Cutting problems
Last updated
Last updated
If you are having problems with cutting the first thing to do is consider the blade you are using. Particularly if you are using a thinner blade, consider changing to Campden Instruments stainless steel blades, which are verified to work well. These do not rust and are very rigid. Buy in bulk: they are cheaper that way. Expect to use a new blade for each sample but in practice you can image multiple sessions with the same blade if you wish.
Next consider your cutting parameters. In general cutting 40 micron cuts at 0.5 mm/s should work. You might need to cut a little slower in some samples: perhaps down to 0.35 mm/s. Sections much thinner than 40 microns might not sink easily and remain floating on the surface or not will away from the agar.
Vibratomes have a tendency to enter a feedback loop where they cut alternate thick and thin sections. Using the Auto-Trim feature or otherwise slowly stepping down to the final cutting thickness should help reduce the chance of this happening. Sometimes some samples are tricky and this doesn't help. If so, try cutting slower: 0.35 mm/s. If that doesn't help, then take a single section of about 80 microns then go back down to your target thickness. If this also doesn't help, consider imaging with sections 10 to 15 microns thicker.
If all sections are coming off at similar thicknesses but not intact and as a series of ribbons (as show below) then probably the vibratome has something loose.
The problem shown above is most likely due to a loose blade. When the system is not cutting, gently place your finger on the short side of the blade in the bath and push. Does the blade come away? If so, remove blade holder and tighten blade.
One form of data loss occurs when the slices fail to detach from the agar block and the flap around under the objective. This happens even though the blade proceeds 2 to 3 mm beyond the edge of the agar block (you should always ensure you set the "Cut Size" in BakingTray as a standard part of the set up procedure). The stuck slice obscures the field of view and you lose data. Things that can help alleviate the problem are:
Try using 5% agar instead of 4%
Round the corners of the agar (red arrows, below) by slicing them off with a razor blade
Larger agar blocks (e.g. those that contain 4 brains) seem to exhibit this problem less. Maybe because the weight of the slices is greater and they pull away.
You may see black tiles or darkened regions of the sample as follows.
The patches visible above originate from dura or choroid plexus that fails to cut and flap around above the sample. These occlude the sample and cause shadows.
The membranes can be removed with forceps, but they will likely return as imaging proceeds. The best way of dealing with this is to stop it happening at all. Most samples do not exhibit the problem, only some do and what these have in common is not clear. Things to try include:
Carefully remove any dura after perfusion but before you post-fix. Only do this if it is obvious what to remove. Avoid damaging the brain.
Improve your perfusion quality
Post-fix overnight in 4% PFA at 4 degrees C
Do not use old PFA.
Perhaps older animals are worse than younger ones.
Note that larger animals, such as rats and ferrets, will have greater problems associated with shadowing artifacts from membranes.
The following image shows a darker strip running down the middle.
The reason for this darkening is that in this area the brain is thicker. The blade is somehow cutting the sample non-uniformly. Often you can see this by looking carefully at the smaple at a steep angle. It is not clear what causes this. If the effect is not severe, ignore it. If it is severe or you really need to fix it, try changing the blade. If this does not fix it, the problem might be due to the brain not being held tightly in the agar.
