Why Cell Cycle Analysis Details Are Critical In Flow Cytometry
The lifecycle of a cell can be described in stages. In diploid cells, much of the time they exist in a resting state, where a cell does what a cell does — such as, undergo differentiation. In some cases, the cells go into a quiescent state, where the level of RNA is reduced. When the appropriate signals are received, cells begin to bulk up and start to replicate the DNA in preparation for division into 2 daughter cells. After the synthesis phase, the cells enter a second period of rest, where everything is checked before the cells undergo mitosis and produce 2 daughter cells. The cycle repeats itself until the cells die. The cell cycle is usually depicted as shown in Figure 1.
Figure 1: The Cell Cycle. Image from Wikipedia.
While there are many differences in cells at each stage of the cell cycle, one of the most obvious is the amount of DNA that the cell contains. At the G0 and G1 phase, the cells have a normal amount of DNA (2N for a diploid cell). Upon entering the S phase, the DNA concentration begins to increase until it doubles (4N) and the cells reach the second gap (G2) phage. The cells eventually undergo mitosis (M), producing 2 daughter cells with 2N DNA content.
Diseases including cancer, Alzheimer’s, Parkinson’s, and more, are all caused at some level by cell cycle dysregulation. Cells, such as the Megakaryocyte, undergo endoreduplication as part of the normal development. In plants, polyploidy is common. The durum wheat used to make pasta is a tetraploid wheat, while a hexaploid wheat makes your bread lighter. Thus, cell cycle analysis remains an important tool in the researcher’s toolbox.
Cell cycle analysis was one of the first clinically robust flow cytometry assays, where it was used to examine the DNA content of tumors to gauge the aggressiveness of the cancer. In fact, Shankey and colleagues published guidelines on how to implement DNA analysis in the clinic.
Cell cycle analysis appears to be a deceptively simple assay, as the base assay only requires 1 fluorochrome. However, there are many steps necessary to optimize to get high-quality cell cycle histograms.
Basic cell cycle protocol
Below are 2 basic protocols that use Ethanol for fixation. One uses PI, and the other, DAPI. The choice of dye will be discussed below.
Ethanol fixation is preferred for cell cycle analysis, and while aldehydes can be used, the crosslinking nature of these chemicals can impair the DNA staining when using intercalating dyes, resulting in a less accurate measurement (Darzynkiewicz, et al. 2017).
Shown here, is a basic protocol for staining for cell cycle only, with the most commonly used DNA dyes, propidium iodide, and DAPI.
The main difference between these 2 dyes is that when using propidium iodide, RNase needs to be added as well.
No matter which dye you are using, take about one million cells and fix them with ice-cold 70% ethanol. It is critically important to add the cells to the ethanol in a dropwise fashion. Have the tube on a vortex, moving at a reasonable speed — not slow, but not resuspend-my-DNA fast. Drop the cells into the center of the vortex and wait until the cells are fully mixed with the ethanol before adding the next drop. It takes practice, but if the cells go into the ethanol too fast, you will end up with goop.
Once you have fixed the cells, they can be stored in the fridge for a few weeks. There may be some signal degradation after a week or two, but it’s very much cell type and cell line dependent. So, when planning to store cells, make sure you do a test first.
After the cells are fixed, they can be stained and analyzed.
First, centrifuge the fixed cells — 10 minutes at 300 g is a good place to start. Ethanol-fixed cells don’t pellet like normal cells — the ring may be more diffuse than what is seen with non-fixed cells.
Next, resuspend the cells in PBS and allow the cells to rehydrate for between 30 seconds to 2 minutes. Spin the cells down again and resuspend in a staining buffer.
When making the staining buffer, for propidium iodide, use approximately 50 u/mL propidium iodide and 100 ug/mL RNase A. Make sure the RNase A is DNase-free. If you cannot find DNase-free RNase A, heat up the RNase A to deactivate the DNase. The RNase A will survive the heat — it’s one of the most stable proteins on the face of the Earth.
Wait at least 30 minutes at room temperature for the RNase A to work, and the propidium iodide to bind.
If you are using DAPI instead of PI, after you rehydrate the cells in PBS and spin them down a second time, you can resuspend the cells in a flow buffer containing 1 mg/mL DAPI.
Wait at least 30 minutes before analyzing on the cytometer to give the DAPI time to bind the DNA. 30 minutes is just a starting point, and depending on your cell line, you might need to change this time to get optimal staining.
Choosing a DNA binding reagent.
There are many reagents that will bind DNA, and they tend to bind in a couple of different ways. The dye can bind to the major or minor groove, may be AT or GC preferential, or may intercalate into the DNA strand itself. This is summarized in Figure 2.
Figure 2: Different binding characteristics of DNA binding dyes (From Thermofisher).
With intercalators, the dye actually wedges in between the base pairs in the DNA helix. Propidium iodide, 7-AAD, and DRAQ5 are 3 common intercalators.
Some dyes are bis-intercalators. Most of the bis-intercalators have a doubled-up name. There’s TOTO and YOYO and POPO.
Methyl green is an example of a major groove binder, but this isn’t commonly used in flow cytometry.
All of these dyes bind stoichiometrically, which means that they will bind at specific ratios to the DNA.
These dyes also come in all regions of the spectrum. So, depending on the instrument configuration and the needs for downstream assays or multiplexing, there are dyes that will bind to DNA and might give an acceptable cell cycle analysis pretty much anywhere in the spectrum.
Shown here are some of the most common dyes used for cell cycle analysis.
DAPI and the Hoechst dyes prefer a UV excitation source, but they can sometimes run off of a violet laser. So, if you don’t have a UV laser, do a little test run and see if there is good enough resolution using the violet laser for them.
DAPI is only slightly cell permeable, so the cells need to be fixed when using DAPI so it can access the DNA.
There are two Hoechst dyes, there’s 33342 and 33258. Hoechst 33342 is cell permeable, and so is a great dye to choose for supravital cell cycle. Meanwhile, Hoechst 33258 is only a little bit cell permeable and would be used on fixed cells.
Propidium iodide is not cell permeable, and requires cell fixation. Additionally, 7-AAD is not cell permeable either.
DRAQ5 is cell permeable but, word of warning, it is known to be cytotoxic. So, using DRAQ5 on live cells is not recommended if those cells need to live afterwards. DRAQ7 is an interesting DNA dye because it has an extremely far red emission peak, so it is an option to get a cell cycle analysis from way out in the panel.
Variables to consider when optimizing your protocol.
DNA staining protocols will need to be optimized based on the cell type, dye, or other situational factors, to ensure the best results and there are a few key variables to consider when optimizing.
First, consider time. This is especially critical when staining viable cells.
Second, consider the concentration of dye. The dye-to-cell ratio is very important when doing cell cycle by flow cytometry. If there is not enough dye, the DNA will not be saturated and the peak CVs become large. The ratio between the G1 and G2 peaks may not be quite what it should be.
So, count the cells you are using and make sure that you’re using the appropriate amount of dye for that number of cells. .
Third, consider the temperature. For live cells, the temperature they are stained at also makes a difference. For example, in the following figure, the difference between staining with Hoechst 33342 at room temperature versus 37℃ is shown. When the cells were stained at 37℃ the peaks were much better.
One additional thing to know about Hoechst 33342 is that cells that have drug transporters will be able to kick the dye back out. So, make sure there is enough dye around to keep the DNA saturated, even while the cells are pumping some of it back out.
Fourth, consider how you will fix your cells. The best cell cycles are usually done with ethanol or a similar precipitating fixative. Formaldehyde is really good for fixing proteins and keeping things around, but it can cross-link the DNA and the chromatin, and it restricts access for DNA dyes. Also, consider whether you will use detergent as a part of your fixing protocol.
Consequences of using fixatives or detergents.
In the example shown above, one well of cells is treated exactly the same, and then split into 2 groups and fixed 2 different ways (with ethanol or formaldehyde) to demonstrate the impact of fixing on outcome.
When these assays were put into ModFit to model them, the CVs are much lower on the ethanol-fixed cells than they are in the formaldehyde-fixed cells. 6% isn’t awful, but it is definitely not as good as just under 5%. And, you can see differences in the peak shoots.
If ethanol fixation is compatible with your assay, this is where you should start.
Next, to determine if you want to use detergent or not, consider the type of analysis you want to do. Do you want to analyze isolated nuclei, or do you want to keep the whole cell around?
If you do an ethanol fix with no detergent, the outer membrane is going be a little bit permeabilized. It will be permeable enough to let dyes in, but it’s not going be enough to let RNAs out. So, with ethanol fixation, most of the cytoplasmic contents will remain.
However, if you fix with detergent, it’s going to dissolve the cells’ outer membrane. The RNAs and other cytoplasmic confounders are going to dissociate, and you will be left with a more or less bare nucleus. Fixing with detergent does, however, get rid of the need for RNase.
But, the biggest problem with using detergent is that during mitosis, there’s the state when the nuclear envelope is actually broken down so that the sister chromatids can get pulled apart, and they go into their new cells. If you expose these cells to detergent and they are in mitosis, the chromosomes and the DNA leak out. So, when using detergent, it’s possible to underestimate the number of cells in mitosis.
Importance of titrating your dyes.
There is a lot of talk about the importance of titration with antibodies. But, it’s not talked about as much concerning DNA dyes. But, titrating your DNA dyes is important.
When doing cell cycle with propidium iodide, it might seem okay to think, “It looks pink, it’s probably fine.” But, this does not allow you to tell if the dye is actually saturated or not.
This becomes more important when working with DAPI, because DAPI is not very water soluble. If you are using too much or too little dye, your peaks will not look as good as they could.
In the image above, you can see how much nicer the correct concentration of DAPI looks. It’s a much tighter peak and the background is a lot lower. When there is a lot of DAPI, there is a lot of junk hanging around. But, if there is not enough DAPI, like down here at 0.1 mg/mL, there is poor separation.
Again, the amount of dye that you need to use is probably going to be cell type dependent. So, do a quick titration. It doesn’t take too long, and it will save you time and frustration in the long-run.
These are the basics of cell cycle analysis. In upcoming blog posts, we will discuss more advanced techniques and data analysis to ensure that your cell cycle experiments are consistent, reproducible, and informative.
Cell cycle analysis is deceptively easy in concept, but details are absolutely critical. It is not possible to hide the data if there is poor sample preparation, incorrect dye ratios, too much (or too little) staining time, etc. Forgetting RNAse when using PI will doom your data to failure. Take these basics into account as you move into performing this simple, yet complex assay.
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