...unless you’re interested in calcium. If you’re using it as a proxy for channel activation then it sucks.
There’s a tendency among those of us who study calcium permeable channels to use fluorescence imaging of Ca sensitive dyes as a way to assess channel activity. And sure, the degree of channel activation will determine the resulting fluorescence signal, and generally speaking more activation will lead to more signal. Plus it’s just easy to do: in a day you can try a huge range of manipulations, you’ll get hundreds of cells as a result (making those statistics sure seem impressively significant). Compare that with a good day of patch clamp where 10s of cells is a GREAT day. I get it. I’ve been tempted by that dark side as well.
Still, frankly, it sucks. There are so many uncontrolled and/or untested variables present in your typical Ca imaging run that if you’re making dose-response curves, or inhibition curves, or whatever, then really, you’re deluding yourself if you think you’re only looking at the channel. And don’t even get me started on those people who treat a given delta ratio as equivalent over the whole range of ratios. GRRR.
In that case, it’s time to suck it up and do the electrophysiology. Yeah, it’s hard, it’s slow, blah blah waa waa whatever. If you want to characterize a channel, do electrophysiology. If you want to see how that channel affects intracellular [Ca2+], do imaging. But then really, WTF are you doing overexpressing that channel in some poor cell line, sitting in the incubator, just minding it’s own business?