Dear Matthijs, a follow-up since you mention "air drying and mounting in moviol". Plenty of people use Mowiol, so I doubt that is an issue. But I'm not sure what "air drying" entails, so ...

My approach is to keep the slices always submerged, as much as possible. During some washes / rinses I might remove almost all the solution, but this is just for an instant before the new solution is added, and the slice itself still retains a fine layer of solution on it during the swap. For mounting I have used DAKO mounting medium, and also Prolong Gold or Diamond. For sealing I use Biotium Covergrip.

When mounting: place a clean coverslip onto a work surface (I find it useful to elevate the coverslip slightly, for example by placing it onto an upside-down cap from a 50 ml Falcon tube). Use a transfer pipette to carefully place the slice upside-down on the coverslip. This ensures that the slice lies flat against the underside of the coverslip during imaging. The slice will be floating slightly in a drop of PBS (or similar). Use a Pasteur pipette to siphon off most of the solution without generating bubbles. Work rapidly to avoid drying out the slice. Use the edge of a fresh Kimwipe tissue to suck away remaining moisture at the edges of the slice (never touching the surface of the slice). It's okay if a bit of moisture remains, don't spend too long on this. Then add several drops of mounting medium onto the slice (how much is a matter of practice and preference - for DAKO I'd say 4-5 drops). Put spacers at the left and right edges of the coverslip: for a 300 um tissue section, I use 4 small square coverslips of #1 thickness (each 150 um thick, two on each side). Place the glass slide down onto this assembly - to avoid air bubbles, it may help to "roll" the slide down from e.g. the lower edge to the top edge (the idea is that air bubbles will be forced towards the edge this way). Cover from light and let it cure in this position (1 hour might suffice), then flip it right-side-up and store it at room temperature overnight to allow adequate curing. For Prolong Diamond, it may take up to 1 week to fully cure. Then seal the edges with Biotium CoverGrip. Once the Biotium is dry, you may optionally re-seal with nail polish. Nail polish alone should be avoided as it can bleach GFP (and perhaps other fluorophores?). Store at 4C in the dark, in a slide box that keeps the slides horizontal.

Some of these details are in the Methods of my papers:

https://academic.oup.com/cercor/article/23/8/1965/354470

http://www.jneurosci.org/content/35/7/2959.short

And reconstructions obtained from such samples are here:

http://neuromorpho.org/KeywordResult.jsp?count=28&keywords="suter_shepherd"

Maybe due to the tissue shrinkage.

Dear Matthijs,

I think the gaps, that you are observing in staining with dye-fills, are most likely due to loss of cellular membrane function and integrity. I think you are recording from the cells until they are dead or close to dying, or they are dying during the electrode pull-out. Obviously the cells are initially filled nicely with the dye at the begining, however following prolonged recordings, current/voltage steppings or simply during electrode pull-out the cellular membrane integrity is lost and the membrane is depolarised due to injury, resulting in gaps you are observing. This is particularly quite common in whole-cell configuration as giga-ohm seal required. I have discussed these issues in my previous papers, for example see my 2008 Pflugers Archive paper.

Best wishes,

Refik

Article Semi-loose seal Neurobiotin electroporation for combined str...

I personally have not seen such "breaks" or gaps in acute slice biocytin fills. Some questions and ideas that might help with troubleshooting:

How deep in the slice are these neurons? I aim for >75 um below the slice surface, better is 100 um. I believe dendrites and axons closer to the surface are more likely to be damaged, or to become unhealthy.

Try handling / transferring your slices with glass transfer pipettes (the wide, bulb-end of a Pasteur pipette, for example), instead of a brush. During e.g. washing and staining there is no need to move the slices - simply use a pipette to remove the first rinse solution and add the second solution, etc.

Try terminating your whole-cell recording earlier, after 15 minutes, at least at first for troubleshooting to see if you can get good stains that way? You can store in a holding chamber for 1-2 hours after this, prior to fixation, and see if that is beneficial.

If you think the problem occurs during the DAB staining, you could try streptavidin-conjugated Alexa instead, followed by fluorescence imaging. This works well for me for neocortical pyramidal neurons in 300 um thick slices, as described in detail here: https://academic.oup.com/cercor/article/23/8/1965/354470

Hi guys, thank you all for your time and useful answers, I really appreciate it!

Changzheng Zhang , That’s an interesting idea, I can see how shrinkage and a following expansion could result in such microscopic tissue damage… I will review our air drying and mounting in moviol procedures to see whether steps can be improved.

Refik Kanjhan, thanks for your detailed answer, I hadn’t thought along these lines yet. Indeed, cells could have been healthier in these experiments but I must say I would be slightly surprised if it were indeed all down to cell health. I am following recording procedures that have given me successful and complete fills for years:

Article Dendritic and Axonal Architecture of Individual Pyramidal Ne...

Benjamin A Suter, also massive thanks for your input. I like to go for deep neurons too, although I’m not sure my optics will permit me too patch 100um deep… I'll definitely use your tips on slice transfer next time and have indeed ordered Alexa 488-streptavidin to have a go at that. Hope that’ll also work for 350-400um slices!

Thanks again and have a nice weekend,

Thijs

Dear Matthijs, a follow-up since you mention "air drying and mounting in moviol". Plenty of people use Mowiol, so I doubt that is an issue. But I'm not sure what "air drying" entails, so ...

My approach is to keep the slices always submerged, as much as possible. During some washes / rinses I might remove almost all the solution, but this is just for an instant before the new solution is added, and the slice itself still retains a fine layer of solution on it during the swap. For mounting I have used DAKO mounting medium, and also Prolong Gold or Diamond. For sealing I use Biotium Covergrip.

When mounting: place a clean coverslip onto a work surface (I find it useful to elevate the coverslip slightly, for example by placing it onto an upside-down cap from a 50 ml Falcon tube). Use a transfer pipette to carefully place the slice upside-down on the coverslip. This ensures that the slice lies flat against the underside of the coverslip during imaging. The slice will be floating slightly in a drop of PBS (or similar). Use a Pasteur pipette to siphon off most of the solution without generating bubbles. Work rapidly to avoid drying out the slice. Use the edge of a fresh Kimwipe tissue to suck away remaining moisture at the edges of the slice (never touching the surface of the slice). It's okay if a bit of moisture remains, don't spend too long on this. Then add several drops of mounting medium onto the slice (how much is a matter of practice and preference - for DAKO I'd say 4-5 drops). Put spacers at the left and right edges of the coverslip: for a 300 um tissue section, I use 4 small square coverslips of #1 thickness (each 150 um thick, two on each side). Place the glass slide down onto this assembly - to avoid air bubbles, it may help to "roll" the slide down from e.g. the lower edge to the top edge (the idea is that air bubbles will be forced towards the edge this way). Cover from light and let it cure in this position (1 hour might suffice), then flip it right-side-up and store it at room temperature overnight to allow adequate curing. For Prolong Diamond, it may take up to 1 week to fully cure. Then seal the edges with Biotium CoverGrip. Once the Biotium is dry, you may optionally re-seal with nail polish. Nail polish alone should be avoided as it can bleach GFP (and perhaps other fluorophores?). Store at 4C in the dark, in a slide box that keeps the slides horizontal.

Some of these details are in the Methods of my papers:

https://academic.oup.com/cercor/article/23/8/1965/354470

http://www.jneurosci.org/content/35/7/2959.short

And reconstructions obtained from such samples are here:

http://neuromorpho.org/KeywordResult.jsp?count=28&keywords="suter_shepherd"

Benjamin A Suter very good point I never let my slices dry up because it can shrink/damage tissue. I usually use fluoromount-G to mount.

I then cure my slides flat, facedown, for 3-4h at room temperature in the dark, and then ~24h flat in the dark at 4C right side up (face up), and then transfer to the freezer box

Dear Benjamin A Suter and Natalia Murataeva, a massive, belated thank you for giving us the detailed descriptions of your mounting procedures. We really appreciate these insights and have adapted our procedures to match yours where we could. Benjamin, I like your method of mounting on the cover slip first, and the use of spacers for these rather thick slices. These and other small details are often lacking in officially published protocols or methods sections, which is a shame really.

Although we never let the slices dry up completely, we now make it an explicit point to ensure they are kept moist at all times before mounting. Unfortunately, we nevertheless still had a subset of stained neurons displaying these ‘breaks’ in a recent staining round. I showed some images to an overseas colleague of mine who is very experienced in these neuronal stains and he too was not familiar with this particular phenomenon… Anyway, we’re now systematically adjusting and re-evaluating every step and batch of solutions used in the protocol. For any interested readers, if we manage to pinpoint the cause I’ll be sure to post it here!