With a spinning disk confocal you probably have to stick to dyes excitable around 488 nm and the emission is around 520-530. The choice of dye depends on the available emission filter set, but I guess that will not restrict you much. I have experience with Fluo-4, which is a better choice than Fluo-3 as it does not compartmentalize so quickly. I have no experience with Cal-520, but its data sheet states it is an improvement to Fluo-4 in S/N and also compartmentalization (stay longer in cytoplasm). With Fluo-4, in my experience it is possible to measure cells up to an hour after dye loading, after which time the dye is mostly incorporated into vesicles or extruded from the cells. So, both dyes can be used in your experiments, check the filter set of the microscope which one fits better. Don't forget that you need the "AM" version to penetrate into cells. In this regard, I've seen a difference, as Fluo-4 AM at 1-2 uM final concentration works fine (30-40 min loading time is OK), whereas the suggested working concentration for Cal-520 is in 5-10 uM (so it might be more expensive to use). Another issue that these dyes are not ratiometric like Fura-2, so if you need to calculate intracellular free Ca2+ concentration than you should produce a calibration curve. Even if you just compare the fluorescence intesity values, I suggest you to check cell's well-being status at the end of the experiments like adding 100 uM ATP to the cells, followed by a positive control (adding 10 uM ionomycin) to see maximum fluorescence, then by a negative control (adding 1 mM EGTA) to see the minimum fluorescence values.

Best, Karoly

Dear Dr. Karoly Liliom,

thank you for explanation and sharing your experience. It is great. Thank you for that. I was considering Fluo-4, however my experiment requires more time than 1 hour of measurement. Anyway, I decided to use Calcium-Green, AM. Good point is to check the loading time. Honestly, I do not know how long the cell will hold the dye. Probably, I have to find it out empirically. I do not need to measure Calcium concentration so far, so there is no need for ratiometric indicator. I am interested in Ca2+ influx/release during contraction only. I will try to perform positive and negative control of min/max fluroescence values, great idea. Would you recommend me to use Probenecid for dye loading? For instance, Cal-250 loading protocol advises to use it, whereas Calcium-Green recommends Pluronic detergent addition only.

Thank you.

Best regards,

Juliana

Dear Juliana,

When we performed intracellular Ca-mobilization assay in adherent HEK cells a few weeks ago, we used both pluronic acid (PF127) and also probenecid during dye loading. We made stock solution as 250 mM probenecid in PBS, and 20 m/m% pluronic in DMSO. We loaded 2 uM Fluo-4 AM for 30 min in HBSS buffer with 0.02% pluronic and 1mM probenecid as final concentrations. Our microscope (Nikon Ti2-E) is equipped with an incubation chamber (Okolab stagetop) to keep cells alive during measurement. I would say under this circumstances only the compartmentalization/excretion of the intracellular dye (most likely by ABC-C and/or ABC-G type of active transport pumps) limits the experiments - the first hour is great, maybe the second hour is acceptable, depending on the cell type and the dye used. Probenecid can inhibit ABC-C type transporters, so it likely facilitates effective dye loading...

Best,

Karoly

I have Pluronic acid, unfortunately not the Probenecid reagent. From your answer Probenecid seems to be very effective to use along with the dye. I will consider it carefully. So you incubated the cells for 30 minutes? In the manual of Calcium-Green 1, AM there is 20-60 minutes range, but I would go somewhere in between, like 45 minutes? Is there any time limit which is better not to exceed?

Thank you Dr. Liliom

Earlier I used also only pluronic. You may either reduce the loading time via increasing the concentration of dye-AM, let's say you can double the recommended concentration. It will be more expensive but you can start the measurement sooner - so the time for the free dye to be compartmentalized can also be reduced. By increasing the loading time compartmentalization may start earlier, especially without probenecid. I suggest you to do a few pilot measurement to optimize the concentration and loading time to have minimum half an hour measurement time after, e.g. uniform intracellular labeling is achieved for the free dye. In my experimence, fluo-4 AM at 1uM loading conc and 40-45 min loading time was successful. Calcium green 1 AM. should behave similarly...

Will do. I will mix the probe from 2 mM stock solution (2 mM stock solution prepared in anhydrous DMSO) with final 0.05% F127 Pluronic acid in ratio 1:1 (product sheet recommendation) and dilute to the final working solution of 1 uM dye concentration (the lowest recommended probe concentration of Calcium-Green 1), then let the dye incubate with the cells for 45 minutes. As a working solution buffer I would use serum-free DMEM cell growth medium. It is solution which cells have been in last 24 hours and I commonly measure cells in it. Once I order Probenecid, I will dilute it in the PBS to get 25 mM stock solution and on the day of experiment I will take appropriate volume to maintain 1 mM of Probenecid in DMEM (working solution). This same buffer will be used for dilution of dye and F127.

What loading temperature works best for you, please? Do you have an experience that decreasing loading temperature reduces indicator compartmentalization?

After dye loading I plan to gently wash the cells once with fresh medium containing 1 mM of Probenecid by pipetting only. Is that a proper way?

Thank you Dr. Karoly Liliom.

Your outlined protocol should work fine - and you still can play around the concentration and the loading time if necessary. Normally, we keep the cells in the incubator (so 37°C,...) during loading. Keeping at lower temp might slow compartmentalization, but the cells would not be happy out of the incubator for long. After loading a single wash by pipetting is recommended - take care not the detach the cells. Good luck :-)

Thank you very much Dr. Liliom. You helped me a lot to understand and feel more confident in the method.

Best,

Juliana