Hi all,
I'm currently working on getting calcium imaging working on my rig. Currently, I'm using 50 ug alloquots from thermofisher and following their electrophysiology protocol which includes the addition of 44 uL DMSO and 9 uL pluronic acid. I then apply the Fluo-4 directly to the slices and incubate for 40 minutes. I was wondering if anyone else had experience with this, as their electrophysiology protocol is for cultured cells. Under the scope, I do have cells that are labeled, but I cannot seem to get evoked responses from stimulation or stimulation mixed with a high potassium solution. I've found a couple of JOVE videos discussing the methods, but most are using the Fura indicator.
Best,
Brandon
Brandon, one of the most difficult bits when labeling slices using AM indicators is ensuring that they remain healthy. This part is much easier when working with cultured neurons.
Also, keep in mind that using Fluo-4, you would expect labeled neurons to be very dim, due to their low calcium. Continuously bright neurons will most likely be dead. Using a ratiometric indicator such as Fura-2 this is easier, as all labelled neurons will be visible. So it is possible that you are actually labeling neurons, but not clearly seeing them. A destructive means of testing this is to apply a high potassium solution to the slice after labeling and see if cells light up.
One way to approach this is to inject the AM-labeling solution into the slice. Stosiek and colleagues described this approach in 2003:
http://www.pnas.org/content/100/12/7319
You may also want to have a look at Rafael Yuste's large body of work on this. He has quite a few method papers and chapters describing the approach.
I have tried a number of solutions myself. In my hand, the easiest was to prepare the Fluo-4 AM with pluronic solution as described by Thermofisher and dilute that into your ACSF. Apply that to the slice while aggressively superfusing the chamber with carbogen. This will at least partially replace the perfusion (which should be switched off for this procedure) and keep the slice close to the right pH and oxygenation.
This method could be enhanced by actually holding the chamber with the slice on ice during the initial incubation. This will slow metabolism, helping reduce damage to the cells. It also nearly stops the esterase activity, allowing the AM-ester to diffuse deeper into the slice without being de-esterfied. But I do not have anything quantitative to back that up.
As always with these things: your mileage may vary. And it will depend heavily on what brain region you are working with as well as how old the animals are. While a lot of work has been done on understanding how to better target AM indicators, a lot of this still remains a bit of a mystery.
Hi Christian,
Thank you so much for the extremely detail oriented reply. I did try the high potassium bath with no increase in spontaneous activity. We've used this prep for other ephys recordings with no problem, so I'm guessing my problem is in the incubation chamber. We are also going to play with concentrations a bit as we are applying Fluo-4 at a higher concentration than originally thought, so we are lowering that a bit as well.
Brandon
For test purposes I would suggest trying other indicators as well. OGB-1 AM tends to behave differently than Fluo-4 AM and uses the same excitation wavelength.
Part of the behavior of AM esters when loading cells can be explained by partial charge distribution in the molecules. Thus, indicators with a striking difference in partial charge distribution will load different cell types with different efficacy.
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