I get emails: cryostat question

August 4, 2024

Question:
I have a technician from [redacted] reaching out to ask for some feedback about your Zeiss Cryostat as they are looking into getting the same model. But as it is new to the market and not many people have this one, they would like to understand the pros and cons of the unit.

I was wondering if you had any feedback that you could provide to them in relation to any other units you may have used previously.

@Daniel Hoops Talking to [redacted] about this he suggested you had experience working on a number of different cryostat I understand it has been a while since you have worked with this one but if you had any feedback that would be appreciated.

My Answer:
Frankly, in general I try to avoid them in favour of using vibratomes, which I find easier, faster, and more comfortable to use. Vibratomes can be operated in a semi-automated way whereas cryostats can’t. Vibratomes also have the benefit of not slowly giving you frostbite on the tip of your nose.

However, there are lots of experiments that require cryostat use. I’ve used several over the years, but I can’t say I’ve noticed any differences between them or even noted what brands they were. They all seemed to function pretty much the same. Sorry I can’t be of more help on this!

If there’s anyone who has informed opinions on this and would like to share, please let me know.

I get emails: frozen edition!

April 25, 2023

Here’s another question I received recently via Slack:

Them: We have ~120 [lizard] heads that we removed post euthanasia from captive born offspring of 4 different populations... The animals were euthanised in liquid nitrogen and then the heads were persevered in 10%NBF [neutral buffered formalin] 24hrs, then 70% EtoH [ethanol]. I am using the heads for scale counts.... However, post that I was wondering if there is the possible application of looking at their brain morphology? I have had a brief look at the literature, and suspect that we probably haven’t preserved them in an appropriate manner. Similarly, I think the liquid nitrogen euthanasia may cause some issues. I thought I’d ask for your thoughts as it seems like a waste if we could apply it to brain morphology questions. … I thought there may be question around relative sizes of different regions of the brain, … etc. This is all out of my wheelhouse so I could be totally off on this one! Any thoughts on if there is anything that can be done with them?

Me: Does "euthanized in liquid nitrogen" involve freezing them [Edit: This seems obvious in retrospect, but I thought it might have been via asphyxiation]? If not it might be worth a go. The 70% ethanol is not great but 24 hrs in neutral buffered formalin might be enough to prevent the EtOH-induced shrinkage, and 70% EtOH is better than 100%... It might be worth a go for MRI but I don't think you're going to have any success with histological methods after the EtOH.

Them: The whole lizard was frozen solid in liquid N.

Me: Ok then no chance.

So this is probably the most common problem I encounter with offers of brains to study: the brains on offer have been frozen. If you think there is any chance that you’ll want to use a brain for research DO NOT FREEZE IT. Put the whole head in a sealed, humid container in the fridge, and contact your nearest local neuroscientist immediately. If you have access to formalin, put the head in formalin for 24 hours and then, again if you have access, put it in saline until you can get it to a neuroscientist. If you don’t have access to saline, still remove the head from the formalin, rinse it in water, and store it in a sealed container wrapped in paper towel moistened with water.

I remembered to post this today because of this tweet. It seems like this freezing thing is not just a me-problem!

Andrew’s Twitter is here if you have any (never frozen) extra bird brains.

If you have any comments or suggestions on what I’ve said here, please don’t hesitate to get in touch. I’d love to hear from you.

And if you have an extra bird brain lying around, please get in touch with Andrew, and DON’T FREEZE THEM FIRST!

Though my method of preserving heads maintains their usefulness for MRI and histology, unfortunately it doesn’t maintain their quality as conversationalists.

I get emails: preserving heads

April 1, 2023

This inquiry was, in fact, technically not an email, but rather a Slack message (though isn’t everything email, really?) Nonetheless, it seems like it might be useful to people, so I thought I’d post it here.

Q: @danhoops My local vet tells me he has to euthanise a [lizard] a week. He is happy to chop off the head and we could do MRI + [immunocytochemistry]. What should I give him as preservative?

A: Place the head in formalin/PFA for 24 hours in the fridge, then store in phosphate-buffered saline in the fridge. For samples you want to store long-term, add 0.05% sodium azide to the phosphate-buffered saline to prevent bacteria and fungal growth.

This answer is my opinion, I’d certainly be interested in hearing from anyone with a differing opinion.

Here is an example of Netrin-1 immunohistochemical staining in a mouse medial prefrontal cortex, revealing “rings” of Netrin-1 protein expression around cell bodies. Images such as this are artificially coloured (the camera attached to the microscope takes only black-and-white pictures) and have been enhanced in photoshop by altering their brightness and contrast. This sort of editing is permissible in scientific publication so long as the entire image is modified uniformly, i.e. the same process (for example increasing the brightness 20 points) must be applied uniformly to every pixel in the image. However, it is important to note that while this image does represent what is visible in the tissue while looking down the microscope, the editing has made the picture prettier than what one would see looking down the ‘scope in person.

I get emails: Netrin-1 staining on cultured cells

January 28, 2023

A day after posting my response to an email asking about CT scanning, I received a fresh question in my inbox asking about quite a different topic: immunohistochemistry using antibodies against the protein Netrin-1.

De : [redacted]
Envoyé : 25 janvier 2023 17:27
À : [redacted, forwarded to me]
Cc : [redacted]
Objet : advice on NTN1 ab

I am writing to seek advice regarding the choice of antibody (and method) for fluorescence immunocytochemistry against Netrin-1 in rat cultured [redacted] neurons. My student found your paper on optimizing immunohistochemistry with the Novus and Abcam Ab https://pubmed.ncbi.nlm.nih.gov/29276694/, and he also found this antibody: https://www.lsbio.com/pathplus-antibodies/pathplus-ntn1-antibody-netrin-1-antibody-aa591-604-if-immunofluorescence-ihc-ls-b5912/141319

Do you still use these antibodies, and if so which one would you suggest for our application? We weren’t sure whether citrate, SDS and/or heat would do well in dissociated neurons, so we haven’t yet tried. When [my student] tried the [abcam Netrin-1 antibody] without any specific treatment (just PFA +/- triton X), he found fairly weak staining (but does detect over-expressed GFP-tagged NTN1).

What do you think ?

Here’s my response, which probably goes into more detail than the inquirers really needed, but hopefully has some useful information for them (and anyone else with similar questions):

In my experience the abcam antibody is more sensitive than the NOVUS one, so I think you’re on the right track using the abcam one. First, a disclaimer that I’ve only ever done immuno optimization on fixed tissue, never on cultured cells, but my understanding is that, in general, it’s easier to get a signal from cultured cells than from tissue. With that being said:

Is the faint signal you’re currently getting sufficient for your purposes? Often I can turn a faint signal into something useable (for data collection, taking images for publication, etc) by fiddling with the microscope settings, in particular illumination intensity, light strength, and gain (but be careful of bleaching!) Also, signal gets stronger with increasing magnification; often I find that a signal that’s barely discernable at 5X magnification looks quite nice at 40X or 100X. Can you do your work at higher magnification?
If none of that works for you, there are umpteenth variations (and combinations of variations) of immunohistochemistry protocols that you can try. The simplest that I’ve had some success with are: leaving the samples to incubate in the primary antibody for longer (so try for 3 days if you were incubating for 2, for example), doing your incubations on a rocker, adding the serum of the animal your secondary is from to your blocking solution (but I think this is more to reduce background than to increase signal), increasing the concentration of triton-X (or tween-20, whichever you are using) in your blocking solution, and using the commercial Protein Block and Antibody Diluent solutions from Agilent instead of home-made blocking solution. In all honesty I don’t think any of these would work to improve Netrin-1 staining in fixed tissue, but because - I think - staining cultured cells is easier, they might be worth a try for you.
If you want to try antigen retrieval methods, be warned that these do tend to make samples more fragile, and I would try them one at a time before trying them in combination. The mildest one I can think of is EDTA, a calcium chelating agent. My recollection is that we never tried EDTA to recover Netrin-1 signal because we had citrate and SDS in the lab but didn’t have EDTA. Citrate is the next mildest, and SDS is the harshest. So I would try Citrate before SDS, and I would try each individually before trying them in combination.

Something I didn’t mention in the email but thought of after is that I should have sent an example protocol for antigen retrieval using EDTA, since I don’t have any personal experience with it and it’s not included in my Netrin-1 immunohistochemistry methods paper (which the inquirer linked to in their email). Here is the paper I’m familiar with that talks about using EDTA for antigen retrieval: Possible role of tissue-bound calcium ions in citrate-mediated high-temperature antigen retrieval - PubMed (nih.gov).

If you have any comments, advice, or corrections on the advice I’ve given here, I’d love to hear from you, so please get in touch!

Here is a lizard brain stained with… something… I suspect it’s stained with Methylene blue. Also, don’t remove the brain from the head before CT scanning! Stain and scan the whole head to avoid brain deformations during extraction.

I get emails: brain staining and fixation

January 27, 2023

These arrived in my inbox from a collaborator recently:

Q: So… I [understand that I] need to stain the brain to estimate the volume or I will not see anything at all [in my CT scan], but I would like to know if there is any staining technique that could be used for both CT scanning and making slides,

A: So recently a paper was published saying that there’s no problem with doing microCT scanning followed by de-staining and then proceed as normal with histological staining. So stain away!

Multiscale imaging of the rat brain using an integrated diceCT and histology workflow | SpringerLink

Q: Also, is there any better (and healthier) way to fix the brain instead of formalin that may lead to the same results?

A: Do you mean like, safer and healthier for you? Not that I know of. 4% paraformaldehyde (PFA), which is 10% formalin without the stabilizing agent (usually methanol), is the standard. You should use freshly made-up PFA, not formalin.

There are other fixatives out there, but I believe they are all more dangerous, not less dangerous, than PFA. I’ve never looked into it very deeply, though, because PFA is just what everyone uses.

If you are more knowledgeable on these topics than I am and have any comments or corrections, please get in touch, I would love to hear from you!