Flow Cytometry Channels

Flow Cytometry Channels

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I did a flow cytometry experiment for the first time. I had three conditions: (1) unstained healthy monocytes (2) healthy monocytes stained with green/red (488/570) from the Live/Dead thermofisher kit (3) dead monocytes stained with the same green/red dyes.

I did those condition to know what graphs to expects, and learn the FACs process and analysis. Would appreciate it if someone can answer the following questions:

  1. So I have two dyes in my samples and a control. Which flow cytometry channel should I use in my plots (FL1, FL2, FL3, or FL4)?

  2. How to know the percentage of live and dead cells in each condition?

  3. Anyone knows a free analysis software?
  4. How practical is R to do flow cytometry analysis?

I'll amend this answer as I get details from you.

What is your cytometer, and how are the optics set up? You typically have 1-5 lasers: UV (355nm), violet (405nm), blue (488nm), green/yellow (~561nm) and red (~633nm) lasers. Your FL number, FL1-4, typically refer to a particular PMT (photomultiplier tube) and filter combination. For example, a typical FACSCantoII has a 4-2-2 setup with 4 PMT on the blue laser, 2 on the violet laser and 2 on the red laser (if you bought the device that way, there are other configurations). These fluorescence channels are set up to detect a particular fluorophore or equivalent with a bandpass filter. The base filters on FACSCanto II red laser are 660/20 and 780/60. These filter light emitted by the stained cells at a particular range of wavelengths, and you read them as 600nm +/- 10nm and 780nm +/- 30nm, for example.

You can set your software up to be FL numbers or fluorophores, but what you need to know is what filter is FL1 and so forth? Typical cytometers like FACSAria, these are FL1 = FITC, FL2 = PE, FL3 = 7AAD, PerCP/Cy5.5, etc., FL4 = APC.

So you used a live/dead thermofisher kit, please provide us also the catalog numbers.

What we want to do is take the information about your dyes and about your cytometer and reconcile them to determine what channel they should fluoresce in. 488nm looks like FITC or FL1 and 570nm looks like PE or FL2, but these look like lasers, whereas you want to tell me the dye emission to match it to a channel, using the information about your device configuration!

As for software, there are a number of good, expensive options like FCS Express and FlowJo but they're costly. See if your core or what have you has these available for you to use on-site. Otherwise, the on-board cytometer software like FACSDiva may need to suffice, and often does. I've used some of the "free" ones but they're very clunky and/or terrible to use.

R has some usefulness for high-dimensional analysis, but you need to be able to use R fluently, and you can't draw manual gates. There do exist helpful packages with good documentation like X-cyt (broad institute), though. And for visualization like PCA and t-SNE, R has a lot of documentation for flow data. Also consider R loads everything into RAM so on a slower 2-4GB computer it might have lackluster performance.

To start with your analysis, I'll ask why did you use two dyes? The LIVE/DEAD product line are reactive with free amines which present inside and outside the cell. Dying cells have more permeable membranes and take up more dye. So when you analyze the cells, everything is positive, but the dead and dying cells are 1-2 logs higher intensity of staining compared to the live population.

So the first thing any flow experiment needs are controls. You have unstained controls, FMO controls and compensation controls. The unstained just gives you an idea your morphology by light scatter, but for low-dimensional experiements can act as an FMO. The FMO or fluorescence minus-one is a cell stained with everything in your antibody/dye panel except one marker, so that you can compare against the fully-stained sample and see where to draw your gate. Compensation controls help the cytometer correct for the fact that filters pick up emission from other fluorophores than the target. You also have a biological control, which is good, because for analysis you always want to see where a clear positive and negative population lay on your dotplots or histograms. Live cells help you see where the positive and negatives are but may have a weak positive signals, and unstained cells will have a negative population that is shifted down 1/2 log or so because they dont have any dye at all. The dead stained cells will show you a strong positive signal and potentially a weak negative signal. Use these to draw your gates.

So then compensate the data electronically with your software (follow the manual), and create a dotplot with FSC-A and SSC-A on the axes. Only experience will tell how your cells look on this plot, but for reference, a blood or leukocyte sample would like like this:

You want to draw a gate around your monocytes and then create a plot based on your monocyte gate (at least gate out debris). It's also useful for further experiments to stain with a monocyte marker so you can dump everything but your target cells, so use CD14, for example, which is specific to monocytes.

On the next dotplot, make the axes FSC-A and FSC-H. This is a doublet discimination step where laser area scaling done before a cytometer run helps a lot (see your core expert). The principle here is to gate out (get rid of) cells that are stuck together:

Then you would want to look for your target cells. If they're both live/dead dyes you're going to have the same signal on two different channels, so you can probably just pick one, like live/dead green.

You can do a histogram of FL1 based on the cells kept in the doublet plot. You should see two discrete peaks on a log or bi-exponential transformed intensity axis (width basis -10 is good to use, but FACSDiva does it automatically). Width basis is a co-factor that compresses the data around 0, so negative populations get smaller and you get better resolution among the positive populations.

You would draw a bifur gate between the positive and negative peak or a range gate around the positive, and select tell the software to generate statistics to get your percentages. And you can repeat for the FL2/red dye to get those percentages, too, but both markers are looking at the same thing.

Then, apply those gates to all your samples and adjust the gates based on what the data looks like.

Please try this presentation:

4 Biggest Mistakes Scientists Make During Multicolor Flow Cytometry Cell Sorting Experiments

Multicolor cell sorting is a complicated process and certain scientific errors can be common.

Unsuccessful multicolor sorts can result in erroneous data and inconclusive results. Successful multicolor sorts, on the other hand, can give excellent results and lead to dynamic conclusions.

Successful multicolor cell sorting requires special attention to planning.

Using specific setup strategies for your experiment can create a streamlined system for an otherwise complicated process. For example, these critical steps and strategies for multicolor sorting experiments can save you time and maximize your results.

When setting up a multicolor experiment, the most common mistakes are failing to set PMT voltages properly, failing to use a viability dye, failing to address doublet discrimination properly, and failing to set the right sort regions and gates. Eliminating these 4 mistakes is important for any kind of flow cytometry experiment, but particularly for flow cytometry cell sorting experiments.

The following 4 mistakes should be avoided prior to the setup phase, which should be executed immediately before the sort. This setup phase should be included as part of the planning, optimization, and trial process of the experiment to give you the best cell sorting results possible.

Here are 4 common multicolor cell sorting mistakes you should avoid…

Flow cytometry in human reproductive biology

Flow cytometry (FC) is an analytical cytology technique which has been extensively used for decades. It has many advantages compared with other similar methods for the study of cell biology, even on a molecular basis. FC allows the cell-by-cell analysis of many optical or immunological features in the same sample, at the same time, and at a rate of thousands of cells per second, generating immense quantities of data and thus providing almost limitless information which is statistically robust due to the number of units studied. The aim of this review is to describe the contribution of FC to the study of physiological and pathological processes related to human reproduction, and to discuss how this technique has been used in research, as well as its clinical applications in this field. We have used some practical examples selected from the most relevant studies within a wide range of investigations published in the literature, and we have also drawn on our own experience of using flow cytometry to study different phenomena related to reproduction. It is conclued that FC is a useful instrument for basic investigation of gynecological issues, as well as for the study of male reproductive characteristics, either in research applications or directly for clinical diagnostic purposes. Future development of these techniques will permit further advances both in our knowledge and in the improvement of assisted reproduction techniques.

The Difference Between Linear And Log Displays In Flow Cytometry

Data display is fundamental to flow cytometry and strongly influences the way that we interpret the underlying information.

One of the most important aspects of graphing flow cytometry data is the scale type. Flow cytometry data scales come in two flavors, linear and logarithmic (log), which dictate how data is organized on plots. Understanding these two scales is critical for data interpretation.

Let’s start at the beginning, where signal is generated, and trace its path all the way from the detector to the display.

Behind every flow cytometry data point is what we call a pulse. The pulse is the signal output of a detector generated as a particle transits the laser beam over time. As the cell passes through the laser beam, the intensity of the signal from the detector increases, reaches a maximum, and finally returns to baseline as the cell departs the laser beam. The entirety of this signal event is the pulse (see Figure 1).

Figure 1: The voltage pulse begins when a cell enters the laser, hits its maximum when the cell is maximally illuminated, then returns to baseline as the cell exits the beam.

This is all good, but an electrical pulse is not useful to us in and of itself. We need to extract some kind of information from it in order to measure the biological characteristics we are seeking. This is where the cytometer’s electronics (which contribute significantly to a particular cytometer model’s performance heft and price tag) come into play.

Modern instruments employ digital electronics. This means that the signal intensity over the course of a pulse is digitized by an analog-to-digital converter (ADC) before information is extracted from it.

This was not the case in the past, when most systems used analog electronics. In analog systems, the information about a pulse is calculated within the circuitry itself, and is digitized for the sole purpose of sending the data to the computer for display.

Regardless of the instrument, the type of data provided about the pulse is the same: area, height, and width (see Figure 2). These three pulse parameters are what are ultimately displayed on plots.

Figure 2: Three characteristics of the voltage pulse: area, height, and width.

Area and height are used as measurements of signal intensity, while width is often used to distinguish a single cell from two cells that passed through the laser so close together, that the cytometer classified them as one event (a doublet event).

Typically, on flow cytometry plots, you will see the axis or scale labeled with an A, H, or W denoting the pulse parameter being displayed (e.g. “FITC-A,” “FITC-H,” or “FITC-W”).

It is important to note that all of the pulse processing is performed in the cytometer electronics system, not in the computer.

The reason for this is that the required speed for processing can exceed what is possible with the computer and its ethernet connection. Given this, the cytometer passes all of the pulse measurements, already neatly processed and packaged, to the computer and cytometer software that graphs the data.

This is when plot scaling becomes important.

The range of signal levels that the cytometer transmits to the computer is extremely large, and is a function of the cytometer’s ADC. The number of bits of the ADC determines how many values comprise this range of signals.

For example, a 24-bit ADC can divide the range of signals into 16,777,216 (2 24 ) discrete values. (Note that each scatter or fluorescence channel gets its own ADC, so the number of ADCs equals the total number of parameters on the instrument.) Therefore, the dimmest FITC signal on this example instrument can be assigned a value of 1 while the brightest FITC signal can be assigned a value of 16,277,216.

Even though the granularity of each signal is assigned 2 24 different values, this kind of resolution is much too fine to be useful on the scales of plots.

If a histogram’s scale reflected this many values, events would be spread out among so many channels that we would need to collect millions of events to see the peaks and populations we are used to.

Furthermore, computer monitors don’t have the resolution required to draw dots on this scale. Even if they did, the dots would be so small we wouldn’t be able to see them on the screen.

The universally employed solution is to scale down the resolution on plots to a more practical, but still useful, degree.

Instead of dividing the scale into millions of units, we divide it into 256 (or, in some cases, 512) units called channels.

For a 256-channel system, we allocate all 16,277,216 digital values equally among the channels, so that each one contains 65,536 discrete values (16,277,216 divided by 256). Channel 1 can contain up to the dimmest 65,536 events, while channel 256 can contain up to the brightest 65,536 events.

This kind of scale is linear because equivalent steps in spatial distance on the scale represent linear changes in the data. As illustrated in Figure 3, moving a distance of x reflects a change of 64 channels, regardless of whether the starting point is channel 0, channel 64, or channel 192.

As such, the key feature of a linear scale is that the channels are distributed equally along the scale: the distance between channel 1 and channel 2 is the same as the distance between channel 100 and channel 101.

Figure 3: On a linear scale, channels are spaced equally.

Linear scale is certainly nice, but what happens if two populations, with very different levels of intensity, must be plotted together? This is a common situation in flow cytometry, in which nonfluorescent cells are visualized on the same plot as brightly fluorescent cells.

In this case, a plot with linear scaling becomes much less useful, as it will be very difficult to see both fluorescent and nonfluorescent cells at the same time, no matter what PMT voltage we use. Either all the nonfluorescent cells will be crammed into the first few channels, or all the fluorescent cells will be crammed into the top few channels.

This is where a logarithmic scale comes into play.

A log scale is one in which steps in spatial distance on the scale represent changes in powers of 10 (usually) in the data.

In other words, moving up a log scale by one quarter of the scale allows us to move from channel 1 to channel 10 (see Figure 4). Moving another quarter distance up the scale brings us not to channel 20 but to channel 100, a power of 10.

Figure 4: On a log scale, channels are unequally spaced so that one can visualize both high and low signals on the same plot.

Log scales are really good at facilitating visualization of data with very different medians, and are organized into decades. A four-decade log scale is marked: 10 1 , 10 2 , 10 3 , 10 4 , so it contains 10,000 channels in total.

Importantly, even though each channel itself contains the same number of digital values, data channels are not distributed equivalently across the scale.

The first decade, from 10 0 to 10 1 , contains 10 channels (channel 1 to channel 10). The second decade, even though it occupies the same amount of space on the scale, contains not 10 but 90 channels (11 to 100). And, the fourth decade from 10 3 to 10 4 — occupying the same space as each other decade does — contains a whopping 9,000 channels (1001 to 10,000).

On the log scale, data is compressed to a much greater degree at the high end than it is at the low end, and it is this very property that makes it so good for visually representing data with very different medians (see Figure 5).

Figure 5: Effects of Linear vs Log scaling on resolution of 8-peak beads. The Spherotech 8-peak bead-set was run on a DIVA instrument with either Log scaling (left) or Linear scaling (right). The 8th peak was placed, on scale, at the far right of the plot. As can be seen, without log scaling of the data, the bottom 6 peaks cannot be resolved.

It is very important to keep in mind that in the digital cytometry world, these scales are solely visualization methods and, like a compensation matrix, have no effect on the underlying data. The scales are applied by the cytometry software, not the cytometry hardware.

Incidentally, this was not the case in older analog systems which applied the logarithmic transformation in the cytometer electronics using logarithmic amplifiers, so the data streamed to the computer was already “log transformed” before it got to the software.

At this point, you are probably wondering about the practicalities of these scales: when should you use linear scale and when should you use log scale?

Typically, linear scale is used for light scatter measurements (where particles differ subtly in signal intensity) and log scale is used for fluorescence (where particles differ quite starkly in signal).

However, it is not always this simple.

For most flow cytometry on mammalian cells, the range of both forward and side scatter signals generated by all particles in a single sample is not wide enough to warrant a log scale for proper visualization.

Particle size may range from a few microns to 20+ microns in a typical sample, so the entire gamut of particles would be happily on-scale using a linear scale. In fact, log scale would be counterproductive in this situation, compressing the range and making it difficult to differentiate different blood cell populations from each other, for example.

However, side scatter on a log scale can be extremely informative, especially when measuring “messy” samples with many different kinds of cell types, like those generated from dissociated solid tissues.

Additionally, make sure to use both forward and side scatter on log scale when measuring microparticles or microbiological samples like bacteria. These types of particles generate dim scatter signals that are close to the cytometer’s noise, so it’s often necessary to visualize signal on a log scale in order to separate the signal from scatter noise.

Fluorescence measurements typically involve populations that differ significantly in intensity, and thus require a log scale for visualization. This is the case when measuring signal from immunofluorescence, fluorescent proteins, viability dyes, or most functional dyes.

However, there is a major exception: cell cycle analysis. Cell cycle analysis by flow cytometry is usually accomplished by measuring DNA content via fluorescence. Cells in G2/M contain up to twice the amount of DNA found in other cells, so we need to see relatively small differences in signal intensity in order to assess cell cycle state.

Therefore, cell cycle analysis must be visualized on linear scale.

We hope this explanation sheds some light on scaling. Knowing how to properly display your data is a critical part of scientific communication. Remember to use linear scaling for most scatter parameters, or when you need to visualize small changes, and log scaling for most fluorescence parameters, or when you need to visualize a wide range of values. As always in flow cytometry, there are certainly exceptions, but armed with this knowledge, you should be able to make educated judgements about which scale types to use in various assays and to better interpret your data. Happy flowing!

To learn more about The Difference Between Linear And Log Displays In Flow Cytometry, and to get access to all of our advanced materials including 20 training videos, presentations, workbooks, and private group membership, get on the Flow Cytometry Mastery Class wait list.

Tim Bushnell holds a PhD in Biology from the Rensselaer Polytechnic Institute. He is a co-founder of—and didactic mind behind—ExCyte, the world’s leading flow cytometry training company, which organization boasts a veritable library of in-the-lab resources on sequencing, microscopy, and related topics in the life sciences.

Talk Overview

Dr. Malte Paulsen gives an introduction to flow cytometry with an excellent explanation of the basic principles governing the technique. He explains how fluid flow is used to focus a sample in a laser beam. Light from the laser is scattered by cells in the sample and the degree of scatter provides information about the cell’s optical density and other characteristics. In conventional flow cytometry, lasers are used primarily to excite fluorescent antibodies bound to specific cell types. A detector with different filters allows specific wavelengths to be dissected from the overall fluorescence. This signal can then be displayed in ways that provide the most information about the cell type of interest.

Introduction to flow cytometry

Flow cytometry is a widely used method for analyzing the expression of cell surface and intracellular molecules, characterizing and defining different cell types in a heterogeneous cell population, assessing the purity of isolated subpopulations, and analyzing cell size and volume. It allows simultaneous multi-parameter analysis of single cells.

It is predominantly used to measure fluorescence intensity produced by fluorescent-labeled antibodies detecting proteins, or ligands that bind to specific cell-associated molecules such as propidium iodide binding to DNA.

The staining procedure involves making a single-cell suspension from cell culture or tissue samples. The cells are then incubated in tubes or microtiter plates with unlabeled or fluorochrome-labeled antibodies and analyzed on the flow cytometer.

If you are looking to get to grips with flow cytometry analysis, check out our free online flow cytometry training.


The flow cytometer: fluidics

Figure 1. Overview of the flow cytometer. Sheath fluid focuses the cell suspension, causing cells to pass through a laser beam one cell at a time. Forward and side scattered light is detected, as well as fluorescence emitted from stained cells.

When a cell suspension is run through the cytometer, sheath fluid is used to hydrodynamically focus the cell suspension through a small nozzle. The tiny stream of fluid takes the cells past the laser light one cell at a time (Figure 1).

Light scattered from the cells or particles is detected as they go through the laser beam. A detector in front of the light beam measures forward scatter (FS) and several detectors to the side measure side scatter (SS). Fluorescence detectors measure the fluorescence emitted from positively stained cells or particles.

The flow cytometer: measurement of forward and side scatter of light

Cells or particles passing through the beam scatter light, which is detected as FS and SS. FS correlates with cell size and SS is proportional to the granularity of the cells. In this manner, cell populations can often be distinguished based on differences in their size and granularity alone.

Figure 2. Light scatter as the green laser interrogates the cell. The direction of light scattered by the cell correlates to cell size and granularity.

A useful example of this is when running blood samples on the flow cytometer.

  • Larger and more granular granulocyte cells produce a large population with high SS and FS.
  • Monocytes are large cells, but not so granular, so these produce a separate population with high FS but lower SS.
  • Smaller lymphocytes and lymphoblasts produce a separate population with less FS. They are not granular cells, so also have low SS.

Therefore, these cells can be separated into different populations based on their FS and SS alone.

Figure 3. Dot plot of FS versus SS. Each dot represents a single cell analyzed by the flow cytometer. The characteristic position of different cell populations is determined by differences in cell size and granularity. Image reference: Riley and Idowu. Principles and Applications of Flow Cytometry*.

The flow cytometer: measurement of scattered light and fluorescence

As well as separating cells based on FS and SS, cells can also be separated by whether they express a particular protein. In this case, a fluorochrome is often used to stain the protein of interest. Fluorochromes used for the detection of target proteins emit light when excited by a laser with the corresponding excitation wavelength. These fluorescent stained cells or particles can be detected individually.

Forward and side scattered light and fluorescence from stained cells are split into defined wavelengths and channeled by a set of filters and mirrors within the flow cytometer. The fluorescent light is filtered so that each sensor will detect fluorescence only at a specified wavelength. These sensors are called photomultiplying tubes (PMTs).

Figure 4. Fluorescent light is filtered so that each PMT detects a specific wavelength. The PMTs convert the energy of a photon into an electronic signal – a voltage.

In the example shown in Figure 4, the FITC (fluorescein isothiocyanate) channel PMT will detect light emitted from FITC at a wavelength of approximately 519 nm. The PE channel PMT will detect light emitted from PE (phycoerythrin) at 575 nm wavelength. Each PMT will also detect any other fluorochromes emitting at a similar wavelength to the fluorochrome it is detecting.

Various filters are used in the flow cytometer to direct photons of the correct wavelength to each PMT (Figure 5).

Figure 5. Filters in the flow cytometer. Band pass (BP) filters allow transmission of photons that have wavelengths within a narrow range. Short pass (SP) filters allow transmission of photons below a specified wavelength.

Long pass (LP) filters allow transmission of photons above a specified wavelength. Dichroic filters/mirrors (such as dichroic LP mirrors) are positioned at a 45° angle to the light beam.Sub heading

In a long pass dichroic filter, photons above a specific wavelength are transmitted straight ahead, whilst photons below the specific wavelength are reflected at a 90° angle.

Measurement of signal

As the fluorescing cell passes through the laser beam, it creates a peak or pulse of photon emission over time. These are detected by the PMT and converted to a voltage pulse, known as an event. The total pulse height and area is measured by the flow cytometer. The measured voltage pulse area will correlate directly to the intensity of fluorescence for that event.

Figure 6. The PMT measures the pulse area of the voltage created each time a fluorescing cell releases photons. When no fluorescing cells pass through the optics, no photons are emitted and no signal is detected. As the fluorescent labeled cell passes through the optics and is interrogated by the laser, photons are emitted and so the intensity of the voltage measured increases. As each fluorescing cell completes its path through the laser beam, this leaves a pulse of voltage over time.

The pulse area is calculated by adding the height values for each time slice of the pulse, determined by the speed of the analog to digital converter (ADC), which is 10 MHz (i.e. 10 million per second or 10 per microsecond).

These events are assigned channels based on pulse intensity (pulse area). This signal can be amplified by turning up the voltage going through the PMT.

Figure 7. A one parameter histogram plotting channel number vs. number of events. The channels are usually viewed on a log scale on the x axis.

Each event is given a channel number depending on its measured intensity the more intense the fluorescence, the higher the channel number the event is assigned.

Figure 8. Fluorescence intensity measurements for a negative and positive result. The negative result shown on the left has no staining and many events at low fluorescence intensity. A positive result is shown on the right, this has a large number of events at high fluorescence intensity.

For a positive result you are looking for the shift in intensity between negative control and a positive samples (Figure 9).

Figure 9. Anti-CCR2 antibody (ab21667) staining of human PBMC gated on monocytes. Data is from an anonymous Abreview.

Antibody staining

    Direct staining:

In direct immunofluorescence staining, cells are incubated with an antibody directly conjugated to a fluorochrome (e.g. FITC). This has the advantage of requiring only one antibody incubation step and eliminates the possibility of non-specific binding from a secondary antibody.

This approach is particularly useful for intracellular staining, where large antibody-fluorochrome complexes including secondary antibodies can become trapped causing non-specific binding, or fail to enter the cell preventing primary antibody detection.

In indirect staining, the primary antibody is not fluorochrome-labeled, but is detected by a fluorochrome-labeled secondary antibody. This second reagent may be an antibody with specificity for the first antibody. Alternatively, the avidin-biotin system can be used, whereby an antibody is conjugated to biotin and detected with fluorochrome-labeled avidin.

With the wide range of conjugated antibodies now available, this method means that unconjugated primary antibodies raised against many different targets can be used in conjunction with a labeled secondary antibody for FACS analysis. This widens the choice of target proteins for the researcher.

Staining of intracellular antigens for flow cytometry protocols depends on various fixation and permeabilization methods to allow access of antibodies to internal cellular proteins. A successful staining procedure in all cases is dependent on optimization of experimental conditions through titering of antibodies, use of appropriate controls to set up the flow cytometer correctly and optimized fixation and permeabilization procedures.

Selecting a fluorochrome conjugate

The ability of a given antibody to resolve a positive signal from a negative signal often depends on which fluorochrome conjugate is used.

A general guideline for the relative intensities of the various fluorochromes is, from brightest to dimmest, PE, PE-Cy 7, PE-Cy5, APC, APC-Cy7, Alexa Fluor 647®, Alexa Fluor 700®, FITC, Pacific Blue, Alexa Fluor 488®. This is a general pattern some differences in the relative intensities are seen for individual antibodies.

A highly expressed antigen will usually be detected and resolved from the negative control with almost any fluorochrome. An antigen expressed at lower density might require the higher signal to background ratio provided by a brighter PE or APC conjugate to separate the positive cells adequately from the unstained cells.

The relative fluorochrome intensity depends on the instrument. This is because of differences in the laser and filter combinations used on the different instruments. Be sure to use the appropriate FACS instrument.

BD Biosciences offers a comprehensive portfolio of flow cytometry instruments and reagents for cancer biology research

From specimen collection to sample preparation to cell analysis, BD Biosciences offers a multitude of tools.

Sample collection

The BD Vacutainer ® products family can be used for blood cell and biomarker preservation.

Sample preparation tools

The BD Horizon™ Dri Tumor and Tissue Dissociation Reagent (TTDR) offers gentle and effective dissociation with superior epitope preservation. TTDR maximizes cell yields, while minimizing cell death, which allows effective dissociation of a variety of tumor types to enable single cell studies.

Cell staining, characterization and analysis tools

BD Biosciences offers a comprehensive portfolio of over 9,000 immunology and immuno-oncology–related research reagents that are designed for efficient characterizations of cells.

The dried reagent cocktails of BD Horizon™ Dri Panels are predesigned, ready-to-use multicolor panels optimized and tested for memory T cell, monocyte subset and TBNK cell characterization.

Fluorochrome Marker Clone
APC CD192 (CCR2) LS132.1D9
Fluorochrome Marker Clone
FITC CD197 150503
PE-Cy7 CD95 DX2
BD Horizon™ APC-R700 CD27 M-T271
BD Horizon™ V450 CD4 SK3
BD Horizon™ V500-C CD8 SK1
BD Horizon Brilliant Violet™ 605 CD45RA HI100
Fluorochrome Marker Clone
BD Horizon Brilliant Violet™ 450 CD20 L27
PE CD16 B73.1
PE CD56 NCAM16.2
PerCP-Cy5.5 CD45 2D1 (Hle-1)
PE-Cy7 CD4 SK3

In addition to predesigned panels, our Custom Solutions offer contract manufacturing of multicolor panels in lyophilized, liquid and/or dried formats to minimize the error(s) and time associated with manual cocktailing of reagents, increase reagent stability, and significantly enhance performance consistency.

The BD Horizon Brilliant™ Polymer Dyes were developed from advanced Sirigen dye technology, enabling high-parameter flow cytometry experiments for discerning cell populations. The bright dyes help in distinguishing dim cell populations, such as tumor-infiltrating lymphocytes or cells that have few receptors on the surface from other cells in a sample.

From sorting to analysis, our cell sorters, cell analyzers and downstream sequencing and informatics analysis tools provide efficient solutions for cancer biology research.

Our research cell analyzers, such as BD FACSymphony™ Flow Cytometer, can be used for the simultaneous measurement of up to 50 parameters. The BD LSRFortessa™ System can analyze up to 20 parameters.

Our multiomic solutions include the BD Rhapsody™ Single-Cell Analysis System, which enables high-throughput capture and analysis of hundreds of single cells. Our single-cell multiomics reagents portfolio enables the analysis of the entire transcriptome or of targeted genes of interest. Targeted panels like the BD Rhapsody™ Immune Response Panel allow profiling of specific immune cell types.


To summarize, there is a distinct difference between simply scaling data by changing the relative positions of the axis coordinates and scaling the data by changing the histogram bin-widths. The valley and picket fencing artifact associated with the logarithmic binning both result from the effective resolution of the logarithmic histogram increasing as the channel number decreases. The valley artifact results from the resolution increasing so dramatically that the widths of logarithmic channels become so small that the number of events per channel is too low to see any structure. The picket fencing results from improper graphing of the data in the region where the histogram resolution exceeds the resolution of the instrument's ADC's resolution. New transformations that transition between linear and logarithmic binning help to minimize these artifacts and provide a way to display negative data values. The primary motivation of these transformations is to make the data appear more visually intuitive. The transition parameters of these transformations should be held constant whenever data is compared to avoid plots that are visually misleading solely as a result of differing axis scales.

Flow Cytometry: An Overview

Flow cytometry is a technology that provides rapid multi-parametric analysis of single cells in solution. Flow cytometers utilize lasers as light sources to produce both scattered and fluorescent light signals that are read by detectors such as photodiodes or photomultiplier tubes. These light signals are converted into electronic signals that are analyzed by a computer and written to a standardized format (.fcs) data file. Cell populations can be analyzed and/or purified based on their fluorescent or light scattering characteristics. A variety of fluorescent reagents are utilized in flow cytometry. These include fluorescently conjugated antibodies, nucleic acid binding dyes, viability dyes, ion indicator dyes, and fluorescent expression proteins. Flow cytometry is a powerful tool that has applications in immunology, molecular biology, bacteriology, virology, cancer biology, and infectious disease monitoring. It has seen dramatic advances over the last 30 years, allowing unprecedented detail in studies of the immune system and other areas of cell biology. © 2018 by John Wiley & Sons, Inc.

Keywords: flow cytometry fluorescence light scatter reagents.

Copyright © 2018 John Wiley & Sons, Inc.


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Flow Cytometry Channels - Biology

Description of the Facility
FACS Facility
The Flow-Cytometry Facility is located on the Nineth floor, East wing of the Hunter North building. This Facility can provide analyses of up to 7 parameters in eukaryotic cell populations and sort cells under sterile conditions. These analyses can be used to identify and isolate rare cell populations, determine chromosome ploidy in individual cells, study apoptosis, and study cell-signaling, among other applications.

User training for the FACSCalibur(Becton-Dickinson Biosciences) and assistance in its operation is available, by appointment, through the Facility's Manager. Periodic user training on the FACSCalibur is also provided through specialized seminars.

The FACSVantage(Becton-Dickinson Biosciences) is operated only by the Facility Manager. The Manager and Project Director are available to discuss projects and to advise investigators on the applications relevant to their project.


FACSCalibur Flow Cytometer

FACSCalibur is a bench top analyzer consisting of Argon, air-cooled laser emitting at 488 nm and 633 nm diode laser. It can provide analyses up to 6 parameters and up to 4-colors.

FACSVantage Flow Cytometer

FACSVantage is a sorter consisting of Helium-Neon (HeNe) air-cooled laser emitting at 633 nm and an Enterprise II, water-cooled laser emitting at both 488 nm and Ultra Violet(UV). It can provide analyses up to 7 parameters and up to 5-colors.

Rules of operations
Rules and Regulations

1. Make an appointment to use the equipment at least one week in advance.

2. Equipment use is limited to a 2-hour period per laboratory per day unless special arrangements are authorized. Users must notify the Facility of any cancellation at least 6 hours in advance. For the FACSVantage, a set-up fee will be charged for a missed appointment.

3. Sign the log Book that is placed adjacent to all instruments.

4. After each session on the FACSCalibur, follow the instrument cleaning procedure.

5. The Principal Investigator of each lab will be responsible for his/her laboratory personnel.

6. Report any problems to the manager as soon as possible.

Safety Precautions

1. Lasers

a. FACSCaliubr is equipped with Air-cooled Argon laser emitting at 488 nm and a Diode laser emitting at 633 nm. Although the lasers are enclosed, the light may be visible (with difficulty) if the cover is opened. Do not look at the laser light. It can damage your eyes.

b. FACSVantage is equipped with Enterprise and HeNe water-cooled lasers, which are visible when the instrument is in operation. No one is allowed in the room without the presence of the Facility Manager while the instrument is in operation.

2. Specimens: Depending on the investigator's specimens, one needs to wear an appropriate protective clothing (latex gloves, lab coats, face shield etc.). No materials requiring BL2 should be brought to the Facility. Latex gloves should be used when changing fluids in the FACSCalibur.

Service Schedules and Required forms
FACS Facility Service Charges

FACSCalibur usage: $10.00 per hour

(minimum charge of $10 for each use)

FACSVantage Usage: $20.00 per hour + set-up fee of $20.00 (per day) (1/2 hr. minimum)

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