Hi!

What is the thickness of your slices? Maybe you could try to make them a bit thinner, around 220-230 micron...

Hi, Thanks for the response,

I'm using 350 micron slices because I want to keep the cells' morphology as intact as possible. Anyway, I believe that currently I can patch only the top ~50 microns and my labeled cells are randomly scattered, so I don't think cutting thinner would help...

Nikon made at one time a 60X water immersion lens with over a 250 micron depth of focus. Its images were very good. May be you can fine someone with this lens. If not have you thought of vital staining the cells, this may give you added land marks. In addition, can you put a florescent maker in the pipette so you can see it?

 Try to make positive pressure a little strong, then go deeper.

I agree with increasing the positive pressure, but I would try also to make at least 1-2 slices 250 micron, you'll see even better the tissue and the cells, it could be helpful anyway... And for sure if you don't see clearly the cell's membrane the recordings are susceptible to be not very stable, because it's hard to patch very good..

Try this: positive pressure (around 30-40mbar with pipettes around 3.5MOhm). Than you go back and forth and left and right when approaching the cell. This will pusch away some tissue above the cell and the DIC will improve. 

The other thing is: what are you overexpressing? Are the cells healthy?

Cheers, Milan

What vibratome are you using? The visibility of individual cells in the depth of slices depends to some extent upon the quality of  the vibratome. With a traditional vibratome we encounter a poor visibility of individual neurons below a depth of about 80  µm. With more recent vibratomes (MICROM HM650V or Leica VT1200S) the visibility is extended considerably.

^I've had good luck with the 'tunneling' technique Milan mentions. All I would add is that I've had better luck when I first tunnel with a cleaning pipette pulled a bit thinner than my patch pipette to make a space to drive through and to clean the surface of the cell.

Increasing the pipette pressure is a very good idea. Having a smaller tip (higher resistance) and a longer shaft might also help. I did a lot of blind patching deep in slices. Also I patched identified cells in thick collagen. You patch the cell based on the resistance signal. You use small, fast steps with the pipette, you see a very short, temporary decrease in the resistance (do not expect a permanent reduction in the resistance, like patching cultures or surface cells, the tissue is moving and you have the positive pressure) thus when you see the fast dip in resistance, release the pressure at once (fast) and just wait for seal formation (it takes longer time. suction does not help too much). The success rate is definitely lower than in cultures. How to find your labeled cell is the main problem. In normal case manipulator is fixed to the stage, objective is moving at focusing. In this case it might be better if the manipulator is fixed to the frame of the microscope, the stage is moving with focusing (traditional upright microscope). In this case you can place the tip of the electrode to the focus point, pull back the electrode, rise the stage with the cell to the focus point, push back the electrode to the focus point to the cell.. In this case you are moving the electrode only axially to find the cell. I did not mentioned, but in any case you better need a manipulator setup to move the electrode axially instead of XYZ.

Hi Asaph,

First, make sure your optics is as good as it can get. Use IR-DIC, Kohler the optics, I found Olympus to be superior and they have a DIC prism specifically for deep patching, etc... There is a very good box about optics in the Dendritic Patching paper in Nature Protocols (2007 I think, Davie et al).

Second, have good slices. As mentioned previously, the latest Microm and Leica slicers are far superior. Again, the Dendritic Patching Protocol has lots of good tips. And do try 200-250 um slices, yes, you may loose more of the dendrites and especially the axons, but at least you will be able to patch. One issue can be that dead cells are more scattering, thus more visible when deep. Make sure you are not poking hard and dead cells. 

Third, fill you electrode with a different dye to help visualise it. If everything else fails, do a semi-blind approach: approach visually as Peter described it but do the last 10-15 micros blind looking at the resistance. I had good success with this approach in highly scattering tissue (cerebellar granule cell layer, patching mossy fibre terminals).  

Now the 'best' solution would be to do this in vivo if you want to preserve the anatomy as much as possible. If your cells are not too deep, you could do Twho Photon Targeted Patching (TPTP, 2004? Neuron paper by Margrie et al.). If you have access to a 2p system, you can do this in slice as well of course. 

Good luck!

Thanks everyone for the detailed answers. A few more (failed) attempts led me to think that as Ede suggested, the problem is not visualizing the cells rather than the fact that they are just dead. Not sure if that's a direct result of my labeling protocol (dendritic electroporation), or maybe the labeling is biased for dead cells, or maybe it's just statistics (a lot of dead cells in the slice, so I'm getting a lot of labeled dead cells). Will try to get better slices and better optics.

Thanks again

Dear Asaph,

If these cells are labeled with a chemical fluorophore or express a fluorescent protein (e. g., EGFP) you could visualize the cells with the corresponding emission filter for your fluorophore, so you could get as close as possible. It would be easier for you if you put  some dyes in the patch-pipettes with emission characteristics overlapping those of the cellular fluorophore, so you could visualize pipette and target at the same time. If you are using thick slices (e. g., 400-500µm) you could try blind patch-clamp recording after roughly presenting the pipette to the targeted cell.

Best

Asaph:

we use IR/DIC which can get you about 150 um beneath the surface.  With labeled cells (usually fluorescent Rhodamine), we locate a "target" that is fluorescing, and carefully draw out its shape and that of its neighbors, as well as its position relative to an eye-piece reticule.

We then switch to DIC/IR, and find it,  using our drawing and the reticule for guidance; once we are confident that we have the same cell, we switch to the TV monitor to visually guide our pipette tip onto the cell.

Good luck!

Usuakky I used a blind patch clamp techniques

http://www.utdallas.edu/~tres/microelectrode/microelectrodes_ch10.pdf