Hi Rania,

To my knowledge, all thicknesses that you mentioned are normally used for general IHC and also for microglia morphology studies. Thus, I do not think you could have any problem using any of them.

"The thicker, the better" I do not think is always true. Using confocal microscopy this statement works because you will be able to stack all the planes and obtain a very nice pic of the total morphology. However, if using a conventional microscope, this is not true at all, in my sincere opinion. Here you would need to do a reconstruction too if you want to see all the structure and it will not be easy.

Hope you find it useful,

Jordi.

Rania, For the purposes of your staining, the first thing to determine is the size of your target. If your target cell is 10um then a 4um slice isn't going to help you. For the purposes of staining, tissue thickness varies as the method of staining and the method of image capture vary. For example, if you are performing free floating immunohistochemistry, you can cut much larger sections as the slices are bathed in the antibody solution from every angle. However, if you are performing IHC on tissue already on slides, you will want to use a thinner tissue section. With slide staining, your antibody solution will only bathe the tissue from one side, so larger sections do not fare well. Thicker tissue sections will indeed allow you to see more cells and more processes distal to the soma of microglia, but this also depends on your microscope power. If you don't have the capability of doing Z stacks on a confocal microscope or higher, then a simple XY image can be done on thin tissue sections. There is a lot to consider. I think for your purposes (microglial staining), any of the thicknesses you mentioned will do, however, I would caution on thinner sections if you are using pre-mounted tissue slides. What device are you using to section them? What species?

The attached paper talks about brain microglial morphology so may be a good starting point.

Jordi-Thank you very much for your kind clarifications.

Sincerely appreciate it,

Rania

Hi Renee,

Thank you very much for your kind clarifications. And thanks for the paper recommendation.

My microglia are from mice species. At our research facility, we have access to confocal microscopy and I believe we can perform Z stacks.. We also have access to a freezing cryostat and a microtome.

I have been reading that with specific microglia antibodies (like this Iba1 antibody from Wako: http://www.wako-chem.co.jp/english/labchem/product/life/AntiIba1/index.htm), many have had successes with one staining technique (slide staining or free floating) over the other.

For this particular antibody I was thinking of using, I have read that free-floating is recommended. So, I had been intrigued to find out if I stain 30-40 um slices, would I be able to see more microglia processes in confocal microscopy than in a 10-20 um slice..

I am using IHC in my project so I could (1) compare microglia morphology in WT and Tg mice mutant for a gene of interest, and (2) compare th number of microglia in WT and mutant mice.

Thanks for your helpful advice,

Rania

Rania,

I have used the Wako Iba1 antibody in mice on both free-floating and pre-mounted slides and ranging from 4um to 30um tissue thickness, so you should be free to choose what works best for your equipment, etc. If price isn't a problem, I think free floating IHC on 30um slices works really well, especially if you have confocal microscopy available. This will allow you the most depth in your imaging and allow the visualization of both cell and processes. I use this Wako rabbit Iba1 at 1:500-1:750. Is your tissue fixed? Post-fixed? Cryoprotected?

Hi Renee,

Thanks very much for your clarifications.I have not yet developed an IHC protocol for my microglia staining, nor have I fixed or processed any tissue. Roughly this is what I plan on doing (I am following an Abcam guide):

1. PBS perfuse through heart followed by cold 4% PFA perfusion.

2. Harvest and transfer brains into 4% PFA and store for 1 week.

3. Transfer brains to 0.02% sodium azide in PBS until ready for tissue embedding.

4. Tissue embedding: Transfer brains from sodium azide to 30% sucrose and wait until brains sink.

- When brains sink, remove excess sucrose from tissue by blotting brain with kimpwipe. Transfer brain to centre of TissueTek well filled with OCT (tissue Tek) Freezing Compound.

- Freeze tissue by floating on methanol/dry ice bath. This should create frozen tissue blocks.

5. Tissue blocks are ready to be slices on cryostat or stored in plastic bags in -20C frost free freezer to reduce drying out during storage.

Does this seem like an appropriate technique for fixing and processing?

Would a cryostat be the suitable equipment to slice brains if I would like to do free-floating IHC staining or would you recommend a microtome?

Thank you very valuable recommendations,

Rania

Rania,

As far as your plans, I have a few comments/suggestions. First, a week is too long in my opinion to post-fix your tissue. Fixation can mask your antigens, necessitating antigen retrieval methods or making antigen retrieval impossible. In my experience, here is the way I would handle the staining:

As far as slicing, cryostat or microtome is fine. If you are cutting thicker sections and performing free-floating IHC, I feel that the cryostat is kind of overkill, but honestly this choice has more to do with which piece of equipment you feel comfortable using, although I would recommend the cryostat for thin sections and the microtome for thicker sections.

If using a microtome, you are ready for cutting after step 3 above. Once brains sink, I usually block out the section I am interested in first to minimize the time it takes to get to the area of interest. I place a little sucrose on the microtome stage and create a platform. Then I use the blade to create a flat surface on the frozen sucrose I just placed. I place my tissue block (with the side up that I want to cut first) on the flat surface that I just created. Now, you need to build a column of sucrose around the tissue. This requires some finesse and skill but the secret is to only apply a small amount at a time as the more you add, the more the sucrose tends to “spread out” instead of “building up” around the tissue. I use a transfer pipet. I fill with sucrose solution and then get the solution to the tip of the pipet. I then touch the pipet tip to the edge of my tissue and circle the tissue edge. Don’t squeeze out the sucrose, just let the surface tension pull the liquid off the tip of the pipet onto the tissue. Then wait a couple minutes for this to freeze and then repeat. Once you have completed the column all the way up the tissue, you are ready to cut. Take a few slices to make sure you have a nice, complete, flat section. Use a paintbrush (wet with 1x PBS) to take slices off the blade and place them in 1x PBS solution (or a special long-term cryoprotectant if not staining within one week). Generally, you want to collect serial sections, starting in well 1, then well 2, well 3, well 4 and so on for a total of 4-8 wells, then start back into well 1. This will give you 4-8 sets of serial sections throughout your area of interest.

If using a cryostat for sectioning, you need these additional steps:

As for the IHC protocol, I use the following for central nervous system tissue staining of Iba1:

I have attached a couple representative photos of the stain.

Good luck,

Renee

Very detailed and informative answers. Many thanks to Renee R Donahue. I think the answers help others who want to stain microglia. Just want to know which mounting media you used?

Thanks

Mohammad, there are many mounting mediums to choose from. They all work but have different capabilities. I personally prefer Vector Labs Hardset Antifade Mounting Medium with DAPI (H-1500). I like this as it includes a nuclear stain and also is hardset, allowing the medium to set, preventing the need for nail polish to seal the edges. It may take a little practice to use since it does set within a short period. I use about 1ul of mounting medium per 1mm of coverslip. I have also used prolong gold with Dapi with success in the past but feel the Vector brand is superior in ease of use. The choice can also matter depending on which type of fluorophore you are using as well as what type of light you are using to visualize. For example, vectashield does not work well with cyanine dyes and has more autofluorescence when excited with UV. Additionally, prolong doesn't work well with BODIPY dyes. I believe Prolong Glass is probably the best all-around antifade mounting medium but the cost may be prohibitive if it isn't necessary for your particular antigen or fluorophore.

A good discussion of mounting mediums as they compare in confocal microscopy is found here:

Article Quantitative Comparison of Anti-Fading Mounting Media for Co...

Thank you very much for your prompt answer and also thanks Rania Faidi for asking the question.

Hi,

Will give you a short answer for this specific staining. I recommend you to do 30um sections. Are very good for handling during cutting and provide you full microglia (including full soma and processes). I had try many Iba1 antibodies, the best one is the rabbit anti-iba1 from Wako. Do not look for more, that is the best one, and is worth you get it. Works perfectly (and just) with PFA fixed sections. You do not need to perform antigen retrieval or something like that.

Renee: Thank you very much for your insightful and informative feedback. Your stained pictures look great and I have just started practicing following your instructions with a microtome on some fixed frozen brains.

I appreciate your time and your valuable guidance.

Sincerely,

Rania