room temperature instead. Sometimes it also helps 1 hr at 37 °C. And time... although O/N is working most of the times, we have Abs that need 2-5 days, either at 4 °C or at RT to give enough signal for confocal or super-resolution microscopy.

Final point, it sounds a bit odd that a specific mouse line should behave differently form others. I would look more into your procedures. If you only tried on one 5xFAD mouse, it is possible that for example the perfusion was not very successful and you have more blood left in the vessels... Also the plaques, although insoluble aggregates, should not particularly affect a staining (although you didn't tell us what you are staining, if it is in the plaques themselves, in the matrix or in the cells!).

Good luck!

Pietro