An introduction to flow cytometry with troubleshooting tips webinar

Review how flow cytometry can be used in a wide range of applications such as cellular phenotyping, apoptosis, or cell cycle analysis. 

Flow cytometry is a powerful technique that allows rapid and quantitative measurements of multiple parameter on individual cells simultaneously. Sheath fluid is used to transport these cells in single file past optical and electronic sensors to detect physical and biological characteristics.

This webinar will review how multiparameter flow cytometry permits precise characterization of multiple cell subpopulations in complex mixtures.

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Webinar Topics:

  • Sample processing
  • Antibody/fluorochrome selection
  • Controls

About the Presenter:

Dr. Ina Schulte is currently a member of Abcam's Scientific Support team. She has a MS in Biochemistry and Molecular Biology from the Friedrich-Schiller University in Jena, Germany, and a PhD from the University of Essen-Duisburg in Germany.

Before joining Abcam, Ina worked as a postdoc at Cambridge University. Her flow cytometry experience includes surface staining, as well as intracellular cytokine staining, and troubleshooting for both techniques. ​

"Our lab enjoyed listening to Ina's talk especially regarding the troubleshooting staining techniques."

- Attendee

Webinar Transcript:

Hello. Welcome to Abcam's webinar on an introduction to flow cytometry. The principal presenter will be Dr. Ina Schulte, a member of Abcam's Scientific Support team. Ina has a MS in Biochemistry and Molecular Biology from the Friedrich-Schiller University in Jena, Germany, and a PhD from the University of Essen-Duisburg in Germany. Before joining Abcam, Ina worked as a postdoc at Cambridge University. Ina's flow cytometry experience includes surface staining, as well as intracellular cytokine staining and troubleshooting for both techniques. ​

Joining Ina today will be Augustine Mzumara, a member of our new products team. Augustine has a MS in Human Molecular Genetics from Imperial College in London. Before joining Abcam, Augustine worked as a research scientist at GlaxoSmithKline, and has a diverse background in molecular biology. Just as a reminder, if you have any questions during the presentation they can be submitted at any time throughout the webinar via the Q&A panel on the right hand side of your screen. Questions will be used during the troubleshooting section at the end of the webinar. I will now handover to Ina who will start this webinar on flow cytometry.

IS:    Thank you, Lucy. Hello everybody, thank you for joining our webinar on flow cytometry. We appreciate your time and hope it's not too inconvenient for you. As Lucy said, in the next hour or so we would like to give you an introduction to this exciting technique. First, I would like to start off with an overview: how flow cytometry can be used, including the basic protocols. Then I will briefly explain the science behind the exposition of data, and finish with the analysis of the results and potential troubleshooting.

So for those of you who are not familiar with flow cytometry, what is it? Flow cytometry is a powerful technique that is used to study individual cells or particles in heterogeneous populations. It allows you to measure multiple parameters rapidly and simultaneously. Furthermore, cells can be separated using a cell sorter. This subdiscipline of flow cytometry is often referred to as fluorescence activated cell sorting, or in short FACS. Due to its versatility, flow cytometry is widely used in basic and clinical research.

The research areas which flow cytometry is used include the investigation of certain cell surface markers, as well as intracellular markers or fluorescent proteins, for example. It also is used in the analysis of the DNA content of a cell and much, much more as you can see from this list. I don't want to read the whole list to you, but would rather like to give you a flavor of what can be done with flow cytometry. Flow cytometry evolved from the analysis of blood cells in leukemia patients, and today it is still used to phenotype cells of the immune system in various experimental settings, such as vaccination studies. Flow cytometry can also be used to characterize cancer cells from a solid tumor, and tumor cells that are maybe circulating in the body. The cells that can be analyzed with flow cytometry it can go from very small like bacteria, up to very large, or even whole organisms. Amazingly, flow cytometry is indeed used to study certain mutants of C. elegans. In these studies, the whole worm is flushed through the flow cytometer in order to check for expression of the marker of interest.

Because flow cytometry is such a powerful technique, it has been and is being used more and more. This is illustrated in this graph, which shows the number of publications mentioning flow cytometry or FACS over the last 35 years, and you can see the number is steadily increasing.

But how do you actually utilize flow cytometry? Flow cytometry is a three-step process of: (A) a proper sample preparation; (B) the preparation of the instrument, followed by (C) the data analysis step. Today, I would like to point out what is most important for each of these steps. This means I will talk about the sample preparation and the staining protocols. I will speak about how to set up the instrument, including the compensation process and which controls are required, and I will briefly mention how data is usually analyzed and displayed.

For flow cytometry it is imperative that you prepare a single cell suspension, and this is because so-called doublets, cell clumps or dead cells in your sample can generate misleading data. In case you are analyzing blood samples, which basically is already a single cell suspension, the blood should be collected in tubes that support your experimental setting. For immunophenotyping, for example, collection tubes with EDTA or citrate as an anticoagulant are recommended. Here I suggest to check with the manufacturer of the collection tube for more details. You also may want to consider the lysis of the red blood cells. If you're analyzing cell culture cells or tissue samples, you can use enzymatic methods or a tissue homogenizer to prepare the single cell suspension. In general, it is useful to filter all samples using a 50 µm nylon mesh. Also, to reduce unspecific signals or staining due to dead cells, it is always advisable to ensure viability of the cells you're investigating. 

The next step is staining of the cells or particles. As for most procedures there are direct and indirect staining protocols, as well as a protocol for staining intracellular proteins. I would like to start off with the direct staining protocol. As mentioned earlier, to ensure viability of the cells it is recommended to keep the cells on ice during the preparation and staining of the procedure. A typical sample contains 105 to 106 cells, which are usually resuspended in a volume of 100 µl in a tube or a round-bottom plate. Depending on the sample preparation procedure, the cells should be washed and this is usually done using a buffer-like PBS that includes serum of BSA. Then you spin down the cells and remove the supernatant.

The next step is a blocking step. As for many other protocols, this can be done by adding a buffer with 1-10% serum of BSA to the cell palette. To improve the blocking, the blocking reagent can also be included in the dilution buffer of the primary antibody, which in most cases might be sufficient. Again, this step is followed by washing and centrifuging the cells.

The third step is incubating the cell palette with the primary antibody. The optimum concentration of the antibody will need to be determined by titration. The aim with this is to obtain the maximum contrast between the specific signal, and the background staining. In this direct staining protocol, the primary antibody is fluorochrome conjugated. The most frequently used fluorochromes are the fluorescein-based dye FITC, and phycoerythrin or PE; but, as you can see, there are many, many more available. In general, the sample is incubated for 15-45 min at 4°C, and in order to protect the fluorochrome from photobleaching in the dark. Again, this step is followed by washing and centrifuging the cells.

The following step, the fixation is an optional step, as there are many pros and cons for the fixation of cells. An advantage is the prevention of infection, for example, with contagious diseases from patients whose blood samples are investigated. Fixation can also be convenient as fixed samples need not be analyzed immediately after staining, but can be stored for some time in the fridge, for example. The disadvantage of fixing your cells though is that you cannot exclude dead cells, as the fixation process kills virtually all cells. Fixation also alters the light-scatter profiles and may increase the autofluorescence, so you might need to adjust the instrument settings. If you decide to go ahead and fix the cells, you can either use alcohol such as ethanol or methanol. These reagents can work for most protein antigens, certainly they are the fixation reagent of choice for DNA measurements. You could alternatively perform fixation with formaldehyde or paraformaldehyde, although this is a cross-linking fixative it generally works well with most antigens.

Finally, before running the sample on the flow cytometer, it needs to be resuspended in a minimum of 250 µl of buffer, depending on the type of assay you're performing or the machine you have available. To avoid fading of the fluorescence, keep the stained cells in the dark stored at 4°C and analyze as soon as possible. Also, remember to vortex the sample immediately before the run to ensure an evenly mixed single cell suspension.

Now, this was the direct staining protocol for flow cytometry; now I would like to briefly talk you through the indirect staining protocol. This protocol starts off with the same steps as the direct protocol, so first you would prepare your single cell suspension, and secondly comes the blocking of the samples, if required. The third step is, again, incubation with the primary antibody. In contrast to the direct staining protocol, the primary antibody in this case is not labelled with the fluorochrome. It may be conjugated to biotin though, as illustrated in these pictures.

Following a washing step, the palette is then incubated with a secondary antibody. This is the antibody which is labelled with the fluorochrome, such as FITC or PE. If you have used a biotin-conjugated primary, you would need to incubate with the fluorochrome-conjugated avidin, or streptavidin. The following steps are as before: after washing is an optional fixation step, before running the sample on the flow cytometer.

Different to the direct and indirect staining protocol is the intracellular staining. In order to make intracellular proteins accessible for the antibody, a fixation and permeabilization step is required. For this, either a membrane solubilizer such as saponin or digitonin can be used. As these induce a reversible permeabilization, they should be added to the dilution buffers which are subsequently used to maintain the permeabilization of the cell. Alternatively, detergents such as Triton or Tween can be used, and these induce an irreversible permeabilization. If you would like to analyze any kind of cell surface marker in combination with an intracellular staining, we would recommend to perform the third surface staining prior to the fixation and permeabilization steps. When performing this step, please keep in mind that the light-scatter profiles can change considerably and you may need to adjust the instrument settings accordingly. 

We now have prepared our precious samples - what comes next? The next step is to set up the flow cytometer. For those of you who are not familiar with the technique, the flow cytometer does not come ready-to-use out of the box. It is the type of your sample, actually, that dictates which setting to use. It's really a very important step, because you don't want to waste your precious samples just because the settings were incorrect. What kind of settings are these? What kind of measurements or parameters are required with the flow cytometer?

Generally, there are two types of properties measured: first, there are the structural properties. These include the size of the cell and its cytoplasmic granularity or complexity, as it is often known. In order to measure these properties, no reagents or probes are required. On the other hand, properties which are related to the function of a cell can be measured. These properties include the expression of cell surface or intracellular marker, or the periodic flux of calcium. It also includes the RNA or DNA content of a certain cell. In order to measure these properties, reagents or probes are required and that's why we stained our samples with a fluorochrome labelled antibody. 

How exactly are these properties measured? How does the flow cytometer actually work? The great advantage of a flow cytometer is that it analyzes one cell at a time. As the name 'flow cytometer' indicates, there is a constant flow of a so-called sheath fluid. Into this flow the sample is injected, in this case coming here from the top.

A process called hydrodynamic focusing occurs, which ensures that the cells are passed through the flow cell in single file, as indicated down here. Each cell in turn enters a laser beam, coming here from the side, and two signals are generated. One signal is the fluorescence that is emitted from the stain cells; this signal is passed on to the fluorescent detectors. The other signal is the forward and side scattered lights coming from all cells. This signal is passed on to the scatter detectors.

What do these different signals actually tell us? First, to the forward and side scatter. As mentioned before, as the cell enters the laser beam light is scattered in different directions, as can be seen here in this schematic, the beam coming from the bottom hitting the cell, and the light is scattered. This can be collected and measured to give information on cell characteristics. To be more specific, the direction of forward scattered light correlates to the cell size; the direction of side scattered light correlates to the granularity or complexity of a cell. Please remember to collect the correct signals, please keep in mind that the sample type dictates which settings to use on your flow cytometer.

So far to the dry theory, how does that actually look like with a real sample? Using this information on the physical or structural properties of the cell that is based on the forward scatter and side scatter readings. Cell populations can be distinguished from each other due to different size and complexity characteristics. The results are typically displayed as a dot plot, as for this scatter profile of a human blood sample. So here we have on the X axis, usually the forward scatter that is the cell size. On the Y axis it's usually the side scatter relating to the complexity of the cell. Each single dot represents an event that has been analyzed, and in this example these cells seem to cluster in three different populations. A population of relatively small cells with little granularity here in the bottom left corner in green, and two populations of cells with a similar size here in red and in blue. The cells in the blue population seem to have a higher intracellular complexity compared to the red population.

Detailed analysis showed that these three populations represent different blood cell types. The green population are rather small lymphocytes, whereas the red and blue populations are monocytes and neutrophils, respectively. So this can help us to distinguish different cell populations from each other, but what can you do when you analyze cells of a similar size or complexity? So if you're interested in the expression of a certain protein in a subpopulation of your culture, then you would stain your cells with a fluorescent dye. This raises the question: what is fluorescence? 

Fluorescence is the emission of light by a fluorochrome that has absorbed light of a different, to be precise, shorter wavelength. Light consists of different wavelengths, as can be seen in this diagram, blue light has a shorter wavelength as red light, some 400 nm compared to 700 nm. You can observe these different wavelengths of light every time we see a rainbow.

The individual flow force of fluorochromes that we use in our experiments have unique and characteristic spectra for the absorption of light, and emission of light. Important to know for each fluorochrome is first the excitation wavelength. This is a signature wavelength at which the molecule is excited by the force of light. Second, the emission wavelength, and this is a signature wavelength at which the fluorochrome emits a photon. So in this example for phycoerythrin, the molecule is excited by the blue laser at 488 nm, as indicated by this white line over here. This is close to one of the excitation maxima indicated by the dashed line, and the solid line shows that PE emits light at a wavelength of 578 nm, as indicated by the peak of this solid line over here. 

As mentioned before, there's a huge variety of fluorochromes available. In general, they are of two different natures, they are either synthetic organic dyes such as FITC, the DyLight family, the Cy dyes, or the Alexa Fluor family, or they are fluorescent proteins such as PE, APC, or GFP.

How are all these fluorescent signals detected in a flow cytometer? This is where I have to go just a little bit into the technical details of a flow cytometer. So this is the setup of a standard four-color flow cytometer. If you remember, earlier I showed you the flow cell and the fluidic system through which the sample is passed here in the bottom left corner, and the laser beam coming from the side over here. The scattered light and fluorescent signals, which are generated here are then split into defined wavelengths and channeled by an array of filters and mirrors, in short the optics. Individual fluorescence is then picked up by so-called photomultiplier tubes or PMTs, and is then, in turn, converted into, in most cases, a digital signal. These are the data which are analyzed on the computer with post-acquisition software.

However, when you have used the flow cytometer already, you may know that this is not all. As mentioned before, the sample type dictates the settings of the flow cytometer. This is particularly true when using more than one fluorescence. During the analysis of data from two fluorochromes, you need to compensate for an overlap of the emission spectra of two fluorochromes; the process, which is called compensation.

Let me explain this with the most frequently used fluorochromes: FITC and PE. So this is a very detailed image, but I will talk you through it. For the moment we can only focus on the solid line, which are the emission profiles. You can see that FITC has got an emission maximum at 520 nm, but it also emits light up to a wavelength of 650 nm, as shown by this tail over here. In contrast, you can see that PE has got an emission maximum at 578 nm, as indicated by this peak here. If you now detect the signals for PE between, let's say, 550 and 620 nm that is around the emission maximum of the PE. You not only detect the signal for PE, but you also obtain a signal that comes from the PE over here, in this range of the wavelength. Therefore, you need to compensate for this so-called spillover by subtracting the fluorescent signal, in this case the FITC, that has leaked into the wrong detector, in this case the PE detector. As there is a huge number of different flow cytometers available and we cannot cover all the details of every machine here, I would like to refer you to the manufacturer's instructions for details.

In order to perform this compensation and also to setup the instrument properly, a number of controls is required. For example, an unstained or negative control is used to determine background staining or autofluorescence, and to setup the instrument. Controls state with only one of the fluorochromes used are required for compensation. Just like for other applications, isotype controls, no-primary controls and positive controls are used to confirm specific binding, and to exclude non-specific binding of the antibodies used. You may find more detailed information of the required controls on our website, and other relevant sources. So having prepared our samples properly and setup a flow cytometer appropriately, the final step, the data analysis should be a piece of cake! Data analysis is facilitated by a huge variety of post-acquisition software that are currently available, and which enables you to display the data in many ways. Most commonly, flow cytometric data is presented as a dot plot. Variations of this kind of plot are the control plot or the pseudocolor plot, as can be seen in the following example.

Here we have the results of a staining for CD4+ and CD8+ T cells in human blood, using our antibodies ab86886 and ab86891. On the X axis the fluorescence intensity for FITC, in this case, labelling the CD4+ T cells is shown. Whereas on the Y axis the intensity for PE, labelling the CD8+ T cells is indicated. In this plot there are four more or less distinct cell populations, in the lower left is a population of cells that reveal little or no green or red fluorescence; these are the CD4 and CD8 negative cells. Along the X axis there's a population of cells that exhibit an increased green, but no red fluorescence; these are the CD4+ T cells. Accordingly, along the Y axis there's a population of cells that exhibit an increased red, but no green fluorescence; these are the CD8+ T cells. Finally, there's a small population of cells that seems to reveal both green and red fluorescence, which correlates to cells expressing both the CD4 and the CD8 molecule.

Flow cytometric data can also be displayed as a histogram, as in this example here. For a histogram the fluorescence intensity, so, for example, the FITC is plotted against the number of cells that are exhibiting the staining, resulting in this curve-like structure. This kind of display is predominantly used for cell cycle analysis. For cell cycle analysis the measurement of DNA is widely used, and histograms allow you to determine the DNA content of a cell that is whether it is haploid or diploid. This kind of assay gives you information about the action of cytotoxic drugs, it also allows the characterization of cells such as cancer cells, which are often aneuploid. 

For those of you who are interested in a certain cell population to subculture it, or to use it for subsequent PCA or microarray analysis, a final word to the subdiscipline of cell sorting. How does this work? So the main difference to an analyzing flow cytometer, which I've tried to explain before, is that individual cells or particles are contained in a droplet of sheath fluid once they have past the laser beam. So, here again, we have got samples coming from the top, entering the laser beam and then being in individual droplets. This droplet is given an electronic charge and is in turn attracted or repelled by the deflection plates, and sorted into appropriate collection tubes down here. Nowadays, most cell sorters can sort up to four populations simultaneously, and this sorting is more specific than magnetic beads. To keep it short, the take-home message for cell sorting is if you can see it on an analyzer, you can sort it on a sorter. 

So far so good, but what can you do when the results are not satisfactory? Typically, poor results are of two kinds, either you obtain no signal at all or there is a high background staining. No signal indicates a need to optimize the protocol or the defective antibody, or a lack of the target in the sample. When you observe this, check if your antibody is specific for the species you are investigating. You could also titrate the antibody again, as you may need to increase the concentration. We would also recommend using other techniques to confirm the presence of your target.

The problem of a high background staining might be resolved by reducing the antibody concentration, or you could block non-specific interactions of the antibodies with serum or BSA. In order to isolate this problem, isotype controls or no-primary controls can be very helpful.

So I hope I could show you today that flow cytometry is a very powerful technique that continues to evolve rapidly with the development of new fluorochromes and dyes, as well as novel flow cytometers. If you would like to gain more information, I can recommend consulting your local, regional or national flow cytometry society, the web, for example, Abcam's website for more detailed information on protocols, or, if you prefer, a number of books and journals that are out there. Thank you so much for your attention, and thank you also for the questions that you have already sent in. If you have any more questions, please feel free to type them in the Q&A panel on the right hand side of your screen. I will try to answer them in a few moments, but first I would like to hand you over to Augustine.

AM:    Thank you, Ina. Hello, my name is Augustine and I'm one of the product managers here at Abcam. I'm going to give you a quick summary of some of the products Abcam offers that can help you with your flow cytometry experiments. Before I introduce these products to you, I'd like to mention that at the bottom of the presentation slides you will find links to resources on our website, where you can find out more information about these products.

The first product I would like to introduce to you is our multi-color B and T cell marker panels. These are comprised of monoclonal antibodies directly conjugated to a spectrum of fluorophores. They can be used with a wide variety of instruments, and are useful for multicolor characterizations. The fantastic thing about these products is that they're a great and convenient starter kit if you're new to flow cytometry, as the antibodies have already been optimized for flow cytometry and you get everything in one nice, tidy and convenient package.

The next product I'd like to introduce to you is our EasyLink antibody conjugation kits. These are a vital tool for direct staining where you cannot find a conjugated primary antibody, which you require for an experiment. EasyLink kits are great and with a hands-on time of only 30 sec. They allow you to rapidly conjugate your antibody to your label of choice, and are a cost-effective solution to having the primary antibody you need for your experiment. Abcam offers a variety of fluorescent conjugates relevant to flow cytometry. However, we have other conjugates avidin and biotin available for other types of experiments as well. One thing to know with antibody conjugation is that it is dependent on the purity and concentration of your antibody sample. We also offer purification and concentration kits to help with optimizing your antibody samples for use in flow cytometry.

If you plan to do a one-off experiment and don't have a directly labelled primary, or it's not cost-effective for you to use a conjugation kit, we would also recommend one of our secondary antibodies. Abcam has over 2,500 secondary antibodies in our catalogue. For flow cytometry we would recommend our pre-adsorbed range, and if you're using Fc receptor-rich samples, we also offer a large selection of F(ab')2 fragments. An additional benefit of using one of our secondary antibodies is that you may find conjugates available in the secondary range, which may not be available with the EasyLink. At the moment, you can also take advantage of a special offer which we are running, where if you place a secondary antibody purchase before the 31st October 2011, you can also receive our free IHC Methods Express book. 

Abcam's catalogue also includes a range of small molecule or the biological dyes, which include DRAQ5, Cytrak Orange and DRAQ7. DRAQ5 is a far-red fluorescent dye with applications in cell cycle analysis, as Ina mentioned earlier. The dye only labels double-stranded DNA eliminating the need for using RNAs, which could shorten your experiment and save you a lot of time. Cytrak Orange is an orange fluorescent dye with dual staining, and can be used as a live cell gate in nucleated cells, and for detecting rare events. DRAQ7 is a far-red dye, which labels permeabilized cells which makes it useful for dead, apoptotic cell exclusion as a viability gate. All these dyes are compatible with a variety of flow cytometry systems, and they also offer the advantage of having very low photobleaching. The majority of the dyes are also compatible with FITC and PE. As I mentioned, there's further resources available for all of these products on our website, and you can find the links to these at the bottom of these slides in this presentation.

As a little thank you for attending this webinar, we'd also like to offer you a special promotion where you can get 25% off the following products: any product in the EasyLink antibody conjugation kit range, any product in our Annexin V assay kit range, DRAQ5, DRAQ7 and Cytrak Orange. If you'd like to take advantage of this promotion, please email us. You can also benefit from a further 25% off in the future, if you send us back data from the initial promotion for any of the products that you may have used. To find out further details about this promotion, please visit our website. Before I hand back to Ina for your troubleshooting questions, I'd like to thank you for your time and to wish you a pleasant day.

IS:    Thank you, Augustine. Hello again and a big thank you to all the participants who have submitted questions during the presentation, and we've received really a lot of questions so far. I will try to go through a few of them now. The first question we had by Sandra, and she wanted to know: How should I perform the lysis of red blood cells? Well, the lysis is usually done using a buffer with ammonium chloride, and this leads to basically the swelling or the bursting of the erythrocytes and thereby you can get rid of them. I would recommend to prepare this buffer freshly, and take care not to treat the cells for too long, as other cell types can be affected as well. Personally, I made the experience that reagents used for fixation or permeabilization also lead to the lysis of erythrocytes and can thereby also be used.

We've got a question - quite interesting - from Richard; Richard asks: Can I use flow cytometry for the detection of proteins that are secreted? Of course, Richard, you can; I did it myself. As I said earlier, flow cytometry has evolved or is frequently used in the immunophenotyping, and immune cells often secrete cytokines and you can measure these by treating the cells with a reagent that prevents the secretion of the cytokines, so the proteins of interest. Usually brefeldin A or monensin is used for this, and these are so-called Golgi blocks and, or brefeldin is Golgi block, and that leads to the accumulation of the protein inside the endoplasmic reticulum, and with this you then can perform an intracellular staining to detect the protein of interest.

I think I'll pick the question from Michelle; Michelle wants to know: I am using DyLight 488 and PE for my protein of interest, can I include DRAQ7 to gate for the live cells? Yes, Michelle, that is a very good choice, actually, as Augustine just pointed out, DRAQ7 stains the double-stranded DNA of dead cells, so you can use it to gate on live as well as dead cells in flow cytometry. The emission maximum is in the far-red for DRAQ5, so somewhere beyond 650 nanometers, and therefore it is compatible with the 488 in the green range and the PE in the yellowish light, if you like.

This is more or less a troubleshooting - Rachel has a problem: I did an intracellular staining and obtained a high background staining, what can I do? Well, that somewhat depends a little bit on your protocol, what you have done already. In general, you can try to wash the sample again, that may help already to reduce the background staining. You could also include a blocking reagent, such as serum of BSA into your wash buffer, or also into the dilution buffer of the antibodies to reduce the background staining. It depends on the dye that you have used, so for intracellular staining I would not recommend to use tandem dyes or large proteins, as they probably cannot enter the cell and may give an unspecific staining.

We have a question by Molly; Molly wants to know: My protein is not abundantly expressed in the cells I'm investigating, which dyes would you recommend? That is a rather short answer, so for rare targets, so proteins that are not highly expressed in your cells, I would always recommend to use bright dyes such as PE or APC to make the signal stand out.

Then we have Derek: How long can I store my samples before running them on the flow cytometer? So this is a question of practical relevance. In general, I would suggest to analyze the samples as soon as possible, so the sooner the better. For unfixed cells it would be definitely within 24 hr also for fixed samples. If you're performing DNA measurements that could be stained, stored longer up to several days, I would say. Maybe also a little bit depending on the dye that you have used.

Then we have Jan; Jan wants to know: Can I use flow cytometry to determine the absolute size of cells? Not really, so flow cytometry is powerful, but unfortunately it does not give you the real size of the cells in µm, for example. So you may refer to a scatter plot, but I've shown you earlier and there you could see that the forward scatter or the signal that is given by the forward scatter is given in arbitrary measurements, so it's not really related to the actual size of a cell. 

A final question. Mark wants to know: How can I study apoptosis? So I think there are many ways, because this is such a complex process you could use flow cytometry to detect the light scatter or the changes in light scatter profiles of the cell that are undergoing apoptosis, so they change their sizes and contents. You could check for the DNA content of the cell using DNA dyes such as, for example, DRAQ5, as Gus just mentioned. Or you could perform an Annexin V staining, which is specific for cells undergoing apoptosis. So I think you could use a bunch of approaches to analyze this with flow cytometry.

I think we are approaching the end of our webinar. Unfortunately, I cannot go through all the questions you had today, but for those of you whose questions I have not mentioned I will be contacting you within the next few days. So from me it's a danke schoën and auf wiedersehen, and I will now pass you back to Lucy who will provide some final webinar details, including where you can download the presentation I've just given.

Thank you, Ina. On behalf of Abcam, I would like to thank you for attending this webinar. I hope you found it informative and useful for your work. The presentation just given is available for download. When you log-off from the webinar you will be directed to a webpage where a PDF of the presentation can be found, along with information about the exclusive promotion for all webinar participants. If you have any questions about flow cytometry, or have any scientific enquiry, our scientific support team will be very happy to help you, and can be contacted at If you have any webinar or event-related questions, please contact the Abcam events team at I hope we can welcome you to another webinar in the future. Thank you again for attending and good luck with your research.