Hi Daniel,

Good question. I have used the cytosolic fluors mainly fura-2 which is loaded into the cell as the acetoxymethyl ester. The fluor has a dual excitation wavelength which means you can carry out ratiometric studies which eliminates errors in dye concentration and other factors. The acetoxymethyl ester form is transported across the cell membrane by diffusion and is concentrated in the cytosol. Esterase cleave the ester and liberated the free acid of Fura-2. Keeping the loading concentration as low as possible is important to avoid any potential impurities. Spectral recording are done continuously and I have recorded agonist induced calcium transients in epithelial cells for over an hour. There are other methods that are better suited to measuring other calcium events such as oscillaions.

Regards,

Stephen Bird

Hi Stephen,

Thanks for your answer! That sounds rather feasible.

Have you ever looked at any cytotoxicity indicators (e.g. increased apoptosis) after a session > 1hr in comparison to a control group?

I would recommend you to use Fura-2 and open the excitation every 10 minutes recording during 1 minute (if that freq suits for you) or different ut trying to avoid to excite all time. Despite fura-2 is ratiometric you will avoid too much photodamage of the cells

Limiting excitation time is a good idea and should be perfectly fine for my purposes, thanks very much Luis!

Hi Daniel,

You may have all the advice you need, but just in case here's my response to your question.

We have done quite a few long term recordings. They are not simple for some of the cell types that we study because of cell movement, which shouldn't be too much of an issue with isolated cardiomyocytes (although the cells will contract of course). The issues you may face during long term imaging include calcium indicator extrusion, indicator bleaching and photodamage. The photodamage may be caused by the illumination you use, or by ROS production via illumination of a calcium indicator within the cells. There are ways around most of these issues, but sometimes they don't work so well.

Using a ratiometric calcium indicator (such as Fura2) is ideal as a way of reducing the impact of indicator extrusion (which can be curtailed using sulphinpyrazone and other compounds) and indicator bleaching. It's not perfect though, especially if you get to minimal calcium indicator levels when the background pixel intensity and noise can become increasingly significant contributors to your recordings. To maintain calcium indicator concentration inside cells, you can do an acute loading with the acetoxymethyl ester, as Stephen said above, and then maintain a tiny concentration, say 0.1 micromolar, in your bathing solutions during your experiment. This will keep Fura2 being loaded into the cells to replace what is lost. Problem is that it could be expensive to add the Fura2-AM to your solutions. Moreover, if you lose Fura2 fluorescence due to photobleaching and have a constant reloading to compensate, the cells will have increasing amounts of a calcium buffer inside them (even if the photobleached indicator is not fluorescent).

Another issue with Fura2 is that it's not the brightest indicator with respect to fold change of the signal from Ca2+-free to Ca2+-bound. Single wavelength indicators, such as Fluo8 and Cal520, have better dynamic ranges. Also, with Fura2 you need to illuminate sequentially at 340 nm and then 380 nm to get the ratiometric advantage of the indicator. There are some fast-switching LED systems that will do millisecond 340/380 illumination (we have a CoolLED system that is very fast; microsecond switching), but most lamp and filter-based imaging systems will not be that fast.

The production of ROS and photodamage can be control for using ROS scavengers such as Trolox/vitamin E, or reducing illumination intensity/duration, but again it's not a perfect solution. It is possible to kill cells with long term illumination, as in photodynamic therapy.

Since you are interested in kinetics of calcium signals over a long period of time, I am not sure that I would use Fura2 or have a continuous recording. My approach would be to do rapid imaging of a single wavelength indicator at regular intervals (say 10 second bursts every few minutes). The calcium signals in myocytes are rapid, as you know, so continuous fast imaging for 2 hours is not that feasible. If you do your imaging using rapid imaging in bursts, you can preserve the indicator fluorescence and cell viability, and get your kinetic data at critical time points. We've used this burst 'off and on' imaging approach many times.

There is a lot of calcium expertise around you in London, but if we can help any further please get in touch.

Best of success,

Martin

Dear all,

Thank you very much for all your helpful suggestions! I actually spoke to the staff from the nikon imaging center here at KCL and they recommended using one of their fast wide-field systems for imaging calcium transients in cardiomyocytes. Since their lasers are not compatible with Fura-2, I actually purchased some Fluo-4 a few weeks ago which is now sitting in my -20 since the covid-19 lockdown.

I also decided against continuous recording and will follow an on/off approach as you suggested @Martin - to limit light exposure (and thus phototoxicity and photobleaching) and file-size. Since the experiments are dose-response experiments for a certain stimulus, I will instead do some pilot experiments at different doses to determine the time until the transients reach steady

state. For the actual dose-response experiments I will then wait the time it takes for steady state to be reached, record a few dozens of calcium transients, stop recording, increase the dose, and begin the cycle again! I will see from the recordings whether steady state has really been reached.

That should limit the total time the cells are exposed to illumination quite significantly. Adding some ROS scavengers sounds also sounds like an interesting idea. Do you do this for all your long-term sessions or only if you have the impression your cells take damage?

I'll post an update on how the pilot experiments turn out.

Best wishes

Daniel

Hi Daniel,

I think you might find that you struggle to keep fluo-4 in your myocytes. We do full concentration response experiments in native endothelial cells and found that fluo-3 and fluo-4 fluorescence was rapidly lost. Instead we use Cal-520/AM, as Martin also suggests.

Our usual protocol is a non cummulative concentration response (9 concentrations, see figure 2 in paper linked below). We record continuously for 5 minutes during each drug addition, and have a 10 minute wash after each recording.

The trick is to use very low excitation light levels. With very low light levels and a sensitive camera (EMCCD), we can image the same cells like this all day and we can perform full concentration response experiments before and after drug treatments. Another trick is to use plutonic f127 from invitrogen in your loading solution.

Best of luck with your experiments!!

https://www.researchgate.net/publication/329775445_Spatially_structured_cell_populations_process_multiple_sensory_signals_in_parallel_in_intact_vascular_endothelium

Hi Daniel,

It's really good to see the community provide so many helpful suggestions. Your question prompted a great chat thread.

We do tend to turn the illumination off during experiments if we can. Irradiating cells is not cost-free (although the damage might be imperceptible in many situations). I have seen the frequency of spontaneous calcium transients in cardiac myocytes speed up if we illuminate cells too much. I haven't thoroughly investigated how that happens, but clearly there is a radiation + indicator + cell interaction that is not ideal. We don't use Trolox routinely, but we have found it to be effective in blocking the presumptive actions of ROS in different situations. For example, if we give cells ionomycin to cause a massive cytosolic calcium signal, it will cause the cellular organelles to fragment. We can block that fragmentation to a degree using Trolox. That's not an example of rectifying an imaging problem, I know, but it shows that Trolox can affect ROS-mediated damage inside cells. Our preference is to establish the least invasive imaging method, such as on/off image capture, minimal calcium indicator loading etc.

I think the choice of wide field imaging of a single wavelength calcium indicator is a good place to start. We do that for many of our cardiac myocyte recordings. We routinely use Cal-520 and can capture images at 10 - 15 Hz for some time using an on/off imaging protocol (an example, if you are interested, is here: Article Examining Cardiomyocyte Dysfunction Using Acute Chemical Ind...

We previously used Fluo4, but have moved to Fluo8 and the Cal dyes for reasons mentioned by Nagendra and Callum above. Importantly, they are brighter and more stable. If you are intending to image at 37oC, you may find that the indicator is lost through extrusion. Many cell types are proficient at extruding calcium indicators when the temperature is elevated above 20 degrees. The options for that are to do your experiments at 20 degrees (i.e., general room temperature) or to use an ABC transport inhibitor (providing that it won't impact on signalling in the myocytes).

Sorry for the essay! No patronisation intended at all (just sharing lots of bitter experience trying to do the same as you!).

Do let us know how it goes!

Best,

Martin