Does anyone have exemplary images of what good, healthy spinal neurons looks like that would be suitable for electrophysiological recording? As a newbie to spinal e-phys, this has proved to be one of the hardest parts, that is, identifying which cells are good to patch.
I know which ones I shouldn’t patch:
These are big and swollen and you can see the nuclei. So avoid these.
But finding the good ones is a bit more challenging. The features I’ve been advised to look for are cells with smooth surfaces, no internal darkening. Can anyone post some exemplary images of what good dorsal horn neurons look like and also maybe comment on the type of optics you use?
I’ve attached an image of what some normal spinal tissue looks like. You are right, you really should be avoiding any cells that look big and round, often with a visible nucleus - the classic fried egg.
The ones to look for, you can see a few in my image should have nice edges and hold their shape - when I’m teaching ppl to patch I usually tell the ‘jelly beans’ are good ones to start with. Having a pipette with positive pressure in the slice will also really help you identify the healthy cells with crisp edges. I’ve sealed onto a cell in the image here, there’s another good one just to the left up, left and below, a bit further right next to a nasty looking round one, and on the right below the myelin running horizontal.
Hi @MGradwell. Thanks. Actually, I used another paper from your lab as the basis for a lot of my buffers this summer:
Overall it was pretty good and standard (sucrose cutting, and then regular aCSF).
Some specific points though about the slicing and lead-up.
Do you do a full agarose embedding with melted agarose? Or do you make a cut-out from a solid block of agarose to hold the cord in place while slicing? And do you make any modifications to keep the cord from sliding out of the cut-out?
Do you ever do transcardial perfusion with ice-cold aCSF or do you go right into the cord? I was using transcardial perfusion so that I could make the cord cold quickly, and that afforded me time to dissect the cord in the animal while still in the body. I know some people just cut out the whole vertebral column and put it in ice/slurry sucrose solution and then dissect in that, but I find it easier to take the vertebrae off when the cord is still in the intact animal.
Up to what age animal do you use the sucrose cutting method? For animals older than 5 weeks, I know some have tried the NMDG recovery method (https://www.brainslicemethods.com/). Have you tried this or do you always use sucrose?
The buffers will be the same throughout all the group’s papers. We don’t use the agarose technique. I’ll point you toward another of my supervisor’s papers - http://jn.physiology.org/content/99/5/2048.long.
You will see in that paper that for transverse slices we use a styrofoam block cut at about the length of the cord, sitting up. We find the cord ‘sticks’ to this pretty well and allows for slicing. For this method it is important to put the thicker end of the cord down, or you have a teetering tower situation.
We don’t use transcardial perfusion. We dissect out the column as you mention and remove the cord in a sACSF slurry.
I did a 2.5 yr old mouse the other day - though this is at the extreme end we typically don’t use particularly young mice, ranging anywhere from 1 month - 2 yrs on avg. I have not tried the NMDG method, always use the same cutting solutions.
Thanks again @MGradwell for the information. The styrofoam sounds interesting. I’ve never heard of that. I’ll give it a try. So just a block of styrofoam behind the cord to give support? No cut out? That’d be an easy solution, easier than the agarose block.
Also good to know that you can use older animals with sucrose and still get good results.
