Which Fluorophores To Use For Your Microscopy Experiment

Fluorophore selection is important. I have often been asked by my facility users which fluorophore is best suited for their experiments. The answer to this is mostly dependent on whether they are using a widefield microscope with set excitation/emission cubes or a laser based system that lets you select the laser and the emission window. Once you have narrowed down which fluorophores you can excite and collect the correct emission, you can further refine the specific fluorophore that is best for your experiment. 

In this blog  we will discuss how to determine what can work with your microscope, and how to refine those options to achieve what’s “best” for your microscope and your experiment.

Matching Fluorophores to Filters

It is imperative to understand that the excitation and emission for a fluorophore is not a single wavelength. For example, Alexa 488 is listed with an excitation max of 490 nm, and an emission max 526 nm. The reality is that there is greater than 10% efficiency to excite GFP from about 450 nm- 525 nm. This means that if you have high light levels, and sufficient fluorophore present, the excitation in that range will result in some amount of emission. This is why you can observe GFP signals with cubes designed for GFP or YFP (Figure 1). However, you will have a brighter signal with less illumination if the cube is matched well with your fluorophore. 

Figure 1. eGFP (A) vs YFP (B)  filters with AlexaFluor 488 dye. Excitation (left curve) and emission right curve) of AlexaFluor 488 is displayed in green. The blue line indicates the wavelengths that are transmitted during excitation, while the red line indicates the wavelengths transmitted for emission. The black line is a dicrotic mirror to prevent transmission back through the incorrect filter. Both filter sets cover some of the excitation/emission curves of AlexaFluor 488, so you would be able to observe the fluorescence with either filter. Curves created with Chroma Spectra Viewer

When I arrived at my post doc laboratory, there was a widefield microscope with a “red filter” and in the freezer there were secondary antibodies conjugated to Alexa 546, 555, 568, and 594. Surprisingly, no one knew which one was best for the microscope since they all “worked.”  But I was working with valuable patient samples, so I didn’t want mediocre data from my experiments. Therefore, I took out the cube and located the technical specifications of the filters (Figure 2). Hopefully, your lab will have those details documented, but it is important enough to take out a cube to find the answer. The emission profiles of these dyes can be found in Figure 3.

Figure 2.  How to Find the Technical Specifications of a Filter Cube. When you remove the filter from the microscope, most manufacturers will have labels with the excitation filter, dichroic mirror, and the emission filter clearly listed.  In these filters from Olympus Microscopy, the labels are the yellow rectangles on the side of the filter

Prevent Bleedthrough

Bleedthrough or “cross talk” occurs when more than one fluorophore becomes excited and emits in the same channel. This can affect your quantification measurements as well as your image quality. Therefore, once you have set up your experiment, it is very important to run controls such as “fluorescent minus one” (FMO). You might not be gating, but the same concepts as used in flow cytometry still apply.

Now, before you get to your wet lab work, there are a few best practices to mitigate future problems. The first and easiest fix is to pick fluorophores that are as far away from each other on the light spectrum as possible. For example, you might have a virus protein tagged with the fluorescent protein mCherry (excitation max: 587nm) and you would like to stain nucleic acids. Something like STYOX Orange (excitation max: 547nm) will cause bleed through, but you are much less likely to have bleedthrough with DAPI (excitation max: 350nm). While performing a two color experiment if you have the option, then pair UV (blue) with red or green with far-red.

Figure 3. Emission spectra of closely related Alexa Fluor dyes. In this example we compared four “red” dyes number 8-11 on this chart. Notice how there is a lot of overlap, but each curve is shifted slightly. Image from Thermo Fisher

Photostability

Photostability is the rate at which a fluorophore will bleach when exposed to light. This trait is treated the same as radioactive decay and is represented as t1/2 (s) for fluorescent proteins and pb (probability of bleaching) for fluorescent dyes. These numbers can be a general guide for which fluorophores are particularly stable, but these rates can vary due to light source, intensity of light source, wavelength, rate of exposure, etc. 

So when should you take photostability into account? One example is experiments involving single molecule detection (SMD) such as fluorescence correlation spectroscopy (FCS). FCS monitors the diffusion of a single molecule within a defined space. Other single molecule techniques such as STORM/PALM depend upon photostable molecules to be able to reconstruct the blinking of the fluorophores over time into a usable image. 

Fluorescence Recovery After Photobleaching (FRAP) is another instance where knowing the photostability of a fluorescent protein is important. The ideal protein for this is something that will bleach easily at high laser power, but remains relatively stable during low light imaging. Otherwise, your intensity measurements will be faulty if you are bleaching the fluorophores that have newly arrived in the previously bleached region.

A collaborator that I’m working with is performing live cell imaging of actin fibers at 10-100 frames per second (fps). The laser needs to be used at a relatively high intensity to acquire an usable image at that rate, but photobleaching causes the experiment to only last  for a few seconds. Therefore, we started to look at new fluorophores that had better photostability. We found that SiR-actin had the stability we needed and were therefore only limited by phototoxicity. 

Maturation Time

Maturation time is a characteristic of fluorescent proteins, which is the time after translation that is needed for a protein to become fluorescent. For example, eGFP requires 25 minutes to mature while mNeonGreen requires 10 minutes. The time to maturation can be over two hours (mRuby3)! 

Why does this matter?

Any experiment that require highly dynamic expression for the measurement needs to have a rapidly maturing fluorescent protein. Balleza et al. demonstrated that a mutation in eGFP that allowed maturation to occur within 5 minutes, was able to pick up specific translational bursts in yeast that were not possible with just eGFP. 

If the time of translation is not important for your experiment— for instance like labeling histone proteins to visualize the nucleus— then this factor can be a lower priority while choosing your fluorophore.

Brightness

Brightness is mathematically defined as:

Brightness = Extinction Coefficient (ε) x Fluorescence Quantum Yield (Φ)

Or more simply put, brightness is a function of how well a molecule absorbs light and the likelihood of that molecule emitting a photon. Having a bright fluorophore means that you can use less excitation power and/or shorter exposures to collect the needed photons for your microscopy image. Less light exposure will also translate to less photobleaching or phototoxicity! 

Let’s take a look at eGFP vs mNeonGreen again for their brightness comparison. Perfect quantum yield would be “1” because if a single photon excites the fluorophore, then you will have an output of one photon for emission. 

We don’t live in a perfect physics world, though.

For our example eGFP has a quantum yield of 0.6 (or 60%) vs mNeonGreen has a quantum yield of 0.8 (80%) meaning that mNeonGreen is significantly more likely to emit a photon upon excitation than eGFP. When combined with the extinction coefficient, we get a brightness of 33.5 for eGFP vs 92.8 for mNeonGreen. Under ideal conditions, mNeonGreen is 3x brighter than eGFP. As you can see, brightness can have a big effect on how you set up your experiments.

Conclusion

There is no universally “right” fluorophore to use for experiments. However, researching some parameters before starting can save you both time and money. No one wants to waste time on an experiment that will not work due to the characteristics of their fluorophore. Matching the fluorophore to your system, preventing bleedthrough, knowing the photostability, understanding the maturation rate, and being aware of the brightness of a fluorophore will put you on the right path. Make sure to check out experiment specific proteins such as the pKa, properties under specific temperatures or different organisms, and if the fluorophore will cluster before you start. Using this knowledge will set you up for success. 

To learn more about important techniques for your flow microscopy lab, and to get access to all of our advanced materials including 20 training videos, presentations, workbooks, and private group membership, get on the Expert Microscopy wait list.

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Heather Brown-Harding
Heather Brown-Harding

Heather Brown-Harding, PhD, is currently the assistant director of Wake Forest Microscopy and graduate teaching faculty.She also maintains a small research group that works on imaging of host-pathogen interactions. Heather is passionate about making science accessible to everyone.High-quality research shouldn’t be exclusive to elite institutions or made incomprehensible by unnecessary jargon. She created the modules for Excite Microscopy with this mission.

In her free time, she enjoys playing with her cat & dog, trying out new craft ciders and painting.You can find her on twitter (@microscopyEd) a few times of day discussing new imaging techniques with peers.

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