Load the cells at 37C, it is much more efficient. If you want absolute Levels of calcium, you have to calibrate your experimental setup by chelating all calcium with EGTA (to substract background fluorescence) and known amounts of calcium (make sure to look at the Kd of the indicators). You would first record the fluorescence signal for your cells (your "concentration"), then permealize them with ionomycin and add EGTA (background value) and finally add known concentrations of calcium in extracellular buffer (calibration values). You can then fit the values and determine your absolute intracellular calcium concentration.

Also have a look here:

http://www.lifetechnologies.com/de/de/home/references/molecular-probes-the-handbook/technical-notes-and-product-highlights/loading-and-calibration-of-intracellular-ion-indicators.html#head2

Good luck!

I agree with Maximilian in that 37 degrees is better, not all calcium pumps work at room temperature so your results will be further from the physiological truth. Furthermore it is always preferable to use ratiometric calcium measurements over single wavelength as your results will not be impacted by variable dye loading and extrusion. There are some compounds out there that are designed to stabalise this but they are from from 100% effective. Microscopy is as you say much more reliable. I have used both plate readers and microscopy and even though the latter takes far longer experimentally, the results can't be matched and you will end up doing fewer experiments and have much less variable data. If you have a good microscopy set up I would chose that way every time.

Have you looked into Indo-1 am? In my hands, it was much better than fura or fluo, and it takes into account the background of the Ca2+-unbound dye.

Véronique

David, i understand that you used ratiometric measurement (Fura-2AM) in microscopy,

how do you do that as you have to excite the indicators at 340and 380nm ? I'd be thankful if you can share your protocol.

37 degrees for fura 2 AM is standard in microscopy. The ratio ext. 340/380 and measurement is for single cell analysis more efficient compared to the single fluorochrome (Fluo 2 or 4), because the normal basal level of the same cell type are different. The absolute measurements of calcium need a large protocol for the interesting cells.

The procedure followed a described technique  with fura-2 as indicator. Briefly, cells in Ca2+ free DPBS were loaded with the calcium-sensitive dye fura-2 AM (to 2 μM) in the dark (30 min with gentle shaking). After washing, the calcium signal (fura-2 ratio, excitation at 340 nm and 380 nm, emission at 510 nm) was recorded using a high-resolution residual-light fluorescence imaging system (Till photonics, Martinsried, Germany). Cells were viewed continuously under the microscope (IX70, Olympus) in an open dish incubation chamber (ΔTC3, Bioptech, Butler, PA) at 37°C.

Indeed, you need a dual excitation hyperswitch for FURA2 measurements, however for cellular (in case of cardiomyocytes diastolic) Ca2+ these measurements are far more superior to Fluo-4 measurements. This is, however, not an easy choice to make, since Fluo-4 has the advantage of a much better loading process and a better signal for kinetics. If you do not need to excite your cells (like cardiomyocytes), then Fura2 is likely the better choice because the dye is ratiometric and multiple experiments can easily be compared. If you need kinetics, e.g. Tau or RT80% of Calcium transients, then Fluo-4 provides a better signal and is also more feasible if you don't have access to a fluorescence microscope with dual excitation capabilities (e.g. Ion Optix setups). Fluo-4 can also be "pseudo-calibrated" to receive actual Ca levels and not just fluorescence values. In my opinion, the Fluo-4 calibration is easier than the FURA2-calibration, which takes some experience. Either way i would recommend a fresh dye for each experiment, stocks are not a good choice for Fluo-4 or FURA since micelles between the essential pluronic acid and the dyes are destroyed by low temperatures.

Special dye packaging would be my choice (eg. http://www.lifetechnologies.com/order/catalog/product/F23917): one vial should be enough for about 16 separate experiments with 250 µl dye each. Also, please note that Fluo-4 is a light Ca buffer, which affects shortening experiments. In this case, I would recommend Fluo-3, which is less of a Ca buffer.

Some resources for Fluo4: "Chemical and physiological characterization of fluo-4 Ca2+- indicator dyes Gee et al"

Have you considered Flow Cytometry?

I use that when measuring Ca2+ in human eosinophils.Instruments with vacuum-system works, as well as the "old" BD FACSCalibur. There might be others, but I have only experience in a few different instruments.

If you use FuraRed, AM and Fluor-3, AM from Life Technology, you need an instrument with only one 488 nm laser and two detectors (Fl-1 and Fl-3)

There are nice chapters in my book, one by Bruton and Westerblad

http://www.amazon.com/Calcium-Signaling-Advances-Experimental-Medicine/dp/9400728875

Hello,

In my opinion, plate reader and microscopy basically answer different types of questions. Microscopy gives a more fine and precise profile of the calcium response. Also it gives the shape of the calcium response at the single cell level. In this case, you have to measure many cells to reach a reasonable n number. However, plat reader may measure the calcium response in a cell population, which could be thousand of cells. Of course, it also needs to be confirmed in 4 to 6 more independent experiemnts.

In both cases, you can and it is better to use ratiometric dye such as Fura-2.

37°C is better since it is closer to physiological conditions.

I hope this help.

Good luck.

Abdallah

Dear Slim,

It appears is a few other people have answered your question to me already, you need to have a fast switching light source to change between the two wavelengths often in the order of tens of switches per second. I agree with the other suggestions and the protocols used are quite standard in their nature. I disagree with Abdallah in part as microscopy can also be used to measure a population of cells, it just depends on the objective you use. The optimal loading temperature will vary with cell type, I study airway smooth muscle cells and it is common to load at both RT and 37 or even RT, then 37 degrees. I would take into account how many cover slips/plate wells you want to do per session which will depend on the length of your protocol and try to avoid having the cells sitting loaded with dye for a couple of hours before the experiment. As Julian mentioned some can act as a Ca buffer.

The answer to your question depends on exactly what you are trying to see.

We have experience  with single dye) , ratiometric dyes (INDO and FURA and also the transfection method based on a  calmodulin fusion protein with extended GFP of Tsien . We flew some experiments  in microgravity using the single wavelength  trying out a number of dyes.  With oregon green and  vitamin E using a flash LED light source we avoided a lot of  bleaching and could measure extended dynamics of calcium..

With FURA the method is supposed to be free of artefacts, but isnt. For One  you need expensive optics  for the 340nm wavalengh and the ligt sources (Xenon produce 50% of the light utput at 380nm, so you need a lot of calibration to try to get a quantitative result. IN addition  it ius usual to remove the background (slicing the  bittoim part of hte  dynamic range, and run a claibration for this. If a fast response time is needed to be measured then the wavlength changer needs to be fast too. To get around this we rebuilt an intense Xenon source (Helicopter landing light) and took off two light pipes each with a  different wavelength. It is also possible to take off 3 or more wavelengths from a single source and use fast shutters Drift in dye changes (dye consumption, bleaching  return into storage etc) can be measured at 360nm with FURA (stays the same ):  YOu can use manganese instead of calcium in the extra cellular medium to check if your calcium source is internal or external  ( Manganese interferes with FURA fluorescence  and also lead (which  potzentiates fluorescence as they come through the calcium channels. IN our experience gadolinium is very tricky, precipitating  at the drop of a hat.

We use thapsigarin also to check of origin of the calcium signal

Calcium channels (in our osteoblasts ) stop working at  19°C so our experiments had to be  at least 23°C to see any channel activity by this method.  Internal release was OK at 20°C using temperature is also a good control for distinguishing between inside/out origin if you do the work to establish the  temperature sensitivity or it is known for your cell type.

Adding hormones  to trigger  calcium is also very tricks as it doesnt mix immediately. A sophisticated method is needed to get the hormone onto the cells at the same time. Sometines you can see the diffusion spreading outwards in a cell culture.

Lastly FURA needs an expensive camera system (originally we used Intensified MCP cameras  and recently cameras with the sCMOS chips from Fairchild like Andor  or  Photonic Science. If you are using high res wide field  optics you can certainly use  a range of modern low light CCD and low light CMOS 2-4MP  chips which can be triggered and dyes like Oregon green which gives yo clear signals and  fast speeds having done a lot of background work using FURA

We built a ratioing  photometer for INDO to check our FURA results (frequency doubled

IR diode) worked well.

AM dyes certainly loaded at 37° in a serum free medium or if problems with cell attachment 1%.

We use thapsigarin also  to check of origin of the calcium signals.

So there are a wide range of dyes and methods including   a fusion protein (dual emission and ratio of the  GFP )

good luck

Thank you guys for the suggestions, pretty helpful !

Hey Martin, I agree with all what you said, but what did you guys eventually use for you ES cells ?

and do you think using FURA-2AM in plate readers would be sufficient to give a clear answer ?

Intracellular calcium accumulation level changes depending upon the source whether it is due to influx or eflux of calcium after your compound treatment. Therefore, it is advised to check calcium accumulation in presence of various extra-cellular and intra-cellular calcium chelators in case you are studying the kinetics. Ignore if it is an end point measure. Try to maintain the temperature of labeled cells at 37 degrees since stress also increases intracellular calcium thereby giving false results. Well to well variation due to time lag can also be a point of concern. I studied intracellular calcium increase after compound stimulation, through flow cytometry in suspension cells, but I guess its better to use plate reader in case of your cells. 

Good luck.

Dear Slim,

i've measured intracellular calcium using Oregon Greenn 488  BAPTA-1-AM, an intracellular calcium chelator. Cell were seeded  (15000cell/well) in 96 well plate (preferably black) overnight at 37°C and 5% CO2. Cells were loaded with Oregon Green 488 BAPTA-1-AM (from 2,5 to 5 uM, final concentration). Fluorescence intensity was monitored using a microplate fluorometer, set at appropriate excitation and emission wavelennght, at intervals that you prefer, until 5-6 hours at 37°C. Details you can read in paper

http://www.ncbi.nlm.nih.gov/pubmed/24440855.

Good luck

I agree with the answers submitted.

I add that I found enhancement in my calcium signalling measurement when I used fluo8, which has a number of improvement respect the other calcium indicator (one for all is less sensitive to temperature changes).

I now routinely use it, both on microplate reader and microscopy set up.

One thing to keep in mind is to get the appropriate controls (e.g. using the right molecules to load, or to deplete Ca++ from your cell system).

Good luck

http://aatbio.com/protocol/A3300d1.pdf

http://onlinelibrary.wiley.com/doi/10.1002/0471140856.tx0205s01/abstract

I agree with most of the comments, microscopy is the more accurate and best way to go.  You have to be careful with Fura2 because it requires a special UV objective, which doesn't come standard on most microscopes.  If you want to do ratiometric (which is more accurate and less dependent upon dye leakage), you need to make sure your objective passes the 340 wavelength light. 

I've used Fura2-AM in single cell microscopy and Fluo-4 in a plate reader. Personally, I prefer the single cell method. If you have a heterogenous population of cells, any changes in intracellular calcium may be lost with a plate reader, as you will be including cells that are not of interest in your analysis. Also, unless you have a plate reader with a pipetting robot, you will miss the first 20-30 seconds simply loading the plate and pressing go. So if you are looking for an acute response this may not be the method for you. Finally, using Fura2-AM it is possible to get an exact measurement of Ca2+ in the cell by calibration at the end of the experiment.

All this said, the plate reader comes into it's own when you want a high-throughput method, and the set-up for Fura2-AM is way more expensive, so it depends on what you need!

Dear Slim,

In my experience only ratiometric measurements allow Ca quantitation independent of loading, cell shape and bleaching. Also, unless you want to do rapid throughput, use a microscope. Individual cells tell you more than field average. I like Fura-2 AM (2-5 uM, 30-60 min at R or 37C) best, just be sure you get it from a reputable manufacturer (eg. Molecular Probes or Teflabs). Never re-use or refreeze. Order 50 ug microtubes and use one/experiment. The conditions for loading should be worked out for every cell type, only do not overload. You do not need a lot of signal with a good camera. Use good UV optics inverted scope with large aperture UV-transparent objective. My preferences- Nikon A40 or Zeiss A25 (a great objective but on the expensive side). You need a good camera (preferably intensified, my favorites either Photonics Science ISIS or Orca) and a good imaging software (Metafluor). A rapid wavelenght changer 340/380  - 2-3 pairs of frames/sec is usually as good as you might need, unless you require very rapid kinetics. To put your acquisitions in the right dynamic range, a set of neutral density filters (0.1, 0.2, 0.3, 0.5 and 1.0 OD) to dampen the exessive 380 signal and balance the weak 340. Use ionomycin (1uM) for max  and EGTA 5mM for min Ca signal required for ratios. And don't worry about absolute concentrations. Ratios will do, as all calculations are usualy rather incorrect (unless you put a lot of work into calibration). And last, but not least, keep EGTA frozen and thaw small aliquots. I never understood why, but fresh EGTA works and refrozen has problems (leaching metal ions from the plastic?).  Good luck and ask if you need more info.

The best way of measuring intracellular Ca2+ is using a flexstation. Load you cells with Fura-2. It is very easy and consistent. Check if there is one in your faculty.

I used to work with primary smooth muscle cells and loaded them with Fluo-4. If I loaded them at 37C I loaded the ER too, however, you may want to try and compare 37 vs RT to see if it's the case for your cells as well. You will have to optimize for your primary cells (including fluo-4 concentration), but here is what I used to do:

Cells were loaded for 40–50 min at room temperature in physiological solution containing 10 μM Fluo-4 AM (in DMSO) in the presence of 2.5% pluronic F-127 and subsequently washed for 30 min with physiological solution for deesterification. 

for intracellular calcium concentration measurements, i work à 37°c,  i used : specific buffer (134.8mMNacl, 4.7mM Kcl, 1.2mM K2HPO4, 1mM MgCl2, 1mM CaCl2, 10mM glucose, 10mM Hepes, pH7.4) be carefull with pH you must have the good pH at 37°C !

i used  : 1.5µM of FURA-2 AM and 0.006% pluronic acid and 30min of loading at 37°c

I would recommend you to use Fura-2 instead of Fluo-4  (according with the proposed protocols) in order to get a more quantitative analysis of calcium levels. Fluo-4 is more powerful if you are planing to do a dynamic study, with internal controls. The good point with Fura-2 is that you will calculate a ratio value between both measurements, so you will reduce the intra-sample variation.

Good luck!

The best temperature for loading with fura-2/AM or fluo-4/AM varies between cell types. However, it is usually safest and best to do it at room temperature, not at 37C. At 37C the dye is actively sequestered inside organelles within the cells (which gives artefacts in the signals measured), and it is also actively pumped out of the cells which reduces the equilibrium loading of the cytosol.

Whether fura-2 or fluo-4 is best depends on your exact experiments. Fura-2 has a higher affintiy for Ca2+ so it will saturate more easily than fluo-4. Hence if you anticipate large Ca2+ signals then fluo-4 may be best. If your Ca2+ signals are brief then fluo-4 is also best as it requires only a single wavelength measurement while fura-2 requires measurement at 360 and 380 nm, which slows down the rate at which measurements can be taken (eg. on a FlexStation this can be a few seconds between reads with fura-2, depending how many wells are being read). If fura-2 is used it is necessary to do a Mn2+ quench at the end of the experiment to establish the basal level of sequestered (unquenched) fluorescence, which can then be subtracted from all the measurements during the experiment. Both fura-2 and fluo-4 can be calibrated equally well: for Fluo-4 in FlexStation just use one well to add Triton+BAPTA to give an Fmin and one well to add Triton+high Ca2+ to give an Fmax - together with the published Kd for fluo-4 this can then be converted to [Ca2+]. For fura-2, just use a prior ratio calibration along with the final Mn2+ quench correction to convert to [Ca2+]. For a protocol on FlexStation measurements of Ca2+ see: Tovey et al., 2006 (Nat Protoc. 1(1):259-63). The intensities of 360/380 and 488 nm illumination along with sensitivies of cameras/photodiodes on your systems may also dicate whether fura-2 or fluo-4 is best.

Whether to use plate reader or microscope also depends on exact experiments: if you are measuring Ca2+ oscillations or responses to single reagents then maybe go for microscopy; if you are comparing a number of treatments or making dose-response curves then the plate reader will save you a lot of time. Not all cell types give sufficient signals in plate readers anyway, so you may have to go to microscopy. Whether your feeder cells will interfere with the measurements from the ES cells will also dictate whether you need to use microscopy?

 David L Prole, I have question  to you;

Both fura-2 and fluo-4 can be calibrated equally well: for Fluo-4 in FlexStation just use one well to add Triton+BAPTA to give an Fmin and one well to add Triton+high Ca2+ to give an Fmax - together with the published Kd for fluo-4 this can then be converted to [Ca2+]. For fura-2, just use a prior ratio calibration along with the final Mn2+ quench correction to convert to [Ca2+].

 -Can I use for Fura-2 single cells measurement 'add Triton+BAPTA to give an Fmin and one well to add Triton+high Ca2+ to give an Fmax -'? if not why?

and why for  fura-2, "just use a prior ratio calibration along with the final Mn2+ quench correction to convert to [Ca2+]."? which concentration of Mn2+ do you use? I am having problems to get my Rmin.. how do you get Rmax with single cell measurement?

Thanks!

Hi Meerim,

There is no need to obtain an Rmin and Rmax for every experiment when using fura-2/AM. Just calibrate your system using drops of fura-2 (pentapotassium salt, not AM ester) in buffers of defined [Ca2+] (eg. Invitrogen calibration kits) to give 340/380 values at each [Ca2+], then use the published Kd for fura-2 to calculate the [Ca2+] in cells for any measured 340/380 ratio (see calibration kits manuals for calculation method). Remember to keep illumination settings the same for calibration and experiments. Mn2+ quench required at end of each experiment (1 microM ionomycin + 10 mM MnCl2) to enable subtraction of signal from sequestered dye (which can be large). See following for method (although generally 20C loading is best):

http://jcb.rupress.org/content/202/4/699.long

fura2 can not be loaded into fetal sheep cardiomyocytes, any comments?

This should be determined experimentally for each cell line and each tracer. I always preferred to load at RT with higher concentrations of Fura rather than lower concentrations at 37. At 37 it is relatively common to overshoot and get some of the dye sequestered in organelles. A simple evaluation is to look for punctate fluorescence, which implies that you overshot the loading. Even if not overtly overloaded, it is always better to work with minimal tracer load, as every tracer is essentially a calcium buffer and will greatly change the shape and kinetics of the response.

Can we measure calcium in aorta? just on confocal microscope?