Flow Cytometry - Science method

If it only reaches 13% of your maximum excitation potential, you won't get a good excitation signal, you are risking incaccurate results.

That might be autofluorescent dead cells. You should add a viability stain so that you can exclude the dead cells. There are many amine binding alternative to chose from. You stain with the viability stain before continuing with surface staining, fixation and intracellular staining. ThermoFisher have many fixable viability stains from eBiosciences, Biolegend have their Zombie Fixable viability kits and BD Biosciences have some too.

To differentiate T-regulatory cells (Tregs) by flow cytometry, you need to use a combination of cell surface and intracellular markers that are characteristic of Tregs. Here's a step-by-step guide:

Thank you so much Kais Khudhair al Hadrawi for your answer!

We use Google Calendar and Google Spread Sheets for booking instruments like LC-MS, autoclaves, thermocyclers, q-PCR, and so on.

Just share the like with editing privileges.

Good luck!

High fluorescence in flow cytometry for an Annexin V assay typically indicates a high level of phosphatidylserine exposure on the outer surface of cells, suggesting early stages of apoptosis or cell death

We do Reference Settings every 60 days, which is the BD recommended interval if using BD FC Beads for "compensation". Are you using the BD FC Beads? Comp Beads? We use BD FC beads for 14 fluors. We use comp beads for other antibody/fluorochrome combinations, and some of those are lot specific (especially tandem dyes). Compensation for a few of our antibody/fluorochrome combinations must use cells. For several others we use SpectraComp beads, and for live/dead stain, ViaComp beads (cell mimics, both from Slingshot Bio). I hope this helps a little.

I would suggest using mouse anti-CD3 and CD19 labeled with fluorescence (FL1) for B and T cells, while using a anti-human Fc labeled with a different FL2 to evaluate the pre-exist anti-lymphocyte antibody. It will be a consecutive gating of first gate is SSC/FSC, second gate is FL1/FL2, while double positive cells are what you want.

Best

Thank you very much Hanh Hong Nguyen

Designing a panel in flow cytometry means selecting your fluorochrome-conjugated antibodies so that their signal won't overlap while taking account of your cytometer specification and allowing you to distinguish your important cell populations.

If possible if you know the frequency of your positive population that can also help determine how many cells to add per well. Rare population of cells would be harder/impossible to see if you add too few cells. The numbers indicated above are good, but its good to have an idea (if possible) of your % positive population (out of total live cells)

Thanks, I got this one already. Kind of a classic.

Since you are staining for a surface bound protein marker, trypsin could cleave your protein of interest during the cell harvesting step. So I suggest you use a gentler enzyme like accutase for flow cytometry purposes or just 10mM EDTA solution in DPBS (works for some cell lines). Also the antibody mentioned in the 5th step contains a fluorescence probe right? As I cannot see a secondary antibody staining step. Also the PFA fix is not necessary if you are doing the analysis immediately after staining and are not storing the samples.

It should be single kind of cell type if it was from thp1, so basically you don't have to distinct different population. Just make sure exclude the cell debris then you should be ok.

In general, if antibodies work in tissue (confocal), they should always work in cell suspension (flow cytometry), but not vice versa.

Reasons are below:

1. Epitope availability. Some epitopes on the protein target might be blocked in tissue that cannot be accessed by antibodies using the same epitope. Then you may need to go through the "finding the right clone" process to make it work in tissue.

2. Fluorophore limitation. Many scopes are limited by the choices of laser they have, most don't have UV laser in 350nm, therefore you can't use BUV dye conjugated antibodies. BV dyes can be excited at 405nm but they are large polymer dye that may not be staining complex cell processes and tissue structures as well as small dyes such as Alexa Fluor dyes. Many flow antibodies are designed in these colors and not available in other dyes such as BD antibodies, while many tissue antibodies are limited in colors.

Have a look at FPbase, there you might find what you are looking for.

Hello Cristina Crespo

Not required. Compensation beads are small particles that are pre-coated with antibodies recognizing species-specific antibody light chains, and there is no Fc-binding antibodies involved. So, if you add serum to the compensation beads before staining nothing is going to change. You will be simply wasting mouse serum.

Best.

It is possible that it can be used for this purpose. If I am understanding correctly, you will likely need to search the literature to find an antibody clone which specifically blocks the extracellular binding pocket, search through commercially available clone descriptions about epitopes which may block the binding pocket, or use trial-and-error to test antibodies that could block the GPCR. The one which you describes might work since it is apparently targeting the extracellular region of GPCR, but you'd need to test it to find out. If you test your antibody and find that it does not block GPCR, it most likely means that the antibody is binding another region of GPCR that's farther away from the binding pocket.

can you kindly show the graph? That will aid understanding of the question.

You might try an anti-SRY antibody as for sperm cell sorting; but honestly, I would rather flow sort the various cell populations and do FISH with a Y-chromosome-specific probe.

CD34 for progenitors. but additional markers should be used when define the subtypes.

Be sure to use antibody coated plates or beads for stimulation. Soluble antibodies are not efficient for stimulation (other than for whole blood). Staphylococcal enterotoxin b is also an option, but only a small percent of T cells (in PBMCs) is activated.

That's water and some preservatives. That's all you need. The blue color is optional. Its purpose is just to help you see when all the tubes are filled.

Hi navit,

There are some proteins in cells like YY1 that they have Histidine repeat and maybe its the main reason for background. You can read following article:

Hello Richard, hopefully this STAR protocol would be of use

Analysis of T cells in mouse lymphoid tissue and blood with flow cytometry

Troubleshooting EdU (5-ethynyl-2'-deoxyuridine) staining in a mouse model in vivo involves identifying and addressing potential issues that could affect the success of the experiment. EdU is a thymidine analog used in the detection of DNA synthesis, making it a valuable tool for studying cell proliferation. Here are some common issues and potential solutions:

If problems persist, consider consulting with colleagues who have experience with EdU staining or reaching out to technical support from the product suppliers. Each experiment can have unique challenges, so sometimes a bit of experimentation is needed to optimize the conditions for your specific setup.

l Take a look at this protocol list; it could assist in understanding and solving the problem.

Not sure if you can get MFI for subgroups but if so you could do a one way ANOVA of post - pre MFI across groups.

Hello Maryam Soltani Asl

I agree with Soumyadeep Mukherjee . You cannot freeze the tumor tissue for flow cytometry analysis. You may cut out many pieces of the tumor tissue and transfer them to a 50 ml tube containing MACS buffer (1X PBS, 0.5% FBS, 2 mM EDTA) for subsequent flow cytometry analysis. You may store the tube at 4°C overnight or use the tissue for flow cytometric staining immediately following digestion into a cell suspension.

There are several digestion methods available. Ensure that you choose the appropriate method (namely, that method which will preserve cell surface protein expression, viability, etc) for your downstream application. After having performed the right digestion step, filter the tumor cell suspension through a 70µm cell strainer. Centrifuge cell suspension at 250 x g, 4 °C, 5 min. Resuspend cell pellet in 1 ml FACS buffer (containing 1-5% FBS in PBS along with approximately 1-5mM EDTA). It is important that you work fast and under cold condition. Count cells using a hemocytometer. Then proceed with the flow steps.

Best.

Hi Wey,

A great way to practice with flow is to use simple stains like Annexin V and PI (measures of early and late apoptosis). The staining protocol is only about 30 minutes as you don't wash out the stain. You can purposely spike in dead cells to make sure you have positive and negative populations. Another suggestion would be to mix two cell lines with different size/morphology so you can practice gating with FSC/SSC.

For beads BD makes great reagents all the way around. I also like ThermoFishers Ultra comp beads (01-3333-41) which would allow you to use the same colors you'll be using in the future.

Best of luck!

Hello Lucas,

Use HBSS (Hanks' Balanced Salt Solution) with calcium and magnesium, which is a balanced salt solution with trace amounts of glucose.

Cell viability is not compromised while cells are in HBSS since it is a balanced salt solution with glucose and it maintains osmolality and physiological pH. Cells will remain relatively happy and survive for at least 2 hours in HBSS.

So, you may use HBSS instead of PBS, and HBSS can also be used in flow cytometry.

Best.

I think it would be OK to compare the fluorescence intensities before and after treatment of the same type of cell, but it would be more difficult to compare different cell types because the cell types could differ in important parameters such as size, volume, and internal composition.

Is the mean intensity of a fluorescent signal a reasonable measure for relative quantification? Yes, but you should also show the distributions, since a heterogeneous distribution could lead to misleading results if only the mean were presented.

Hello again Henry, Just occurred to me - have you titrated the live/dead fixable stain or using it at the manufacturers recommended concentration? I have found you often(almost always!) find these concentrations will simply be so high that they appear to stain everything. If so try titrating 1:10, 1:100 and 1:1000 to get an idea of actual concentration required for optimal discrimination. These dyes are amide binding dyes not DNA dyes, designed to covalently bind to intracellular amines. However there are surface amines, too much dye availability may saturate everything or lead you to interpret live material as dead be reference to debris rather than live cells. Is your 10% 'live' cellular material suprisingly low FSC material? Can you upload a plot(s) of FSC versus ef780 staining? Sorry I missed this in the last message!

Sorry Maya for getting this but it is normal in the research filed. Did you check all the buffer components used and the cell count as well because these technical issues are important to get accurate results. If you still have issue, I recommend you ask the FACS expert from the FACS department in your research institute and he can fix the problem with you. Good luck.

Sudeep Kumar Maurya Thank you so much!

If this is for flow cytometry staining, count all the cells! Disregard any difference in cell type.

Flow cytometry data analysis can benefit from various statistical and bioinformatics tools, and the choice of tool depends on the specific goals of your analysis. In your case, where you want to identify the differential expression of meta clusters generated from FLOWSOM in 18 samples across 4 groups, the selection of an appropriate tool can be crucial. Here's an overview of the tools you mentioned:

For your specific analysis of flow cytometry data to identify differential expression of meta clusters generated by FLOWSOM, CITRUS is a tool specifically designed for this purpose. It integrates well with flow cytometry data and is user-friendly, making it a suitable choice. However, you may also consider using edgeR if you prefer an approach that you are more familiar with and are willing to adapt it for your flow cytometry data.

Ultimately, the choice of the tool should align with your expertise, the nature of your data, and your specific research goals. It's a good practice to explore multiple tools and potentially consult with colleagues or bioinformatics experts to ensure the most appropriate and reliable analysis for your project.

Hello Alex Peh

Yes, it will. The dead cells present in your sample can greatly affect your staining and therefore the quality of your data. The dead cells have greater autofluorescence than living cells and will increase non-specific antibody binding, thereby leading to false positives. This may make the identification of weakly positive samples and rare populations difficult. Moreover, cell clumping may occur due to the release of DNA from dead cells which is sticky in nature. Thus, identifying and removing data points representing dead cells is a critical step in ensuring accurate results.

There are dyes available which can distinguish live from dead cells. Depending on the fluorescent markers used, either PI, 7AAD or DAPI may be added to cells just prior to analysis. Only dead cells will take up these dyes. A very small amount of dye may be used to check viability and will not be detrimental to the cells. As little as 0.5 - 1.0 ug/ml of the dye may be added before the start of acquisition. However, if your sample is fixed, these dyes will not work. You will need a fixable live/dead marker. These nuclei acid binding dyes, when bound to double stranded nucleic acid will fluoresce. The dead cells can be identified and removed from the final analysis by gating on the unstained population (live cells).

Best.

Hello Shuo Wang

No, you cannot use non-conjugated secondary antibody in flow. Flow cytometry can be performed in two ways,

1) directly, using conjugated primary antibody or

2) indirectly, using a conjugated secondary antibody to bind an unconjugated primary antibody.

The indirect method would be more appropriate as indirect flow cytometry allows the choice of a wide range of probe molecules, enabling the user to match the desired probe with any primary antibody. Secondary antibody conjugates can improve a flow cytometry experiment by preserving the active site of the primary antibody, and by signal amplification.

In flow cytometry, a laser light is involved. Therefore, fluorescent detection with a secondary antibody conjugated to a fluorochrome is the requirement.

Best.

Hello,

It's normal to have a certain number of doublets or even triplets in FC. If you think you have above average doublets events on your SSC-A/SSC-H or FSC-A/FSC-H graphs, you can try to adjust your tryspinisation protocol to make it harsher (longer incubation time, higher trypsin concentration).

I don't think you will be able to tell the difference between doublets and syncytial cells on the SSC-A/SSC-H graph.

I'm not familiar with the protocol, but my suggestion is to make additions of BODIPY to the wells at time intervals corresponding to the amount of time it takes to analyze the samples, so that all samples are treated with BODIPY for the same length of time.

Hi,

This is a very simple and effective method of Kupffer cell isolation:

Good luck!

Online training e.g.

and many more, a google search will lead you to several other training sources

Morning Argha, propidium is being used as a viability dye and/or evaluate DNA fragmentation @ 1-2µg/ml(final dilution) in combination with AnnexinV. Similar concentrations can be used for DAPI and Hoechst to allow combinations of other AnnexinV conjugates. While 7-AAD is generally used a higher concentration(5µg/ml) and requires longer incubation.

Best wishes

It is not recommended to place an ice bucket directly on the collection tube place in a FACS BD Aria machine during cell sorting. The collection tubes are designed to be kept at a specific temperature, typically room temperature or slightly warmer, to maintain the viability and functionality of the sorted cells. Placing an ice bucket nearby could cause the temperature of the collection tubes to drop significantly, which may negatively impact the cells' health and sorting efficiency.

Instead, you can consider placing the ice bucket elsewhere in the workspace, such as on a separate table or shelf, and transferring the sorted cells to the ice bucket after they have been collected. This will help keep the cells at the appropriate temperature while being sorted, and then rapidly cool them down after sorting to prevent any potential damage from heat stress.

It's also worth noting that some FACS machines, including the BD Aria, come equipped with built-in cooling systems that can regulate the temperature of the collection tubes. In these cases, it may not be necessary to use an external ice bucket at all. Be sure to consult your machine's user manual or manufacturer's instructions

All the best

Waqas Nawaz Quantifying cytoplasmic versus nuclear localization of proteins within the same cell using flow cytometry typically involves a process known as intracellular staining and flow cytometry analysis. Here are the general steps to perform this quantification:

It's essential to optimize each step of the protocol for your specific experimental conditions, including the choice of antibodies, fixation, and permeabilization buffers. Additionally, controls and validation experiments should be included to ensure the accuracy of your quantification.

While the Beckman Coulter article you mentioned may not have provided the buffer details, you can try reaching out to other researchers in the field or your institution's core facilities for guidance on suitable permeabilization buffers and protocols. Collaboration with experts in flow cytometry can also be valuable for developing a robust assay.

Testing CD4 and CD8 T cell responses in splenocytes using flow cytometry involves a series of steps to prepare, stain, and analyze the cells. Below is a detailed protocol for this procedure:

**Materials:**

1. Splenocytes (isolated from mouse spleen)

2. Appropriate cell culture medium (e.g., RPMI-1640)

3. Fetal bovine serum (FBS)

4. Penicillin-streptomycin solution

5. Phosphate-buffered saline (PBS)

6. Red blood cell lysis buffer (if working with mouse splenocytes)

7. CD4 and CD8 fluorescent antibodies (fluorochrome-conjugated)

8. Cell surface marker antibodies (e.g., CD3, CD45)

9. Fixation and permeabilization buffer (e.g., BD Cytofix/Cytoperm)

10. Intracellular cytokine staining antibodies (e.g., IFN-γ, IL-2, TNF-α)

11. Appropriate isotype control antibodies

12. Flow cytometry tubes or plates

13. Flow cytometer equipped with appropriate lasers and filters

14. Software for flow cytometry data analysis

**Procedure:**

1. **Isolation and Preparation of Splenocytes:**

a. Sacrifice the experimental animals or obtain spleen tissue from human donors following ethical guidelines.

b. Isolate splenocytes using standard procedures. If working with mouse splenocytes, use red blood cell lysis buffer to remove erythrocytes.

c. Wash the isolated splenocytes with PBS and resuspend in culture medium (e.g., RPMI-1640) supplemented with FBS (10%) and penicillin-streptomycin (1%).

2. **Stimulation and Activation:**

a. Plate the splenocytes at a density of 1-2 million cells per well in a 96-well plate.

b. Add a T cell activation cocktail that includes an appropriate mitogen or antigen (e.g., PMA and ionomycin) and a protein transport inhibitor (e.g., GolgiStop) to prevent cytokine secretion.

c. Incubate the cells for 4-6 hours at 37°C in a CO2 incubator to allow for T cell activation and cytokine production.

3. **Surface Staining:**

a. Wash the activated splenocytes with PBS to remove the activation cocktail.

b. Incubate the cells with fluorescently labeled antibodies against surface markers, including CD4, CD8, CD3, and CD45, according to the manufacturer's instructions. Use appropriate isotype control antibodies as controls.

c. Incubate for 30 minutes in the dark at 4°C.

d. Wash the cells with PBS to remove unbound antibodies.

4. **Fixation and Permeabilization:**

a. Fix and permeabilize the stained cells using a fixation and permeabilization buffer according to the manufacturer's protocol. This step is necessary for intracellular staining of cytokines.

5. **Intracellular Staining for Cytokines:**

a. Incubate the fixed and permeabilized cells with fluorescently labeled antibodies against intracellular cytokines (e.g., IFN-γ, IL-2, TNF-α) in the dark at 4°C.

b. Incubate for 30 minutes, following the manufacturer's recommendations.

6. **Flow Cytometry Analysis:**

a. Analyze the stained splenocytes on a flow cytometer equipped with appropriate lasers and filters.

b. Gate on lymphocytes based on forward and side scatter characteristics.

c. Analyze CD4 and CD8 T cell responses by examining the expression of surface markers and intracellular cytokines.

d. Collect data for a sufficient number of events (e.g., 10,000-50,000 events) for statistical analysis.

7. **Data Analysis:**

a. Use flow cytometry analysis software to quantitate CD4 and CD8 T cell responses, including the percentage of positive cells and mean fluorescence intensity (MFI) of cytokine expression.

b. Compare experimental samples to appropriate controls (e.g., unstimulated cells, isotype controls) to determine specific T cell responses.

8. **Interpretation:**

a. Interpret the flow cytometry data to assess CD4 and CD8 T cell responses based on the expression of surface markers and intracellular cytokines.

9. **Reporting:**

a. Compile and report the results, including the percentages and MFI values for CD4 and CD8 T cell responses, as well as any relevant statistical analyses.

Remember to follow biosafety and ethical guidelines throughout the procedure, and adjust the protocol as needed based on the specific requirements of your experiment and the flow cytometry equipment available in your laboratory.

This depends on which cytometry (brand, model) you are using.

When analyzing mitochondrial staining data by flow cytometry, the gating strategy can vary depending on the specific experiment and the markers used. Here are some general guidelines for gating in mitochondrial flow cytometry experiments:

Single-cell gating: Start by gating on single cells to exclude debris and cell aggregates. This can be done using forward scatter (FSC) and side scatter (SSC) parameters.

Mitochondrial mass: To analyze changes in mitochondrial mass, you can use a mitochondrial dye such as MitoTracker Green or MitoTracker Deep Red. Gate on the population with high fluorescence intensity for the mitochondrial dye to select cells with intact mitochondria

Mitochondrial membrane potential: To assess changes in mitochondrial membrane potential, you can use a dye such as tetramethylrhodamine, ethyl ester (TMRE) or JC-1. Gate on the population with high fluorescence intensity for the mitochondrial membrane potential dye to select cells with functional mitochondria

Superoxide production: To measure mitochondrial superoxide (ROS) production, you can use a dye such as MitoSOX Red. Gate on the population with high fluorescence intensity for the MitoSOX Red dye to select cells with increased ROS production

Mitochondrial DNA content: To study mitochondrial heterogeneity and mitochondrial DNA dynamics, a nanoscale, multi-parametric flow cytometry-based platform was used. In this case, mouse liver homogenates were prepared and MTG-labeled

Mitochondrial ROS production under hypoxia: In this case, a gate was drawn to exclude mitoSOX-negative cells and include mitoSOX-positive cells. This gating was then applied to any sample by drawing an icon of the "gate" on the corresponding plot

Dear Sergei Rozhkov ,

CD25 is usually not expressed on Jurket cells.

Please find attached DOI of an article, explaining the production of CD25 on Jurket cells.

10.1134/S1990519X15050119

Hi Sergi Betriu Méndez

I briefly wanted to reach out concerning your inquiry.

For a detailed list of flow cytometry agents for the characterization of activation markers on the cell surface please consider the following paper from my old boss.

For quick insights into activation markers for cytotoxic T cells and a basic flow cytometry protocol, please consider:

For a comprehensive cytokine analysis (48plex with absolute quantification) use:

And for a detailed proteomic analysis (using cell culture supernatants or serum (prior to incubation)) please consider the T96 INF, IR and IO panels (albeit with relative quantification of expression) as per:

and more specifically used in the following papers:

I hope that helps.

All the best & good luck with your experiments,

Michael

Felipe Andrés Cordero da Luz thanks anyway. I'll keep trying

Dear Ana,

As Dr Sudeep stated, the blockinf step os importante tô reduce unspesific ligations. However, more importante in some cases os the use of isotype controls/the secondary antibody that could adress unspesific internacional with the fluorophore. Especially, tandem fluorophores can be metabolized by some cells (DOI: 10.1002/cyto.a.20774.)

There are 3 options for polymer dye buffers:

1. Super bright staining buffer

2. Brilliant stain buffer

3. Brilliant stain buffer plus (more concentrated version, which requires less volume)

One of these buffers should be used when using 2 or more polymer dyes to prevent dye-dye interactions. These buffers don't need to be included in single stain controls. Most compensation issues are due to poor single stains. If you have dim single stains on cells you should consider compensation beads.

I hope this helps. Happy flowing!

Thank you Xavier and Souheib for your answers!

Hello Malihe Naderi ! Sure thing, let's get straight to it:

Hi Ellie Alberts

I briefly wanted to reach out concerning your inquiry. There are a couple of options for your experiment.

For murine cells you may consider IL4 + aCD40 + aIgM (for BCR crosslinking). However, I would consider switching IL4 out for IL21 (as this seems to work irrespective, if you take mouse or human cells).

For details please see:

And for details on plasma cell characterization please consider:

And for a more general overview with more details, please have a look at this excellent review by Leen Moens and Stuart Tangye. It's a bit older, but key references especially for experimental protocols are included.

All the best & good luck for your experiments,

Michael

Thanks

Hi Junaid Ur Rehman

You can use a combination of both NGS and BSA (5%), as it would help in providing more effective blocking when compared to using them alone.

BSA and NGS can bind to the Fc region of IgG, as they have different properties that can complement each other. BSA is a small protein that is relatively easy to penetrate into cells, while NGS is a larger protein that can more effectively block Fc gamma receptors on the cell surface.

Thanks,

Based on the information you've provided, it seems likely that the lower concentration of cells observed in the PI stained samples after treatment with an antimicrobial is indeed due to cell lysis and subsequent washing steps. Let's break down the possible reasons for this:

Putting it all together, after the antimicrobial treatment and washing steps, the lysed cells release their DNA into the medium. When you stain the samples with PI, it primarily stains the extracellular DNA, as it can no longer penetrate the intact membranes of viable cells. Since the PI is staining mostly the released DNA from lysed cells and not the intact intracellular DNA of viable cells, the observed concentration of PI-positive cells will be lower in the treated samples.

To verify this explanation and further interpret your results, you may want to consider the following:

All the best buddy

It depends on how compact they are. I would add Trypsin to the wells and put them in the incubator for 10 min. Then, pipet them gently to obtain single cells. If The spheroids are still dense after pipetting, I would repeat the steps (incubating and pipetting) 2 or 3 more times until I get single cells.

Bets,

Hello,

Sending us an example of your peaks would be great. I you get only one peak, I think that your staining didn't work, all your cells are negative. Do you have a compensation control Alexafluor488 on compensation beads that shows both negative and positive peaks?

After acquiring single cell suspension and Fc block, I usually stain using biotinylated-antibody together with other conjugated primary antibodies in one master mix for 30min at 4C, followed by wash with FACS buffer once, and then stain for conjugated streptavidin alone for another 30min, followed by wash twice with FACS buffer. The cells are ready to be analyzed.

Hi Joshua Torres , have you tried using a spectral viewer to design a panel?

The manufacturer of your flow cytometer will have a spectral viewer. For example, Beckman have this: https://www.beckman.com/flow-cytometry/fluorescence-spectrum-analyzer

It allows you to select your cytometer, laser configuration, and bandpass filters. Then, you can use manufacturers websites (do they produce a fluorochrome that is used for your marker and laser of choice) to design a panel.

Brilliant Violet is excited around 405nm (violet laser) but emits light at ~710nm. PerCP-Cy5.5 can be excited on violet, blue, yellow and red lasers, but peak emission is ~675nm. There will be significant overlap of emitted light between brilliant violet 711 and PerCP-Cy5.5.

Alexa Fluor 700 and APC are both excited by the red laser (more so for APC) and there will be overlap in emitted light.

Hope this helps!

There are many packages that can be used for cytometry data. The most common are flowCore, flowStats, openCyto, and flowClust. They provide a comprehensive set of tools for preprocessing, analyzing, visualizing, and interpreting flow cytometry data. Though I didn't use any of them but my suggestions are based on some literatures of cells studies.

Using unstained cells is probably not a good idea for fixable viability dyes since, as you said, there is usually some degree of background staining on live cells.

I would recommend generating some dead cells, staining them with the viability dye and setting the gate based on that. In other words set the gate below the dead cell population. You can also kill some cells, mix them with freshly gathered (i.e. presumably alive) cells in a 1:1 ration, then stain that, so you should have 2 clear live and dead populations that you can extrapolate to your actual samples.

I do this by heating some cells in media or PBS for ~5 minutes at 65 C. Then I proceed with washing, viability dye staining and fixing (the fixing may be a bit redundant but it keeps them consistent with the other samples).

If you have a cytotoxic compound handy (like staurosporine) this could also be used.

I have attached an example of viability gates, in panel C you see the analysis of cells killed by a toxic compound, then stained with the dye.

Hi Danielle Fernandes Durso

I briefly wanted to reach out concerning your inquiry. As you haven't specified, if you want to use human or mouse B cells, please use the following protocol for human B cells with Miltenyi reagents (which always worked very well in my hands.

For mouse cells, please use the flow cytometry protocol outlined here:

with the following BD reagent.

I hope that helps! Good luck with your experiments.

All the best & kind regards,

Michael

Hi Shahrooz Nasrollahi Shirazi

I briefly wanted to reach out concerning your inquiry.

There are a couple of things that you could try.

1. I would cut down the PMA+ionomycin restimulation period (to 4hours or less) and possibly titrate the reagents as they are cytotoxic and do impede de-novo synthesis of proteins (including cytokines). For ideal concentrations for re-stimulation, please consider;

2. Please check the sequence and possibly methylation of your CPG as both may influence cytokine production (especially when pro- and anti-inflammatory cytokines are simultaneously produced). For details, please consider the following papers (I only found a reference for Tregs, but would assume that the same may hold true for in vitro induced Bregs)

3. Elongate you culture time to include a 96 hours (as 24 hours may double the fraction of IL10⁺ cells) time point possibly skipping the 24 and 48 hour time point as per;

4. Possibly include additional mitogens like anti-CD40, anti-IgM or (what I would specifically recommend) R848 (resiquimod), which is a TLR7/8 agonist as per;

I hope one or a combination of these considerations will help with your experiment.

All the best & good luck,

Michael

Shaswati Athaiya may be you can give accutase a try.

A.B Bayazid

Perfect! And it has worked well in your own experiments?

To remove axis labels from copied graphics in Flowjo, you can follow these steps:

By enabling the "Hide labels" option in the Flowjo preferences, the axis labels should be removed from the copied image. This allows you to add your own labels in PowerPoint without the need to trim off the default labels.

If you have previously achieved this and are unable to reproduce it now, it's possible that the preference settings might have changed or there could be other factors affecting the copied image. Make sure to double-check the preferences and ensure that the "Hide labels" option is enabled before copying the image.

CytoNORM is a normalization method for flow cytometry data that is implemented in OMIQ, a software tool for the analysis of high-dimensional cytometry data. CytoNORM is designed to correct for technical variation between samples, such as differences in staining intensity, instrument settings, and instrument performance.

The cytoNORM normalization method is based on the assumption that the majority of cell populations in a sample have similar staining properties and that any differences in staining are due to technical variability. CytoNORM uses a reference sample to calculate normalization factors that adjust for technical variability across all samples in the experiment.

The use of cytoNORM in OMIQ for the normalization of flow cytometry data can help to reduce technical variability and improve the accuracy and reproducibility of flow cytometry experiments. However, the effectiveness of cytoNORM may depend on specific experimental conditions, such as the type of samples being analyzed and the quality of the data. It is important to evaluate the effectiveness of any normalization method, including cytoNORM, on a case-by-case basis.