Mn should completely quench Fura-2 at these concentrations, so my suspicion is that you are not picking up any Fura-2 signal - it is overwhelmed by GFP. You could check this by instead using a high concentration of extracellular EGTA (mM) in the absence of calcium. This should show a reduction in signal if you are indeed picking up calcium, especially after digitonin. Are you using "classic" GFP or eGFP? The latter has a longer max excitation wavelength (488nm), and for 340/380 nm excitation would show considerably less emission (see http://www.biotek.com/resources/articles/green-fluorescent-proteins.html) . We have used this relatively successfully in the past to look at calcium in eGFP tagged cells. The alternative of course is to use a longer wavelength dye - as these are cells you could get away with a non-ratiometric dye such as calcium green.

Mn should completely quench Fura-2 at these concentrations, so my suspicion is that you are not picking up any Fura-2 signal - it is overwhelmed by GFP. You could check this by instead using a high concentration of extracellular EGTA (mM) in the absence of calcium. This should show a reduction in signal if you are indeed picking up calcium, especially after digitonin. Are you using "classic" GFP or eGFP? The latter has a longer max excitation wavelength (488nm), and for 340/380 nm excitation would show considerably less emission (see http://www.biotek.com/resources/articles/green-fluorescent-proteins.html) . We have used this relatively successfully in the past to look at calcium in eGFP tagged cells. The alternative of course is to use a longer wavelength dye - as these are cells you could get away with a non-ratiometric dye such as calcium green.

If you are using eGFP, then there should be little interference with the Fura-2 signal, which is excited by UV. In addition,since Fura-2 is a ratiometric dye, increasing calcium should result in a drop in 510 nm emission when the cells are excited at 380 nm light. Thus if you are looking at qualitative changes in calcium, then GFP fluorescence, even if you can pick it up at 380 nm excitation, shouldn't change as the calcium goes up or down. The digitonin/Mn quench idea probably doesn't tell you much since the digitonin permeabilization probably liberates all the free Fura-2 in the cell. Thus, whatever fluorescence you are seeing most likely results either from the GFP or background cell fluorescence.

We routinely make Fura-2 based calcium measurements of cells expressing GFP fusion proteins and have had no problems with interference from the GFP emission. Our GFP has an excitation maximum of around 470 nm. What type of GFP are you using?

Thank you for your answer! I am using cloned protein in pCAGIG vector with my cDNA and IRES-EGFP that seems maximum emission at 510. I would like just to probe that GFP is not affecting changes in Fura-2 emission

You can use manganese as a surrogate for calcium. Mn2+ totally quench the fluorescence of FURA 2. See the work attched. Gistavo Benaim

You can calculate how much fluorescence is contaminated from GFP to Fura-2 and Fura-2 to GFP with "linear spectral unmixing". Briefly, you can check the how much % of fluorescent contamination from GFP to Fura-2 channel and Fura-2 to GFP channel, separately. Then you can make simultaneous equations. You can apply the equations to your real data to subtract contamination. Please see the papers below.

1. Niino, Hotta, Oka, Simultaneous Live Cell Imaging Using Dual FRET Sensors with a Single Excitation Light. Pros One. 4, e6036 (2009)

2. Zimmermann, T., Rietdorf, J. & Pepperkok, R. Spectral imaging and its applications in live cell microscopy. FEBS. Lett. 546, 87-92 (2003).

As Jeremy said, Mn should effectively quench the fura2 fluorescence. The trick is to make sure it is getting into cells. We routinely use ionomycin to get Mn into cells. A23817 can be used, but is is fluorescence in the UV range, so bromo-A23187 could be better. Digitonin will permeabilise your cells and potentially let lots of constituents (including fura) out. I would avoid digitonin unless you have no joy getting Mn in. If the fura2 is trapped in organelles, you may have trouble quenching it. Calcium indicators can accumulate in organelles and potentially complicate cellular responses.

We have also had problems mulitplexing GFP and fura2, because GFP has a small excitation peak around 380 nm. In this situation, the prescence of GFP increases the signal obtained with 380 nm excitation (thereby making 340/380 ratio from fura artificially low). I think this was first described in a paper from Stephen Bolsover, who also offered a simple mathematical method for unmixing the signals. I hope this help.

Best wishes, Martin

To really get an accurate measurement, unfortunately fura-2AM should not be used with GFP or really RFP reading to get an accurate measurment of cytosolic Ca transients. However, my suggestion instead of quenching the dye is simple. Added a 0 Ca + 1 mM EGTA to a Normal Tyrodes or Saline solution. You can look at the effect and see if it is measuremable by changing bath concentration. The reason I say this is because fura-2AM is esterified and sticks in the membranes of the cells which you are transfecting with the GFP tagged protein. My opinion would to use either calcium orange or calcium crimson depending on your how fast you are scanning and depending the rig in which are using. If you are not using high speed (2ms), line scanning or 2D imaging, either off these would work and you would not have to worry about having to quench your signal which I suggested for the most accurate results.

I apologize for the post above, it should read if you are NOT using line scanning and/or 2D imaging, calcium orange or calcium crimson would be appropriate. Even if you use ionomycin, which I have used myself to try calibrate with fura-2AM, in non transfected cells it simply does not work because of the mechanism of fura-2AM. Fura-2AM is esterified and the mechanism of the way it works is that it gets cleaved and stuck into the membrane of the cells it is exposed to. So even, if you used ionomycin to poke holes in the membrane you will get as I would imagine residual calcium fluorescence contaminating your recording.

Sorry William, but I completely disagree with your post. 'Stuck in the membrane'. I really don't know what you mean by that. The ester groups that allow Fura2 to pass acorss membranes are cleaved inside the cell, thus yielding the free acid (calcium-sensitive) form of the indicator in the cytosol. Calcium Orange has a very poor dynamic range. I would stay well clear of it.

I would also be curious if people in the community have had success with Calcium Crimson. To my knowledge, that dye also has a poor dynamic range though if people have had success with it, I'd like to know as I have about 20 tubes of it sitting in my freezer and I don't know what to do with it!

If you use probes.com, calcium crismon and calcium orange are perfectly fine DEPENDING on the rate in which you scanning and also rig in which you are using. We had an Ionoptix rig that you could use it fine because it was 1D imaging and we were just measuring cytosolic calcium. Now when I worked with a Zeiss Five Live doing 2D calcium imaging or line scanning (2ms-8ms/scan) you will need a dye such as flou-4AM.

The calcium orange and calcium crismon were made to offset the problem the original questioner asked. As Martin pointed out above, they do not have a great have a great dynamic range, HOWEVER they were designed for this purpose. They were to designed for slow scanning imaging where you have GFP tagged proteins and using a 1D imaging rig.

Another option if you are using a slow calcium imaging rig, my next best suggestion is to transfect with GFP and then with the same GFP with your tagged protein. You can study the transients and calcium puffs to see if your protein is making a difference.

People can disagree with me that is fine. If you would like to see an example, check out my paper on my profile.

Best,

Wil