Hi, About  thickness, you can use 7-10 micrometer, and about antibodies, I recommend you to answer from  Dr Pariver  or Dr Koochesfahani, in research gate. The best Shiva,Nasiraei

Dear Julia,

cut cryosections 10um thick with the cryostat (let the tissue equilibrate at -18°C in the cryostat for 30 min before cutting) and let them air dry overnight at RT. Then you can store them at -80°C or proceed with the staining. My protocol: post-fix sections in 4%PFA for 7 minutes at RT, rinse in PBS 4 times 3' each. Acetone fix is also good but some antigens as CD20 and CD3 prefer PFA fixation. Then incubate the sections with 10% normal serum (raised in the same species from which the second antibodies you will use are produced) for 1 hour at RT. Without wash, incubate the sections overnight with the following purified antibodies diluted in PBS + 1% BSA:

anti-CD20 (mAb Dakocytomation clone L26, 1:50)

anti-CD3 [Rb polyclonal Ab, CD3 epsilon, Santa Cruz biotech (M20) 1:40] or mAb Dako

anti-CD8 [mAb Dako, clone C8/144B or Rb polyclonal (Pierce) 1:50]

anti-CD4 [Novocastra™ Mouse Monoclonal Antibody CD4Product Code: NCL-L-CD4-368]

CD27: [Novocastra™ Mouse Monoclonal Antibody CLONE 137b4 1:40] 

CD79a [Abcam Rb polyclonal (ab4056) 1:80]

CD138 for plasmacells [mAb BC/B-B4  Abcam or Rb polyclonal Spring Bioscience].

After ON incubation wash in PBS 4 times 3' each and proceed with incubation 1 hour at RT with the mix of your second antibodies (for multiple labelling) or with your second antibody fluorochrome-conjugated (I use second Alexa-conjugated antibodies of Invitrogen or Jackson lab). Wash well and mount sections with a Prolong anti-fade DAPI medium Invitrogen). For multilabelling use a combination of monoclonal + polyclonal  primary antibodies (!)

This methods gives a very good results in my lab.

Good luck and let me know

Barbara

Dear Barbara, Thanks so much for the protocol, that's great - I will try that. Few more question as I have an expert at hand (and in my lab or department no one is working with these things) - the spleen sections I cut now seem quite destroyed and some also float off during the washing ... anything I can do about that? Is there a part of the spleen where I see follicules better - anything I should look at during embedding in OTC when I get the tissue samples to decide which part I cut? Close to the capsule or rather further inside? Thanks so much for your help! Julia

Hi Julia,

Although I id not stain for markers you mentioned I did a lot of spleen staining and used B220 and TCR as markers. Perhaps the protocol used there might help you in case the above suggestions do not help:

I cut 7 micron spleen sections and freeze them, immediately without bringing them to room temperature. Then when I need to use them I give them 30 minutes at room temperature then fix them in 10 min in methanol at -20deg. Thereafter it is straight to PBS so that the sections do not dry out - this tended to reduce the background. I mount the sections on poly-L-lysine slides but you can also use AAS-coated slides or positively charged slides. Once the 23x5 min PBS is done, I incubate my sections in 1% BSA for 30 minutes then in primary antibody for an hour, followed by 3x5min PBS wash then 45 min in secondary ab conjugated to either Cy or Alexa fluorochromes. All my primaries are from BD and all secondaries from Jackson Lab.

If you see 'cuts' through your sections (spear-like) these are most liklely due to crystals formed if the tissue was frozen too quickly - you can slow that down by freezing your samples in isopentane bath.

Sometimes you need to test fixatives and in addition to above suggestions it is also worthwhile trying ethanol but also you can sometimes stain tissues then post-fix them once you've done your stain. This is useful if the antibody does not work on any fixatives tested.

Good luck!