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There are several methods for analyzing live, dead, and apoptotic cells by flow cytometry. As cells die, the membrane becomes permeable. This allows for antibodies to penetrate the cells, which can now mimic live cells. For this and other reasons, it’s important to remove dead cells from further analysis during your flow cytometry experiments. For example, let’s say you merely need to generate an accurate cell count. If you fail to remove your dead cells first, you might think you’re seeding 10,000 cells, but in reality only 7,000 of your cells are actually viable. Since the dead cells in your sample will not divide, your culture will take extra time to reach the needed level of confluence. Don't make the mistake of forgetting to add a live-dead cell marker to your next flow cytometry experiment. Here are the top 3 markers available to you.

8-peak beads, sometimes called “rainbow” beads, are a set of beads in a single vial that contains 8 different populations that differ only in the amount of fluorophore contained within them. One of the peaks, termed Peak 1, is unlabeled, and the additional seven, termed Peaks 2-8, contain increasing amount of fluorophore. 8-peak beads are designed to fluoresce in all channels on most flow cytometers and cell sorters. These beads are used to check fluorescence sensitivity and resolution by measuring the position of the unlabeled peak and the separation between all of the peaks, respectively. They are also used to check linearity in fluorescence detection channels by correlating the amount of fluorophore on each population of bead with the position on the scale onto which the flow cytometer places the beads. You may have noticed that when you use your 8-peak beads that your peaks have different CVs and intensities - some are wider and taller than others. But do you know why? If not, how do you know if your cytometer or cell sorter is performing correctly? Here's everything you need to know about using your 8-peak beads.

FMO controls are samples that contain all the antibodies you are testing in your experimental samples, minus one of them. When analyzing the minus, or left out parameter in an FMO control, you give yourself a strong negative control to work with. It’s a strong negative control because the left out marker in the FMO control allows you to take into account how the other stains in your panel affect the respective minus parameter. Many flow cytometry gates are difficult to define. This is especially true when you’re looking at activation markers within a continuum or accounting for the large data spread that occurs when compensating a 10+ color experiment. The only way to convince reviewers that your gate is in the proper place is by using FMO controls. Here's why you need to use FMO controls for any multicolor flow cytometry experiment and how to prepare these controls properly.

Electrostatic cell sorting is a complicated process that continues to be improved. It can be a struggle to understand exactly how all of the sorting components coalesce to accomplish the cell sorter's tasks. For many scientists, the most difficult parts of the sorting equation are how droplets are charged and how drop delays are calculated. By understanding these two things, you will be in a better position to set up a successful fcell sorting experiment, which will help you achieve high sort recovery values, allowing for the accurate analysis of your cells and more cells to work with for your downstream experiments. Here's how cell sorting droplets are charged and how cell sorting drop delays are determined.

Flow Cytometry is a remarkably powerful tool for the study of T cells. It has been successfully used for many decades to accurately visualize and enumerate a variety of T cell subsets. With a large sensitivity range for fluorescent probes, >95% sampling efficiency, and the ability to sort populations of interest for further study, fluorescent-based cytometry remains a tool of choice for T cell analysis. The key is to define your T cell populations of interest with correct gating strategies and to back up your T cell subset findings with functional analysis of these subsets. A cell’s actions should guide its definition, not the other way around.

The International Cytometry Certification Exam was developed over a period of several years. The goal was to ensure a base level of flow cytometry knowledge in certificate holders. Some cytometrists have deemed the ICCE as unnecessary. Others have voiced concerns about the specialization of certain exam subsections. However, despite these concerns, the ICCE is here to stay. The exam and certification process as a whole has the support of multiple companies that are providing training, as well as the support of ISAC, ICCS, and the Wallace H. Coulter Foundation. In the end, the most telling test of the value of the ICCE will be when flow cyotmetry job postings in basic research start asking for the certification. This day may not be too far away.

Microvesicles originate from cells and have the same analysis requirements as cells. For these and other reasons, flow cytometry is a popular choice for microvesicle analysis. However, there are pitfalls with small particle flow cytometry that have led to many conflicting publications. The only way to avoid these mistakes is to first identify them and then take measures to prevent them. The following are 4 common mistakes researchers make when preparing microvesicle flow cytometry experiments, as well as how to prevent these mistakes.

All the experiments and experience in the world do not count if you are unable to communicate your results to the scientific community. As part of that communication process, your paper will undergo the dreaded ‘Peer-Review’ process. If you wish your paper to survive this process, you must collect, analyze, and present your flow cytometry data properly—before you submit your paper. A review of the following questions, as well as how to answer them, will help ensure your paper is not rejected. Here are 5 specific questions reviewers will ask when reviewing your flow cytometry data.

To make certain your instrument is set up correctly for your experiments, manufacturers have developed defined polystyrene beads. These beads’ consistent nature helps you to assess how your instrument is behaving, helps you set up proper compensation matrices, and helps you generate volumetric counts of your cell populations. Alignment, sensitivity, and fluidic quality control beads will help you to ensure that with the same wattage on the laser and the same voltage applied to the detector returns the same median fluorescence. The right compensation capture beads will bind antibodies of multiple isotypes from multiple species and give you a very bright positive signal from which you can calculate a correct compensation matrix. The use of counting beads allows you to easily calculate your cell concentration in your sample. Together, these beads will make your life easier and help you get your data published. Here are the 3 beads you should use.

In today’s world, many scientists have access to instruments capable of running experiments with 10 or more colors. The leap from 2 to 10 colors may seem small, but here are many factors to consider in the design and analysis of experiments that makes full use of instruments that can handle these additional colors. Imagine analyzing a 2-color experiment. With 2 biaxial plots and a single quadrant gate, you have only 4 populations to report. Now add a 3rd color. By doing so, you’ve increased your population count to 8. With 4-colors, you’ve increased your population count to 16. On and on it goes until you get to 10-colors. Now you have 1024 possible combinations! With this kind of complexity, careful experimental planning is not a luxury, it’s a necessity. Here are 7 tips for preparing and analyzing 10-color flow cytometry experiments.

Flow cytometrists use the Jablonski diagram to aid in understanding and explaining the kinetic events of fluorescence. Fluorescent compounds start at the ground state until they are excited by interacting with a photon of light. This photon excites the compound, promoting an electon to a higher energy state. Some of this energy is lost by emission of heat and other non-radiative processes, leading to the previous energy state. Finally, an electron falls back to the ground state while releasing a photon of light. This photon has a lower energy (higher wavelength) than the exciting photon of light. Here's how understanding this process can help you get published.

The field of flow cytometry is moving beyond the use of isotype controls, with many suggesting they be left out of nearly all experiments. Yet, isotype controls were once considered the only negative controls you should ever use. They are still very often included by some labs, almost abandoned by others, and a subject of confusion for many beginners. What are they, why and when do I need them? Are they of any use at all, or just a waste of money? Most importantly, why do reviewers keep asking for them when they review papers containing flow data? Here is everything you need to know about using (or not using) isotype controls in your next flow cytometry experiment.