Hi,
I am trying to use patch-clamp to fill medium spiny neurons (MSNs) from striatal brain slices with a dye (Alexa 568). I patch the neurons and then wait for 15 minutes for dye diffusion. After that I fix the tissue in 4% PFA (30 min) and wash it with PBS (twice, 5 min). After mounting the tissue I use an epifluorescence microscope to detect the cell I patched but by then it looks like it has blown up (all small fluorescent dots with no particular identifiable structure).
If anyone has done this before (even if it's with a different dye) can you give me some advice on this? Is there something I should be paying particular attention to?
Thanks in advance!
Hi Joao - could just be the way you remove the patch pipette from the cell. We use sharps in vivo but really necessary to remove very slowly so that the cell body doesn't get pulled off of the processes when withdrawing pipette. Same principle with patching in vitro I believe - withdraw very slowly at the end of recording. Failing that, check the osmolarity of your internal with the dye included is OK for patching.
Good luck!
John
Hello, I already did it with msn of mice, it was working. I was patching the cells more time as i was studying the cell. Of course, sometime, you have nothing as the cell died during the removal of the pipette, but most of the time, it was fine.
Hello, I have done in the stellate cells in the cerebellum. First, you need choose healthy cells, which means it should not be on the surface of the slice, try to choose the deeper one; Secondly, withdraw the electrode in proper speed, not too quickly but also not too slowly, it like you are doing outside-out patch. The last, the tip of the electrode should be smaller that you use for whole cell. But too small, which will increase the difficulty to break the cell membrane. However, I found the dye will go into the cell when you break the membrane, how the opening it is, doesn't matter. Of course, you have good opening, it will help you get good imagine. In general, I will see the whole cell structure in 1-3min, and usually I wait 10 min to make sure the dye spread very well.
Good luck!
Are you using a fixable dye? Not sure if the Alexa 568 will maintain fluorescence excitation/emission after PFA treatment, or at least it may not be as good as Lucifer Yellow for that purpose. Lucifer Yellow will withstand PFA fixation and will give incredible signal to noise for morphological analysis of your MSNs. It may not be perfect for whole cell recordings if you care about detailed electrophysiological properties, but can be titrated to a low level that is tolerable with relatively minimal effects on ion channels. Alternately you could try live cell imaging (e.g. 2-photon) after filling MSNs with Alexa dye, as this works quite nicely.
Regarding the patch pipette withdrawal and possible damage to the filled cells, you can directly view the filled cells after the detachment to see if the cell retains good membrane integrity. As long as it does not get ripped out of the slice (shouldn't happen with slow withdrawal) and as long as you don't accidentally pull out a nucleated patch, the cell should stay in good shape for fixation and post-hoc imaging.
Concerning the Alexa 594 or other Alexa dyes...there are several derivatives/variants, so my point was to verify the particular one you use will be ideal for imaging after PFA fix.
Out of curiosity regarding Ting's response: which Alexa-594 variants are fixable, and which are not? Thank you.
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