Hi Andrew,

As you say, IHC with fresh frozen brains is rarely as nice as perfused free-floating tissue, but still often possible depending on your antibodies and structures of interest.  Even though I know very little about the primary you mention.....on the points I CAN comment on, the main differences are as follows:

First, most people will typically use coated or charged (e.g. superfrost plus) slides.  If you just use plain glass, the sections will not survive the staining (mangled and detaching from the glass).  If it's already sliced, maybe do 1-2 test staining to make sure your tissue survives the process and that antibodies are working well.  If they aren't sticking well, make sure that the tissue has at least 20-30 min to dry onto the glass at room temp after you thaw the slides, this may help.

Second, depending on whether you snap-froze the brains, left them for a day or two in sucrose to cryoprotect, or maybe gave them a 24h postfixation in 4% para before freezing, you may need to think about fixing your tissue.  This usually means having the slides in PBS for an initial wash, doing 10-20 min in para at room temp, then washing again before any staining.  

Third, there are fancy systems that you can purchase (or borrow) to limit the amount of antibody, which can be important if you are using high concentrations.  My low-tech solution to this has often been to cut up "coverslips" of parafilm.  This way, you can add 200-400ul of your primary solution per slide, then gently lay the parafilm on top and do your incubation (while the parafilm keeps the section covered in antibody solution without too much evaporation etc).  When you remove these, it helps to add 1-2 ml of PBS around the edges and then lift it off with tweezers or fingers.  Generally, these are all laying flat in a humidified slide box (humidification usually just means that PBS is covering the bottom of the box).

http://www.pharma.uzh.ch/dam/jcr:00000000-22c3-f196-ffff-ffffddd37f89/protocole_immuno_rev2015.pdf

Fourth, you will need a new titration run or two to optimise concentrations, they will likely be different than your free-floating protocol.

Finally, good luck!

michael

Hi Andrew,

As you say, IHC with fresh frozen brains is rarely as nice as perfused free-floating tissue, but still often possible depending on your antibodies and structures of interest.  Even though I know very little about the primary you mention.....on the points I CAN comment on, the main differences are as follows:

First, most people will typically use coated or charged (e.g. superfrost plus) slides.  If you just use plain glass, the sections will not survive the staining (mangled and detaching from the glass).  If it's already sliced, maybe do 1-2 test staining to make sure your tissue survives the process and that antibodies are working well.  If they aren't sticking well, make sure that the tissue has at least 20-30 min to dry onto the glass at room temp after you thaw the slides, this may help.

Second, depending on whether you snap-froze the brains, left them for a day or two in sucrose to cryoprotect, or maybe gave them a 24h postfixation in 4% para before freezing, you may need to think about fixing your tissue.  This usually means having the slides in PBS for an initial wash, doing 10-20 min in para at room temp, then washing again before any staining.  

Third, there are fancy systems that you can purchase (or borrow) to limit the amount of antibody, which can be important if you are using high concentrations.  My low-tech solution to this has often been to cut up "coverslips" of parafilm.  This way, you can add 200-400ul of your primary solution per slide, then gently lay the parafilm on top and do your incubation (while the parafilm keeps the section covered in antibody solution without too much evaporation etc).  When you remove these, it helps to add 1-2 ml of PBS around the edges and then lift it off with tweezers or fingers.  Generally, these are all laying flat in a humidified slide box (humidification usually just means that PBS is covering the bottom of the box).

http://www.pharma.uzh.ch/dam/jcr:00000000-22c3-f196-ffff-ffffddd37f89/protocole_immuno_rev2015.pdf

Fourth, you will need a new titration run or two to optimise concentrations, they will likely be different than your free-floating protocol.

Finally, good luck!

michael

As Michael already said, it depends very much what already happened to the tissue between death of the animal and the staining. Fixation might change epitopes, also non-crosslinking fixations like ethanol, methanol, acetone an so on! If the tissue is unfixed and remains unfixed you don't need an antigen retrieval. So it might be good to get more information on this. And think about a good control to be sure that your antibody is working and the signals you get are specific.

For staining: I guess you already use a hydrophobic pen (from Dako or similar) to draw circles around your sections? This allows you to save antibody solution and to put several sections on one slide. Instead of putting parafilm on top of the slide as Michael suggested I heard about using a cover glass for the same purpose. It works good if the whole slide ist incubated with the same antibody solution, but they might mix if using different solutions on the same slide.

For the humid chamber it doesn't matter, if you put PBS or just water inside - the salts in the PBS won't evaporate and don't do any help for creating a humid athmosphere. I normally put paper towels on the bottom of a box and let them soak in tap water. Then I put some kind of tubes as "holders" on top on which I place my slides.

Hi Andrew,

I agree with what Michael and Vivica have suggested.

By the way, how thick are the sections that you are collecting?

I'm new to doing IHC and started off by collecting 10 micron thick frozen sections, which is quite thick for this kind of work apparently. Higher background is the biggest issue I guess.

It seems to me that people who are comfortable with an antigen retrieval procedure are more likely to do paraffin sections instead of cryo. I guess that's mainly because the morphology of the sections can be improved with paraffin. Does anyone have any thoughts on that?

Hi Andrew, I agree with all guys and I can recommand what they have written. My protocol for fresh frozen nativ cryosections:

Put 10-20µm sections on  superfrost plus slides  (4 sections/slide is possible), let them air dry until they become transparent.; put them in a slide box and store them at -80°C until you will do the immuno.

Immuno: Take your sections of interest out of the -80°C ; let them thaw at room temperature until the become tansparent (approx. 1 hour); put them in 4%PFA  in the fridge (4°C) over night; after the fixation 2-3 short rinses in destilled water,  3 x 5 min PBS; 70% Ethanol short rinse; Ethanol abs. plus H2O2 (1:100)  4°C (fridge) for 20 min; prepare this solution in advance so it has the correct temperature; 70 % Ethanol short rinse; PBS 3 x 5 min; now start with your immuno protocol. I recommand  to run a dilution raw and a negativ control to check the optimal dilution of your primary antibody as well as the quality of your dection system. If the result is not suficient that means the staining is to pale and/or the concentration which you need is to high you have the possibility of differnt enhancementt techniques. 1. DAB plus heavy metals like Ni-ammoniumsulfate or cobald chlorid;  when you are using the Anti-rabbit-HRP antibody you can do an enhancement with biotinylated tyramide (CSA System DAKO; Adams JC; J Histochem&Cytochem 40:1457-1463 (1992); or you switch from 2-step to 3-step (biotinylated Anti-Rabbit , Avidin-Biotin-complex;Vector) or you use Streptavidin-Peroxidase and enhance this with Anti-Streptavidin and again Streptavidin-Peroxidase.  I hope it will help you. If you have any questions about the different enhancement techniques, let me know. Good luck and best wishes, Ute

Dear Andrew,

I agree with the other answers.  However, in my experience, it is easier to use the parafilm coverslips than the glass coverslips for incubation.  You can roll the parafilm a bit during application so that you don't get bubbles underneath (and therefore no antibody!), but with the glass you have to be much more careful (just like coverslipping with mountant).  Not to mention that the parafilm is a lot cheaper than good glass coverslips.

I have had to stain many sections at one time for tract tracing, but for me, it was less about efficiency and more about making sure that each slide had equivalent time for enzymatic activity.   I titer my antibodies, if possible, so that I react my positive control slide for 10 min in DAB and the negative control slide has no background after 10 min.  So, since I can rinse the DAB off of one slide per minute, I react my slides in batches of 10, with DAB added to a single slide removed from the rinse jar every minute (I hope that makes sense).  That way, no slide spends longer than 10 minutes in DAB.  This is particularly true for me when using the enhancement techniques.  In my one-person-by-hand operation, that means that I usually can process batches of 30 to 40 slides. 

Part of how fast I can rinse my slides is dictated by the way our EH&S has us collect our DAB waste.  If you can rinse your slides more quickly, then you can process larger batches.   

Good Luck!

Jill

Hi Everyone, thank you so much for your answers and advice! My first time doing it with parafilm didn't work out well (primary antibody exposure wasn't good enough), but I think I need to refine my methods and try it again. 

Evan, my sections are 20 micrometers of rat brains, but I typically do 40 micrometers for free-floating. I know people do thinner sections, but I haven't had trouble so far with 40 micrometers (or even 50 micrometers for human and monkey brain tissue).

Jill, do you use the ready-to-use DAB from vector labs or do you make your own DAB solution using the tablets?

Thank you again everyone!!!

Dear Andrew,

I've been working with frozen sections from tumor specimens and I usually cut at 4micron thickness only. My protocol for doing IHC on frozen sections are quite straightforward. I've never used S100 but this protocol works for my routine antibodies. 

1) After sectioning, I'll collect the sections on positively charged slides.

2) Before staining, I'll thaw the slide at room temperature for a few seconds, which is just enough to let the OCT melt away.

3) Next, I fix the tissue section using 70% ethanol for 30 seconds, and then rinse adequately with PBS. If you want to stain immediately then you can proceed with the rest of the steps such as peroxidase blocking, primary antibody incubation, HPR-polymer, DAB and counterstaining. For this protocol, I don't need the antigen retrieval steps. If you are not going to stain immediately, you can store the section at -20C after you fix them using 70% ethanol. This preparation works well using manual and also automated IHC stainer (fixing in 70% ethanol).

However, I would recommend that you also prepare side-by-side another sections which serves as a negative control tissue. Basically, you just prepare another section from the same tissue, and instead of putting the primary antibody, you substitute it with the antibody diluent or PBS. This will help in identifying the background staining of your specimen so that you have more confident in your interpretation.

Most importantly, you have to check that your antibody can work on frozen section either as recommended by the manufacturer or you need to use proper positive and also negative control tissues to validate the antibody. 

Hi Andrew,

I have been doing IHC on fresh frozen tissues for many ,many years ...including using 20um thick sections. Forget about any need for antigen retrieval - that is reserved for paraffin sections and will only cause your tissue to come off of the slide. Although I haven't used S100 I have used many other antibodies. Hopefully your sections are mounted on Superfrost Plus charged slides.

My 'go to' fixative for fresh frozen sections in acetone/100% ethanol (3:1) ice cold for 10 minutes. That works for most antigens and of course fixation is essential for any IHC work. 10% neutral buffered formalin (pretty much the same as 4% paraformaldehyde), or even 2% NBF is another option, but may over crosslink if you fix too long. Again, I have used these on sections and fixed for 10 minutes.  Coverslipping during your incubations is iffy, sometimes the  solution can wick off the slide or you damage the sectionwhen placing or removing the coverslip, etc. and since you are doing 160 slides it is pretty time consuming.

Here is my standard protocol I start with...and thne adjust accordingly. Do not let the sections dry once you begin - that will likely cause non-specific background staining.

Fix sections in ice cold Acetone/ETOH (3:1) for 10'

Wash in PBS 3 x 3' RT ( between the 2nd and 3rd wash I use a hydrophobic pen (I.e.Pap )en) and draw vertical lines on either side of the section about 0.5 cm outside the tissue)

H2O2 peroxidase block, 15' RT

Wash in PBST 3 x 3' RT

Block w/ modified goat serum blocker ( or a commercial Casein blocker) 1 hr. RT

Incubate optimally diluted primary antibody O/N 4ºC

Wash in PBST 3 x 3' RT

Incubate  Goat anti-mouse (or rabbit depending on primary antibody species) IgG HRP polymer (i.e Dako Envision+ or BioCare Medical Promark micro-polymer secondary)15-20'  RT

Wash in PBST 3 x 3' RT

Develop w/ DAB  ~1-5 ', monitor under microscope for optimal time

Wash in tap water, counterstain w/ hematoxylin, dehydrate, clear and mount

I am a fan of BioCare Medical products, If you don't want to mark the goat serum blocker, use there Background Sniper of Punisher.

Normal Goat Serum Blocking Solution (modified from ihcworld.com)

       5% goat serum (blocking)

       1%BSA - IgG & protease free (stabilizer)

       0.1% cold fish skin gelatin (blocking)

       0.1% Triton X-100 (penetration enhancer)

       0.05% Tween 20 (detergent and surface tension reducer)

       0.05% sodium azide (preservative)

       0.01M PBS, pH 7.2,  qs to 1L

        Mix well and store at 4 ºC

As others have stated a negative reagent control is critical. most people omit the primary, but non-immune rabbit serum (in your case) diluted to the same concentration as you determine to be optimal for S100 staining is better.

Hi Brett,

Thank you for sharing your protocol.

You mentioned that you have used up 20 um thick sections for IHC.

With sections more than 5 um thick, are the 1 hr blocking step and overnight incubation of the primary the key to obtaining good signal:noise?

Thanks,

Evan

Hi Evan,

It generally helps when working with polyclonal antibodies made in rabbits and other species like goats which can be 'dirty'. Remember, with polyclonals you can have a higher chance of off target or non-specific binding because your primary antibody is a collection of antibodies directed against different epitopes on your antigen (protein) of interest. Some of these epitopes may be also be partially expressed on other cells  in the tissue sample, but hopefully with lower binding affinity to the polyclonal primary than the affinity for your true target antigen.

If one incubates overnight at 4C you titrate your primary to a higher dilution  and generally weed out those lower binding affinity antibodies while retaining more of the specific high affinity antibodies. it is also thought that the lower temperature slows down antibody-antigen interactions which again favors binding of high affinity. While it is not always necessary incubate overnight, it can also be a convenience.

Blocking solution's/times etc. really depend on what you sees on your pilot experiment when you are optimizing the protocol and looking at the negative and positive controls. Sometimes it may have more to do with the blocking solution then the length of time. I have used 1 hour, 30 minutes, and on occasion used the blocking solution as the antibody diluent. If you use a serum based blocking solution it needs to be made with species serum of the secondary antibody, NOT the primary...that is crucial. Many use serum-free blockers which are usually casein/BSA based at different %'s, and these are commercially available.  

Also, don't feel committed to a single primary antibody. My colleagues and I often test antibodies (different clones/ against different epitopes) generated against a particular antigen obtained from several sources and find a wide discrepancy if their effectiveness...from total crud that doesn't work at all (i.e Santa Cruz), or yes it's OK, but we still can't get rid of the non-specific staining without losing the robustness of the specific staining,  to VIOLA - this is the greatest! So don't be afraid to look at multiple suppliers.  And always look for rabbit MONO-clonals which are nice when working with rodent tissues and you want to avoid mouse-on- mouse issues, etc. 

Most of what I have stated pertains primarily to fresh frozen specimens. My experience has been that if you can process to paraffin, you get much better morphology and the processing helps alleviate a lot of background issues as long as you have a good primary to start with.

Last - anyone starting in immunohistochemistry needs to read the Dako IHC handbook, it explains a lot!!!

http://www.dako.com/08002_ihc_staining_methods_5ed.pdf

And get away from the old avidin-biotin technology and embrace the secondary antibody conjugated polymer technology... IMO

Thanks Brett for these great suggestions.

In terms of thickness, I guess thinner sections are always going to allow for better penetration of fixative and antibodies and better overall morphology.

With thicker sections though (>5 um), which steps of an IHC protocol do you think need to be optimized or given more attention?

Evan,

Honestly, I don't worry about penetration rates with slide mounted sections or about section thickness when your talking 5-20 um. To me they are all very thin versus the thicknesses some people use for free floating IHC.

Working in a peroxidase system you have to be sure the endogenous tissue peroxidases are killed. If you see rbc's in vessels turning brown with DAB you probably are not doing sufficient peroxidase blocking.  Any DAB staining seen on a control slide where you did not apply the primary is either from endogenous peroxidase or biotin (if you use avidin/biotin detection) that wasn't blocked - or-  your secondary HRP-conjugated antibody is binding to some IgG in the tissue if you use a peroxidase-HRP polymer detection system

To answer your question, optimization of the primary antibody dilution is my primary concern. I try to titrate the antibody out as far as I can while maintaining a robust signal. This can help reducing or eliminating non-specific background staining.

Antibody data sheet suggested dilutions are only a guide, in some cases we've diluted an antibody 10-fold over the suggested dilution from the supplier...others we needed  to use more concentrated. If background is still a problem after optimizing the dilution then start looking at your protein blocking step or add blockers to the primary diluent.  It is also important not to wash between the protein blocking step and the primary antibody incubation step. Also maybe shorten the secondary antibody incubation step.

One problem with IHC is that there is little or no standardization...different labs have different protocols for the same antibody, there is a variety of detection and amplification systems, chromogens, etc. All lot of this stuff needs to be empirically worked out by the researcher.