Tag Archives: spectrum

An excelent new tool for comparing fluorescent protein properties

Screenshot of the new tool

Screenshot of the new tool

Two very useful tools for visualizations of many fluorescent and photoswitchable proteins have been developed by Talley Lambert and Kurt Thorn (UCSF). Continue reading

Red alert: the three color dilemma continues

So, I sat with the biophysicist who built the microscope that I intend to use. We discussed my proposed experiment and the different capabilities of that particular microscope.

Immediately we discovered that I will not be able to use LSSmKat1. If you recall, the emission peak of LSSmKate1 is at 624nm. However, the dichromatic mirrors set in that microscope for the red channels are from ~500 to 620 and from 650 to 750. So the 624 falls just in the waveband that I cannot detect.

So I remain with LSSmKate2, with em. Peak at 605.

Ok, now, the available lasers in this microscope are 407, 436, 488, 561 and 640nm.

GFP is excited at 488. That the peak and that’s good. We will use the 525/30 filter (i.e. from 510 to 540).

mCherry is excited at 587, so using the 561 laser we can get a fair signal (~60%). We will use the 593/40 filter (i.e. from 573 to 613).

This means that in both cases I collect only part of the emitted photons (the ones within the filter range, see fig 1).

spectra of GFP & mCherry, with filters. (image: BD spectra viewer)

LSSmKate2 can be excited with the 436 or 488 laser.  The 488 will give a ~8% signal for mCherry.  But the 436 will not. Although LSSmKate2 is 3-fold less bright than mCherry, and is less photostable, to object using this marker is only for time zero, when the GFP and mCherry signal should be low. However, this needs to be experimentally tested. It could be that the mCherry signal could be mathematically subtracted later.

The problem is that I would not be able to excite simultaneously at 488 and 561 to get simultaneous data of GFP and mCherry. This is because I would also have LSSmKate2 excited. However, for my purposes, I do not have to take the images simultaneously. The microscope can take images with the different lasers at 50-100ms time intervals. Since I expect to detect changes in the seconds to minutes range, a 0.1sec difference between the green and red channel is acceptable.

There is another option and that is to use a far-red protein, such as TagRFP657.

FP Max ex. Max em. QY EC(M-1 cm-1) Brightness Relative brightness Photostability
TagRFP657 611 657 0.10 34,000 3400 0.1 55 sec
LSSmOrange 437 572 0.45 52,000 21400 0.82 10 sec

The advantage is that we can use the 650-750 dichromatic, and use excitation at 640, which still gives a fair excitation, and it will not excite mCherry.  The down side is that the 561 laser will excite TagRFP657 too. However, the TagRFP657 emission at the 593/40 filter is fairly low and with a less than maximum excitation to begin with, and a lower brightness and photostability (compared to mCherry), this should not be a big problem (see figure 2) . But again – should be tested experimentally.

figure 2: TagRFP657 spectra (From: Morozova et al. Biophys J. 2010 99(2):L13-5.)

So here is another lesson: when designing a multi-color experiment, you need to take into account both the FP properties, and the capabilities of your microscope (lasers, filters, dichromatics, speed etc…).

Update 22.6.12:

After further discussion, another LSS FP was suggested, LSSmOrange. I added its characteristics to the table above.  It is brighter than TagRFP657, and the Stokes shift of LSSmOrange is better than LSSmKate2 in relation to mCherry (bluer excitation, which means NO excitation of mCherry.) So I will test all three, and hopefully have an answer which is better within 2-3 weeks.

The red color dilemma: how to choose the right fluorescent protein

This post continues the previous post.

I encountered a serious dilemma in choosing the right protein. This is also a great opportunity to learn about the many properties of fluorescent proteins.

Let us start with the obvious: excitation and emission maxima and spectra.

We all know that EGFP has an excitation maximum at 488nm. That is, EGFP protein that is excited with photons at 488nm will give its maximum emission intensity at its emission maximum, 509nm.

However, we must remember that this is the maximum emission. The emission spectra is much wider, and for GFP it goes from ~470nm up to ~630nm. Figure 1 shows the EGFP excitation (dashed line) and emission (full green) spectra.

Fig. 1: GFP specrum with 488 excitation

And now, what if we want to look simultaneously at two colors, EGFP and dTomato?

Figure 2 (upper panel) shows you that excitation at 488nm (the max ex. of EGFP) also excite dTomato, leading to almost 30% emission at its peak of 581nm. Therefore, if we would collect all the emitted photons, we would detect the combined excitation of EGFP and dTomato.

Fig. 2: GFP dTomato spectra with 488 excitation. Lower panel: with filters.

We therefore use filters with a narrow band of wavelength (fig. 2, bottom panel). Thus, if we use the 510/20 filter, we would detect only photons emitted at the band, in this case only from EGFP. If we use the 580/30 filter, we would detect photons coming from dTomato and EGFP.

Now what happens if we add the long-stokes shift protein LSSmKate1 to the system? Figure  3 shows LSSmKate1 is maximally excited at 463nm (red dashed), and has max. emission at 624nm (full red) [note- I drew the LSSmKate spectra, based on prior publications]. However, 460nm also excites EGFP and dTomato. The EGFP signal, at 624nm is negligible. However, dTomato gives emission at 8-9% of its maximal emission. Is this negligible? We will soon learn.

Fig. 3: GFP dTomato LSSmkate1 spetra with 460 excitation

Now let’s look at mCherry. With excitation laser at 460, we have virtually no excitation of mCherry. We would therefore prefer to use mCherry in our 3-color system, instead of dTomato.

Fig. 4: GFP mCherry LSSmkate1 spectra with 460 excitation

This table summarizes the excitation & emission maxima:

FP Max ex. Max em.
EGFP 488 509
dTomato 554 581
LSSmKate1 463 624
LSSmKate2 460 605
mCherry 587 610

This table shows the relative emission of the different FPs with different excitations (the LSS data is estimated based on publications, since the BD

spectrum viewer does not include LSSmKates in its database):

  % from max emission % from max emission

Em   (nm):

509 580 605 610 624 509 580 605 624
EGFP Ex.488 100% 8% <5% Ex.460 65% 5%
dTomato 27% 20% 19% 15% 15% 11% 9%
LSSmKate1 50%? 100
LSSmKate2 50%? 100
mCherry 8%

However, excitation & emission spectra are not the only considerations. There are other parameters to consider.

Brightness, QY and EC

Different fluorescent proteins have different brightness: some are very bright, some are dim. As a matter of history, we usually consider the relative brightness, compared to EGFP (which is set to 1.00). You can see in the table below the relative brightness of the proteins discussed above.

The brightness is determined by two parameters: quantum yield (QY) and Extinction coefficient (EC).

QY is simply the ration between the number of photons absorbed to the number of photons emitted. If QY=1, then for each absorbed photon you get one emitted photon.  However, we never get 100% efficiency. The protein with the highest QY that I encountered is called ZsGreen with QY=0.91.

EC (or ε) relates to the formula A = εcl in which A is the absorbance, l is the path length (in cm, usually) and c the concentration in Molar units. ε, in M-1cm-1 units, is a measurement of the capability of a certain fluorescent protein to absorb light at a certain wavelength.

The formula ε*QY gives a measure of the brightness, so for EGFP, the brightness is 55,000*0.60=33,000

For ZsGreen, the EC is only 43,000 so the relative brightness is only 1.18. This example shows that a high QY does not necessarily mean a very high brightness.

This table summarizes al the brightness data:

FP QY EC(M-1 cm-1) Brightness Relative brightness
EGFP 0.60 55,000 33,000 1.00
dTomato 0.69 69,000 47,610 1.44
LSSmKate1 0.08 26,000 2080 0.06
LSSmKate2 0.17 31,200 5304 0.16
mCherry 0.22 72,000 15,840 0.48

Let’s go back to our dilemma: dTomato or mCherry?

We now know that with a laser excitation of 460nm, we get 100% emission from LSSmKat1 at 624nm, and only 9% emission from dTomato at 624nm.

However, dTomato is 24-fold brighter than LSSmKate1.  If we look only on the QY data, at 8% emission, dTomato will produce ~6 photons for each 100 photons (100*0.69*9%), whereas LSSmKat1 will produce 8 photons (100*0.08*100%). In other words, although we get very low emission of dTomato, relative to its maximum, it is almost as high as the LSSmKate1 emission in that wavelength. Since eventually, we just “see” the photons with no knowledge of their source, the signal that we will get will be ~40% from dTomato. This will create a problem if we wish to detect changes in fluorescent intensity over time due to changes in protein localization (or other changes) because we will not be able to know if the change is due to dTomato or LssmKate1.

That is why mCherry is a better choice than dTomato for this experiment.


Another parameter to take into consideration is the protein’s photostability.

Photostability is a measure (in seconds) of how long it takes for half of the number of proteins to bleach at the maximum excitation.  This matter is important for any experiment involving fluorescent proteins (or dyes) but is crucial when doing live imaging, particularly for time-lapse experiments.

LSSmKate1 is more stable than LSSmKate2 (see table below), and therefore, it might be a better choice than LSSmKate2 for my time-lapse experiment.

FP photostability
EGFP 174 sec
dTomato 98 sec
LSSmKate1 60 sec
LSSmKate2 44 sec
mCherry 96 sec

There are other parameters to take into account, some of them I discussed in earlier posts.

These include:

pH stability (and the effect of pH on the absorption and emission spectra).

Maturation rate – how long does it take your fluorescent protein to fold correctly and create the chromophore? If you are using your protein to measure expression rate, and the maturation time is longer than the time frame of your experiment, then you will not get the information you are looking for. Maturation rate  depends on protein characteristics, as well as oxygen levels and temperature. Several fast-folding FPs were developed in recent years.

Oligomerization state – many FPs are naturally monomeric. However some FPs particularly in the orange & red range) are dimeric or tetrameric.  Since in many cases the FP is genetically encoded as a fusion with a protein of interest, fusing a dimeric FP may cause your protein of interest to dimerize, through the FP, thus altering its biology. If your protein is naturally oligomeric, fusing it to a dimeric or tetrameric FP may create large protein aggregates. One solution for dimeric FPs is to create a tandem fusion proteins (i.e. two FPs are fused one to the other and to the your protein).

Generally, the letter m at the beginning of the FP name indicates that it is monomeric (e.g. mCherry, mKate), d indicates it is dimeric (dTomato) and td indicates tandem dimer (tdTomato). However, there are many FPs with no indication of their oligomeric state in their name (e.g. TurboRFP is dimeric). EGFP and its many derivatives are monomeric (except at high concentrations when they form weak dimers).

Phototoxicity  – although the FPs themselves do not emit harmful radiation, the excitation light can be harmful to the cell. For instance, UV light, that is used to excite BFPs, as well as some LSS FPs and photoactivatable FPs, can cause direct DNA damage, create reactive oxygen species that will cause an oxidative stress etc…  This is one reason why BFPs are the least popular FPs, particularly for live imaging.

Three-color problem

The most common application of fluorescent proteins in biological research is to visualize the localization of the FP, fused to the protein of interest (thus learning about the localization of your protein). Though looking at the localization of one protein can be informative, it is even more informative if one can detect co-localization with another protein or molecular marker (e,g, a dye that stains a specific organelle).

However, in order to detect co-localization it is required that both FPs will have different emission spectra (e.g. GFP & RFP). The problem of finding non-overlapping colors becomes more difficult when we want to use three colors (or more).

Using the spectrum viewer (see previous post), you can see that in some cases the excitation and/or the emission spectra overlap. Such is the case with CFP and GFP, for instance. Although the excitation maximum for GFP is 488nm and for CFP is 436nm, using a 436 laser will excite some GFP (~35% at peak emission). More importantly, though CFP emission peak is 477nm, the spectra of CFP  goes all the way to 600nm (~5%), and definitely overlaps that of GFP. Therefore, the signal (photons) that we detect at the 509nm (GFP peak) can come from CFP emission.

In order to reduce the overlap excitation or emission, we use special filters and dichromatic beamsplitters (“mirrors”) that allow passage of light at specific band of wavelengths.  These filters are housed in a filter cube. There are different kinds of filters with different ranges of bandpasses for each color.

However, even with filters, overlap can occur, particularly if the filters in your microscope are wide-range. A possible solution to a three color problem (assuming you expect all three to co-localize) is to use a large Stokes shift (LSS) protein. Stokes shift (as mentioned in an earlier post) is the shift in wavelength from excitation to emission. For most FPs, Stokes shift is less than 50nm (usually much less). For LSS proteins, the shift is over 100nm. For example, the fluorescent protein mKeima has an excitation maximum of 440nm (blue), and emission maximum at 620nm (red). Thus, using the correct emission filter for> 600nm, excitation at 440nm will result in emission of CFP, GFP and mKeima, but only mKeima emission will be detected.  However mKeima is not a good choice if you have RFP in your system as well, since mKeima also has a small excitation peak at 590nm that give a similar emission peak at 620nm. This is true for acidic pH, and therefore crucial for the experiment I design in yeast (yeast cytosol is mildly acidic, pH between 5.5 to 7.5). Furthermore, mKeima has low photostability and low brightness.

Therefore, I am going to use a protein called LSSmKate1, with ex/em peaks at 463/624nm or LSSmKate2 (460/605) with no other excitation peak at longer wavelength (except at very basic pH=11).

Two papers on LSSmKates:

Monomeric red fluorescent proteins with a large Stokes shift (Kiryl D. Piatkevich, James Hulit, Oksana M. Subach,Bin Wu, Arian Abdulla, Jeffrey E. Segall, and Vladislav V. Verkhusha. PNAS (2010) 107(12):5369)

Engineering ESPT Pathways Based on Structural Analysis of LSSmKate Red Fluorescent Proteins with Large Stokes Shift (Kiryl D. Piatkevich, Vladimir N. Malashkevich, Steven C. Almo and Vladislav V. Verkhusha. J. Am. Chem. Soc., (2010), 132 (31):10762)

Fluorescence Spectrum Viewer

During training on the FACS machine in our faculty facility, I encountered the Fluorescence Spectrum Viewer  from BD bioscience.

It is a JAVA interactive site that allows to to view excitation/emission spectra of up to 8 differnt fluorophores simultaneousely.

It lets the user to choose the desired excitation laser, and shows the excitation & emission spectra *to that laser*. moving the cursor over the spectra gives you actual numbers of % excitation & % emission efficiency.

You can use filters to test which filter is most suitable for your fluorophores.

This site is intended for FACS users, so you can choose from any one of a number of cytometers from BD. However, I think this site can be usefull for microscopists too.


EDIT: I found more sites like that, each with its own set of fluorephores –

Fluorescence SpectraViewer at Invitrogen (Life technologies) site.

Fluorescent Dye Spectra at U Arizona (only chemical fluorophores)

Evrogen Spectra Viewer at Evrogen (only a very limited list of fluorescent proteins)


There is also this Cell staining tool from invitrogen. The concept is cool, but I think it is a very limited tool and I don’t think it is very helpful.