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Cell cycle seems like such a straightforward assay. At its heart, it is a one-color assay and should be a simple protocol to follow. However, as discussed before, fixation and dye concentrations are critical. Once those are optimized, it becomes important to run the cells low and slow in order to get the best quality histograms for analysis — the topic of another blog. Adding the critical CEN and TEN controls will help standardize the assay, and ensure consistency and reproducibility between runs while helping identify non-standard (aneuploid, polyploid) populations from normal ploidy. Trying to isolate and focus on specific components of the cell cycle can be done by addition of specific antibodies or using thymidine analogs. In the end, cell cycle analysis is a simple assay that has a great deal of potential. With work and optimization, a great deal of information about the life of a cell can be extracted.

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Cell cycle analysis appears to be 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 amazingly informative assay.

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With the added emphasis on reproducibility, it is critical to look at every step where experiments can be improved. No single step makes an experiment more reproducible, rather it is a process, making changes at each stage that leads to reproducibility. Antibodies comprise a critical component that needs to be reviewed. As Bradbury et al. in a commentary in Nature pointed out, the global spending on antibodies is about $1.6 billion a year, and it is estimated about half of that money is spent on “bad” antibodies. This does not include the additional costs of wasted time and effort by the researcher using these bad antibodies. Using tools to identify the best reagent to use, considering a switch to recombinant antibodies, and properly validating reagents for use in an assay, are 3 steps that will improve the reproducibility of your experiments.

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Flow cytometry is designed to measure physical and biochemical characteristics of cells and cell-like particles using fluorescence. Fundamentally, any single-particle suspension (within a defined size range) can pass through the flow cytometer. Beads, for better or worse, are a sine qua non for the flow cytometrist. From quality control,to standardization, to compensation, there is a bead for every job. They are important — critical, even — for flow cytometry.

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Good flow cytometry depends on a high quality, single cell suspension. If the cells put through the instrument are not of high quality, the ensuing data will be difficult to analyze. Likewise, if the sample is clumpy, one will not be able to readily distinguish cells of interest from the clumps they are attached to. Sample preparation becomes the critical first step in any flow cytometry experiment. To get high quality results, follow these 3 sample preparation steps.

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Compensation in flow cytometry is a critical step to ensure accurate interpretation of data. It is also one of the areas that’s steeped in mystery, myths and misinformation. Manually adjusting the compensation values based on how the populations look, or so-called ‘Cowboy Compensation’, is not the correct way to determine proper compensation. The best practices for compensation involve following some very specific rules. Here are 4 steps to correctly compensating 4+ color flow cytometry experiments.

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T regulatory cells (Tregs), formerly known as T suppressor cells, are a T cell subset with direct roles in both autoimmunity and responses to pathogens.

Tregs decrease inflammation via the secretion of immunosuppressive cytokines (IL-10, TGF-b) and also through direct suppression of inflammatory effector T cells (such as Th1 and Th17 cells). Given the importance of this unique T cell subset in so many immune responses, many investigators feel remiss if they immunophenotype their cell populations of interest without including a Treg measurement in the mix. But quantifying Tregs can be complicated. This article will show you how to quantify Tregs and how to ensure you’re measuring true suppressor T cells.

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When setting up a cell sorting experiment, there are many things to consider.

You must consider which controls you’re going to use, how you’re going to compensate the experiment, which instrument and which instrument settings are ideal, and how you plan to analyze, gate, and present your data. With so many things to consider, it’s easy to lose site of the small things that can drastically affect the viability of your cells, including the composition of your suspension buffer. The composition of the suspension buffer for preparation, staining, analyzing and sorting is perhaps the most important parameter for maintaining viability during a cell sorting experiment. While the precise components of a buffer can differ depending on the cell type, there are a 5 key points to keep in mind.

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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.

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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.

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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.

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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.

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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.

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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.

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Written by Tim Bushnell, PhD Pairing highly expressed antigens (like CD3) with dimmer fluorochromes, and the antigens of interest with the brightest fluorochromes, is a key part of panel design with few tools to help. With early generation instruments, this was relatively easy to determine, since fluorochrome choice was limited. With the advent of instruments…

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