r/flowcytometry

Hey, I am titrating FOXP3-AF647 (Clone 206D from Biolegend) to identify CD4+ regulatory T cells. Everything went well, but I noticed that there was a weird population extending from the negative population at the higher concentrations which disappeared at the 1 in 200 dilution and beyond. Wondering what it could be as the far right population (which I suspect are the CD4+ Tregs) remains consistently visible.

*NOTE - I hover around the population I am confused about in the video using my cursor.

https://reddit.com/link/1x2qynl/video/676c0nyrmpuh1/player

I am working on an Imiquimod-induced psoriasis-like inflammation model. I applied imiquimod on the shaved back skin of mice for 3 days and then removed the back skin. I washed the skin with PBS once and then cut it into tiny pieces. Then I digested the sample in 1.5 mL digestion buffer (PBS with 10% FCS, 0.2 ug/ml liberase, 0.5 ug/mL DNAse) at 37 degree celsisu for 70 minutes; then I stopped the reaction by adding 5 mL of FACS buffer. Counted the cells, and the viability was around 50-60%.

After that, I put the cells in the activation cocktail from BioLegend (1:500 dilution of stock) for 4 hours. Then I stained the cells for the viable+CD45+CD3+CD4+IL17A+ population. However, from the plot it's obvious that 90% of the cells died. I had an unboosted control without the cell activation cocktail. There, the viability was around 30%. However, there was no clear signal for IL17A ( as expected).

Can anyone suggest me what to do in this case to increase the viability and still detect IL17A-secreting cells?

Good morning. Our lab is going to get a new cytometer. We currently have a MACS Quant 16, whose software is a headache. Also, it’s very delicate and requires technical service. I’ve been offered cytometers from BD, Agilent, Attune, and Cytoflex. Could you recommend a brand? I’m looking for easy-to-use software and a very sturdy device that needs little maintenance from the manufacturer.

They are offering me the following cytometers:

Milteny MACQuant 16. 3 lasers, 16 fluorochromes.
CytoFlex LX 6 lasers, 21 fluorochromes.
Attune NxT AFC 3 lasers.
BD FACSymphony A1. 4 lasers, 19 fluorochromes.
Novocyte Quanta 4 lasers, 25 fluorochromes.

Thank you

Can't access or download anything

They need to put this stuff in s3 or something

Hi everyone! Thanks so much for all of your help :) I have started optimization for a large tetramer screen (18 x MHC-I specific CD8 tetramers) I had initially optimized the screen for 1 tetramer per well, but would like to increase to as many tetramers per well as possible. I had tried to multiplex this to 18 but unfortunately have a lot of what I suspect is nonspecific staining. :( (see images below) I have plans to titrate the PE/tetramer and try at room temp vs. on ice. Is there any other condition that I should try to optimize or does anyone have any tips? I am expecting very few tetramer positive cells in these conditions. Thank you! (also- the cd19 + population looks strange/nonexistant because this protocol requires B cell depletion.) TIA!

https://preview.redd.it/ibtpd5g104uh1.png?width=1980&format=png&auto=webp&s=a41e1f36d13e88b56766939dd389edf22e78d271

Hello, I'm new to flow and am having trouble understanding how to determine the dilution for each antibody. I read that we are supposed to do titration to determine the dilution but what if I have so many antibodies to test? I'm planning to stain 2E5 of human cells with 100ul. For example, if the manufacturer says 20ul per 1e6 cells in 100ul then for my case would it be 4ul instead?
Thanks a lot !!!

Hey, I have done a lot of reading on flow cytometry and OMIPs but I have seen very different ways researchers use to decide what dilution to use for their antibodies. There does not seem to be one correct way of doing things and some methods seem very impractical or generally not realistic (such as using the same cell population genetically engineered to not express the antibody's target🤦).

Please share suggestions below on how to titrate antibodies for markers that produce smears or plots whereby it is hard to identify the positive population.

During data acquisition, is a FMO a necessity for higher quality flow publications or is manual gating strategies an equal methodology?

https://preview.redd.it/7rs0ngybl3th1.png?width=2044&format=png&auto=webp&s=c705f74c0249c3dffe3327b72c4f94ddbde2062d

Hey all, I titrated my antibody from Thermofisher, ROR gamma T - PE (clone AFKJS-9). However, this antibody looks rather odd and would love some advice or confirmation if this is what is expected of such an antibody. Please

Context: these are done on fixed PBMCs, fixed using the FOXP3 ebioscience thermofisher kit, stained for 16 hours in the fridge (100 microlitres stain volume), intracellular blocking step for 15 minutes on ice using a mix containing 2% rat serum and 2% mouse serum.

The order is ROR gamma T FMO, 1 in 1000, 1 in 5000, 1 in 10000, 1 in 12500, 1 in 15000, 1 in 17500, 1 in 20000 and 1 in 25000.

The concatenated plots are done on CD3+ cells and CD3 plus CD4+ cells.

Hi,

I have a phosho flow panel largely to look at phospho-STATs, I usually try to do all staining and run on the same day but due to some cytometer rescheduling issues I need to run my samples the day following the harvest. My work flow is:

- Samples are B6 mouse spleen, largely focused on T cells.

- collect spleen, process, fixable viability dye and surface stain as usual using (to the best of my abilities) MeOH-compatible flurophores. Wash.

- Fix with BD CytoFix/CytoPerm, 30min RT. Wash.

- Methanol permeabilization with ice cold 100% Methanol, incubate 4C 30min. Wash well.

- Intracellular staining with anti-phospho STAT antibodies, in PBS/FACS buffer. Wash. Acquire on Cytek Aurora.

My question is, being that I need to have an overnight stopping point, which buffer/post fixation step is best?

The Options in thinking:

1. After BD CytoFix/Cytoperm, switch to PBS and leave at 4C. I'm assuming once this fix step is done, the phosphorylation events are fixed and stopping at this step wouldn't affect my pSTAT detection?

2. IN Methanol at -20C. Many protocols recommend this, but -- although my surface antibody fluorophores do OK with Methanol, they do lose quite a bit of brightness and I'm admittedly scared of leaving them in MeOH overnight. This is a 20+ color panel so couldn't just choose the most perfect MeOH-resistant fluorophores.

3. After methanol fix for 30min, wash well and leave in PBS at 4C. Just want to make sure that once I come back the next day, I would be able to go straight into Intracellular staining.

Thanks a ton

Hi everyone! I would like to ask for your help! I need to learn how to use a flow cytometer. My only problem is, that I don't have anyone on the department that can help. And the machine that I'm using is old. Like... they don't even produce it anymore old (partec sysmex ml). And the software that I should use is the same... Flowmax 3.2. Even google corrected me that I'm not looking for this.

So I'll have to measure cells between 15-40 micrometer. And I have no clue how to set up a gain for that or a lower threshold.

If anyone can help, I would be forever grateful for that!

I'm trying to enrich for cells that have stably integrated a transposon construct using FACS. Unfortunately, our construct has no fluorescent or other expression marker, so we're relying on two cell-surface markers, CD81 and CD47, that should be overexpressed following transposition. The problem is that both markers are already constitutively expressed in our target cells, and we don't know the transposition efficiency or how much their expression will increase following integration.

I've performed antibody titrations for both markers, but I'm unsure how best to determine the optimal staining conditions for sorting.

My understanding is that the usual goal of an antibody titration is to identify the lowest antibody concentration that provides good separation between negative and positive populations, with minimal background and sufficient staining of the positive population.

However, in my case, I'm not trying to distinguish truly negative cells from positive cells. Instead, I'm trying to distinguish cells with normal endogenous expression from cells with increased expression due to stable transposition.

Wouldn't it make more sense, in this case, to use an antibody concentration that preserves the differences in fluorescence intensity between cells with normal and elevated expression, rather than one that saturates the epitopes? If the antibody concentration is too high and staining approaches saturation, could that compress the fluorescence differences between normal and high-expressing cells and make them harder to distinguish?

Or am I misunderstanding this?

I'm also unsure how best to optimise the sorting strategy when I don't know the expected increase in CD81 or CD47 expression, or the proportion of cells that have successfully integrated the construct.

Since we're using a transposon system rather than lentiviral transduction, increasing the MOI isn't an option for improving separation.

I'd appreciate any advice, particularly from anyone who has experience sorting cells based on relatively small increases in endogenous cell-surface marker expression rather than using a distinct positive/negative marker.