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Flow Experiments Come Alive - Featuring ZE5™ Cell Analyzer, Bio-Rad Antibodies and Reagents
Flow cytometry is a powerful tool with many applications. It can allow researchers to analyze marker expression, DNA levels, cell health, signaling pathways and apoptosis in both single cells and populations of cells to name just a few. Here we will walk you through these common flow cytometry assays, showing you how and when you should perform them with examples of easy-to-follow protocols and results generated with the new ZE5™ Cell Analyzer featuring Bio-Rad antibodies and reagents.
Identification of viable cells is vital in many cell assays. Toxicity or side effects of drugs or treatments can affect viability and therefore the assay result. Alternatively, sample viability can be used to ensure reproducibility of your experiments or to simply ensure that you are studying a live population. The presence of dead cells can greatly affect the quality of your data in flow cytometry as they have increased autofluorescence and non-specific antibody binding, potentially leading to false positives and increased background noise. This can be more of a problem when using frozen samples, building multicolor panels and ultimately may make weakly positive populations harder to identify. Removal or identification of dead cells is useful in most flow autofluorescence and non-specific antibody binding, potentially leading to false positives and increased background noise. This can be more of a problem when using frozen samples, building multicolor panels and ultimately may make weakly positive populations harder to identify. Removal or identification of dead cells is useful in most flow cytometry applications including immunophenotyping, apoptosis, cell cycle and proliferation experiments.
Whilst using the forward and side scatter to gate on cell populations allows debris and some dead cells to be identified and excluded from your analysis, it will not allow complete removal. The use of a viability dye in this situation will allow complete exclusion of dead cells and may significantly improve the quality of your data. This can be seen in Figure 1 where the use of propidium iodide to exclude dead cells leads to significant improvements in the ability to visualize cell populations that are Gr-1 and CD11b positive.
Fig. 1. Use of a viability dye to improve staining.
and side scatter may not be sufficient to remove dead cells. B, dead cell exclusion using propidium iodide allows easier identification of positive and negative populations. Images shown here are mouse bone marrow stained with Rat Anti-Mouse CD11b Pacific Blue (MCA711PB) and Rat Anti-Mouse GR-1 FITC (MCA2387F). Data acquired on the ZE5 Cell Analyzer.
The use of a viability dye will allow a simple calculation of the live to dead cell ratio in a sample based upon the uptake of the dye in dead cells and not live cells, giving you a quick and easy result. The most common viability dyes are nucleic acid binding dyes such as propidium iodide, DAPI and 7-AAD which are excluded from live cells by a viable membrane. Simply adding the dye five min prior to acquisition will be sufficient to stain dead cells. There is a second type of viability dye based upon its ability to bind to free amines present on proteins. Greater fluorescence can be observed on dead cells compared to live cells due to increased content of accessible amines in dead cells again due to the lack of membrane integrity. Whilst these dyes require a 30 min incubation step to label the cells, they have an advantage over nucleic acid dyes in that the cells can be fixed, allowing analysis at your convenience. Furthermore being available in a wider range of excitation and emission spectra, they can be easily added to flow cytometry multicolor panels.
Viability dyes should be a standard addition to all flow cytometry panels to ensure reproducible results.
This is the most common flow cytometry assay and is used for the identification of cells, most often those of the immune system, by their surface markers and increasingly intracellular markers. Populations of cells can be identified by a single marker, such as CD19 for B cells, or using multiple markers including CD3, CD4, CD25 and CD127 for T regulatory cells. This allows the characterization of cell population in a sample, such as B, T, NK and myeloid cells in peripheral blood. An example of simple seven color immunophenotyping, acquired on the ZE5 Cell Analyzer, is shown below in Figure 2, where human peripheral blood has been stained with antibodies to identify T cells, B cells, granulocytes and monocytes.
Fig. 2. Common leukocyte panel with gating strategy. A, seven color panel of CD3, CD4, CD8, CD19, CD20, CD14 and CD66b to identify T cells, B cells, monocytes and granulocytes in human peripheral blood. A, cell populations were gated. B, T cells were identified by CD3 expression. C, helper and cytotoxic subsets were identified by the expression of CD4 and CD8 respectively. E, B cells were identified as CD3-, CD19+ and CD20+. D, myeloid
further identified as CD14+ monocytes, and CD66b+ granulocytes. Data acquired on the ZE5 Cell Analyzer.
The complexity of immunophenotyping panels can be increased to include homing profile, activation state, cellular signaling and cytokine or hormone release to quickly build into complex panels of over 18 colors in a combination of surface and intracellular markers. In addition to basic research these types of panels can be used for clinical diagnostics, disease classification, disease monitoring and evaluation of disease progression.
To successfully immunophenotype a cell population, there are many things to consider and careful planning will help you obtain better results. One of the first considerations is ensuring you have a healthy sample as a single cell suspension at the right concentration, as poor cells will give poor data. The inclusion of a viability dye, as discussed in the earlier section, will allow you to gate out dead cells which can bind antibodies non-specifically.
Choosing the right antibody is crucial as flow validated antibodies may be different clones than those used in alternative applications. Titration is also important to ensure optimal staining. Knowing your instrument configuration of lasers and filters will allow you to choose suitable fluorophores. The more lasers and filters the more flexibility you will have. For example the ZE5 Cell Analyzer has five lasers and can detect 27 colors giving you a wide choice. The best choice of fluorophores to build your panel will depend on many things including fluorophore brightness, fluorescence spillover, cross-laser excitation, antigen density, marker expression patterns, changes to antigen expression in your assay and the number of cells you expect to detect.
If you have an antibody that is not conjugated to a fluorophore, two options are available. You could use a secondary antibody which will give useful amplification of signal. However you may also get amplification of non-specific binding and your multiplexing options may also be reduced due to similar primary antibody species or immunoglobulin isotypes. The alternative is to use an antibody conjugation kit (Figure 3), removing the need for a secondary antibody and reducing the number of wash steps in your protocol, which may be important for samples with rare populations. Bio-Rad offers two distinct types of antibody conjugation kit, with a wide range of fluorophores suitable for microgram to milligram amounts of antibody. They are easy-to-use and require minimal hands-on time.
Fig. 3. Comparison of LYNX Conjugation Kits®
with directly conjugated antibodies. Purified CD4 (MCA1267) and CD8 (MCA1226) were labeled with LNK021RPE and LNK241D650 respectively and used to stain human peripheral blood. As a comparison, directly labeled antibodies were used. The staining shown are lymphocytes gated on the CD3 positive population. CD4 and CD8 positive T cells can be identified in both plots. Data acquired on the ZE5 Cell Analyzer.
In addition to staining with antibodies conjugated to fluorophores, flow cytometry is an ideal application for detecting cells transfected with fluorescent proteins. There are now well over 50 variants of fluorescent proteins spanning almost the entire visible light spectrum. The proteins are often co-expressed, expressed as a fusion or under the control of a specific promoter helping to elucidate cellular processes. Flow cytometry allows quantification of number and fluorescence intensity of transfected cells and can be combined into standard immunophenotyping panels.
Performing the right controls is important for all experiments. Typical controls for flow cytometry include unstained, isotype, Fc blocking, compensation, fluorescence minus one, and biological controls.
Surface staining of samples is a relatively straightforward process involving incubation of sample and antibody in the most appropriate buffer. However intracellular staining will require more steps. The sample has to be fixed to then allow permeabilization. The choice of fixative and permeabilization reagent may have to be optimized or alternatively pre-optimized buffers for intracellular staining can be purchased.
Finally after your cells have been acquired the next stage is analysis. Again we have some simple tips to help you. Plotting the height or width against the area on FSC or SSC will allow you to detect doublets as they will have double the area and width values of single cells whilst the height will be roughly the same. This will help prevent detection of false positives. Sequentially selecting or gating on populations, in histograms or two-color dot plots, that are positive or negative, for increasing numbers of markers, will allow identification of specific cell populations.
To find out more about how to immunophenotype successfully, refer to our flow cytometry resources.
Click on the links below to find more in-depth information on each topic and view our popular flow cytometry basics guide.
Flow cytometry is crucial in the analysis of cell populations to determine levels of programmed cell death, or apoptosis. This highly regulated mechanism for selectively eliminating cells without the inflammation associated with necrosis, plays an important role in embryogenesis, maintaining organ size and removal of damaged or aberrant cells. The importance of apoptosis is underscored by the many diseases such as neurological disorders, cardiovascular disorders, autoimmune diseases and cancer resulting from dysregulation of this process.
Apoptosis can be initiated through the intrinsic and extrinsic pathways and follows distinct signaling pathways (Figure 4). The identification of the pathway and which mediators are activated can be important when studying apoptosis as it may help determine a disease mechanism or allow intervention and potential therapy. Furthermore when combined with surface marker detection, specific cell subsets which are undergoing apoptosis can be identified.
Although an indication of apoptosis can be determined by the use of an assay detecting a single marker, such as the externalization of phosphatidyl serine in the plasma membrane using annexin V, confirmation of apoptosis is usually confirmed using another test such as viability (Figure 5).
Fig. 5. Annexin V detection at the cell surface during apoptosis. Jurkat cells were treated with staurosporine at 1μM to induce apoptosis. The cells were then stained with Annexin V FITC (ANNEX300F) and ReadiDrop™ Propidium Iodide (1351101). Apoptotic cells positive for annexin V can be seen in the bottom right quadrant and dead cells positive for both annexin and PI in the top right quadrant. Healthy cells are negative for both stains. Data acquired on the ZE5 Cell Analyzer.
As cells can die by a variety of processes such as necrosis, necroptosis, pyroptosis or through cellular processes such as autophagy, it is important to confirm that cell death is due to apoptosis. Caspase initiation, whether for specific members or general caspase activation, or loss of mitochondrial membrane potential (Figure 6), are common assays which, when combined with other apoptosis assays, can give definitive results.
Fig. 6. Loss of mitochondrial potential during apoptosis. Jurkat T cells were treated with DMSO (black) or CCCP (red) for 30 min followed by staining with TMRE (ICT946) for 15-30 min. Loss of mitochondrial potential results in a reduction in fluorescence. Data acquired on the ZE5 Cell Analyzer.
Choosing the right apoptosis assay for your experiment can be crucial to obtain the right result and give you the most valuable information. As with immunophenotyping, planning is very important to obtain meaningful results. Knowing the filters and lasers on your flow cytometer will help you choose the most appropriate apoptosis assay, especially if you plan to use them in combination with antibody staining. The analysis of your data should also follow the same procedures as outlined above, removing doublets and carefully gating your sample. Gating strategies may be different to other flow assays, especially with forward and side scatter as cell shrinkage and membrane blebbing may affect the scatter.
Titration of apoptosis reagents and usage of the appropriate controls are key to achieving optimum results. Prior testing on a known apoptotic sample as well as a negative control, including vehicle-treated controls to account for the effects of solvent, may be required to allow optimal instrument and experimental set-up. Necrotic cells are often positive for apoptotic markers due to the loss of membrane integrity. Therefore using a viability dye to exclude necrotic cells from your assay is a crucial step in identification of apoptosis.
Experimental optimization may be required to get optimal apoptosis induction, depending upon the assay required. In addition the order of antibody staining, apoptosis induction and detection may have to be considered to obtain the best results. Don’t forget, compensation controls, fluorescence minus one and unstained controls will still be required to show autofluorescence and fluorophore spillover.autofluorescence and fluorophore spillover.
Whilst flow cytometry is a powerful tool for identifying apoptosis and many antibodies and kits are available, there are other assays and applications such as microscopy and western blotting useful to detect different components of the apoptotic pathways such as members of the Bcl-2 family involved in the intrinsic pathway. Table 1 below summarizes the assays that can be used for detecting the various stages of apoptosis. See our range of specialist apoptosis products including antibodies, kits and reagents.
Table 1. Summary of assays for detecting hallmarks of apoptosis.
Reagents to Allow Detection
Annexin-V conjugates, pSIVA probes
Flow cytometry, microscopy
Signaling cascades initiated
FLICA, caspase antibodies
Flow cytometry, microscopy, western blot
TMRM, TMRE, JC-1
Flow cytometry, microscopy, microplate reader
Hematoxylin + eosin
DAPI, Hoechst, PI, 7-AAD
TUNEL, Sub-G1 assay, DNA laddering
Agarose gel, flow cytometry, microscopy
Phagocytosis of apoptotic bodies
Acridine orange, hematoxylin + eosin
Flow cytometry, light and electron microscopy
To find out more about apoptosis, click on the topics of interest below.
Activation and proliferation of cells provide an effective method to determine immunocompetence and cell reactivity. This can be in response to drug treatment or as part of an immunological response to a pathogen. The strength or lack of a response can be an important indicator of underlying problems or potency. Activation and proliferation can be determined by specific marker staining, calcium signaling or phosphorylation of proteins. However there are specific proliferation assays available.
The simplest method for assessing proliferation is counting cells, if you have more cells after treatment than you put into the assay, you can assume your cells have proliferated. However a quick and easy quantitative method to determine proliferation in cells is to use cell labeling dyes such as CytoTrack™ Cell Proliferation Assay. As seen in Figure 7, with each cell division the relative amount of dye is reduced allowing each division to be measured. Furthermore as these dyes are available excitable by different lasers they can be included in multicolor panels allowing identification of specific subsets within the proliferating populations.
of human peripheral blood lymphocytes. Human PBLs were stained with CytoTrack Red 628/643 Cell Proliferation Kit and stimulated for 5 days with PHA. Cells that have proliferated show a reduction in the amount of dye with each cell division. You can see stimulated cells (in red), labeled cells but unstimulated (in green) and unlabeled cells (in blue). Data acquired on the ZE5 Cell Analyzer.
An alternative popular method for determining cellular proliferation is to use antibodies against BrdU. BrdU is an analog of thymidine which is readily incorporated into the DNA of proliferating cells when added to culture media. The proliferating cells can then be identified using an antibody against BrdU and when incorporated with a DNA dye the relative proportion in each stage of the cell cycle can also be identified.
For more information on BrdU antibodies, click on BrdU link in the Proliferation Resources section below.
Careful planning will ensure you get valid reproducible results. As with all flow cytometry experiments healthy samples are crucial to obtaining good proliferation. The inclusion of a viability dye will allow you to exclude dead cells that have taken up dye but are not proliferating as this may impact on your results. The limitation of this assay is that it does not give you any effector function of the cells. However measurement of specific cytokine release or cytotoxicity assays will provide this information.
The inclusion of negative controls, a fully stimulated control if possible, and unlabeled cells will allow the establishment of a normal range which can be used to optimize other parameters such as dye concentration and instrument settings. If you include other fluorophores, ensure you also use compensation controls. If you use antibodies to identify specific cell populations, make sure you also add Fc blocks, unstained, isotypes and FMO controls.
We have grouped a few simple protocols for T cell stimulation, with some background information to help you choose an appropriate stimulation method. These protocols are useful to determine proliferation responses and may also be used as a positive control for your experiment, although they may need optimization in your assay.
Identification of cells in the different stages of the cell cycle was one of the first flow cytometry assays developed. It is particularly useful for diagnosing cell cycle anomalies or checkpoints and DNA damage. The cell cycle phases can be followed using nucleic acid binding dyes which bind stoichiometrically to DNA. The relative amount of cells in G0 and G1, S and G2 can therefore be determined as cells in G2, which have twice as much DNA as cells in G1, will have double the fluorescence. In addition cells that have >2N amounts of DNA, often indicative of cellular transformation and sub G1 an indication of apoptosis and cell death, can be identified. An example of cell cycle quantification is shown in Figure 8 below.
Fig. 8. Cell Cycle Analysis. Jurkat cells were fixed in 70% cold ethanol, treated with
and stained with PI to reveal the stages of the cell cycle. Doublets were excluded in A, prior to analysis in B. Data acquired on the ZE5 Cell Analyzer.
For DNA analysis the best results are obtained when the cells are fixed in cold 70% ethanol. The cells are pelleted and cold ethanol is added dropwise whilst vortexing to ensure good cell separation. The cells can be kept at 4oC for several weeks at this point.
If you are using dyes that bind DNA and RNA, such as propidium iodide, the RNA can interfere with the staining. It is therefore useful to remove the RNA by incubating the cells with RNAse. As the binding is stoichiometric, the data should be analyzed on a linear scale and often better resolution of the peaks is observed with a slow flow rate when acquiring the data. A crucial part of the analysis is the removal of doublets. Doublets will have a DNA profile of a cell in G2/M or a cell that is >2N, which could skew your results. Cell cycle experiments may require protocol optimization especially when combined with immunophenotyping or fluorescently labeled cells as ethanol fixation can have a detrimental effect on fluorescent proteins and antibody binding.
The ZE5 Cell Analyzer was designed based on real user feedback and developed in collaboration with our innovative partners at Propel Labs. The result is the only flow cytometer on the market that gives you the flexibility to run up to 30-parameter experiments, the convenience of an integrated universal sample loader that accommodates any sample tube or plate format, and the security of Bio-Rad's world-class technical support and service.
Seasoned flow users will appreciate a sample delivery system that ensures quick processing and minimal sample loss and allows you to reuse sample for downstream experiments. Users new to flow will find the ZE5 Analyzer and Everest™ Software easy to learn and use. And all users will fall in love with the ZE5 Cell Analyzer's fast electronics and short laser transit time that enable run speeds of up to 100,000 events per second with no loss in data, perfect for analyzing rare cell events.