r/flowcytometry

Hi Everyone,

II’m running into something confusing with my flow data and hoping someone here has seen this before.

I’ve been seeing that, for the same bacterial spore sample, under different staining conditions (unstained, PI only, Syto 9 only, and PI-Syto9 double stain), the FSC:SSC profile changes depending on the stain I use.

https://preview.redd.it/rqyag0zo6jbh1.png?width=2136&format=png&auto=webp&s=6ddc50eb69277d615e54a7a21274c732c521437d

I'm thinking, would it be because the stain changed the refractive index of spores? is this something normal?

Would love to hear from people. Thanks in advance!

Hi everyone, I am setting up an overnight co-culture cytotoxicity assay using strongly adherent target cells (solid tumor models) and suspension effector cells. My final readout is flow cytometry (FACS), using a cell tracker dye to gate the target population and a viability dye (like 7-AAD/PI) to measure specific cell death. The issue: After the overnight incubation, the target cells are tightly attached to the bottom of the flat-bottom plate. Standard mechanical detachment (pipetting) is not effective enough—cells stay stuck, and I risk physically destroying the fragile, dying target cells. On the other hand, I want to avoid standard strong trypsinization at the end of the assay because I am afraid it will completely lyse the compromised target cells or interfere with the viability readout. Has anyone optimized a gentle detachment protocol for very sticky cells in a similar flow cytometry setup? Do you use alternative, gentler dissociation buffers (like Accutase or specific non-enzymatic buffers)? Does a cold shock help? Or is it generally better to just perform the entire overnight co-culture in round-bottom (U-bottom) / ultra-low attachment plates to avoid the attachment issue entirely? Thanks in advance for any tips or methodological advice!

What are some tips, tricks, or best practices y'all use to maximize post-sort recovery? MY core has had some users complain about low recovery recently, but we aren't doing anything differently than we always have. I wanted to see how y'all keep yours up. Also, how high do y'all shoot for? I've heard some folks say 50% is the best you can do, while I've heard others say anything less than 90% is unacceptable. Thoughts?

I'm building a 32 colour panel that is an extension of an existing 27 colour panel that has already been optimised. However I'm going to need to use Fc Block which wasn't needed on the original 27 colour panel.

In the past I've put on Fc block for 10 mins then added antibody mix directly onto cells from that. My concern in this case is that the final staining volume is going to be higher than in the original panel. I'd quite like to avoid having to re-titrate the other 27 antibodies so wondered what people's experience is changing that staining volume e.g. from 50 to 75uL for the same amount of antibody?

Maybe I just need to test it and see...

Thanks!

Hi everyone, I'm currently running a 2nd biological replicate using an 11-color spectral flow cytometry panel on leukemia cell lines to track surface receptor density during forced differentiation. Everything is going fine, but I have hit a hard wall with my CD-135 (FLT3) inventory.
I have exactly enough CD-135 left for my remaining 80 samples if I use 2.5 µL per sample with absolutely zero waste. Because I normally use a 10% volume overage for my master mixes (to account for pipette dead volume), including CD-135 in the daily cocktail means I will mathematically run out before the end of the project. I can't order a replacement in time.

My proposed workaround:
For today’s run (16 tubes total):

1. I calculate and mix the 10% overage cocktail for the other 7 antibodies + Immunofluorescence Buffer, completely omitting the CD-135.

2. I pipette 47.5 µL of this CD-135-free master mix into each cell pellet

3. I take a P10 pipette and manually spike exactly 2.5 µL of pure CD-135 directly into each of the 16 individual tubes, changing tips every time and striking off the tubes to ensure I don't miss any.

My questions for the experts:

1. Will manually spiking this one antibody alter the binding kinetics or MFI reproducibility compared to a fully pre-mixed cocktail, assuming the final volume (50 µL) and incubation time (45 mins at room temp in the dark) remain identical?

2. Has anyone else resorted to this manual spiking method to survive a low-reagent situation, and did it skew your data?

I know it increases the risk of technical pipetting error, but it seems like the only mathematically sound way to get to the finish line. Any advice or validation would be hugely appreciated!

Hi everyone,

I'm optimizing a flow cytometry protocol using mouse cervical lymph nodes from a disease model. Our goal is to identify the immune cell populations present and determine which cells are associated with dextran-coated iron nanoparticles injected into the mice.

As a test, we used snap-frozen lymph nodes, but after dissociation the cells were almost all dead and the staining was very poor. I'm trying to figure out whether snap-freezing is simply not suitable for preserving viable immune cells or if something else may have gone wrong.

Ideally, I'd like to preserve the tissue for a few weeks before processing.

Does anyone have recommendations for the best way to do this while maintaining good cell viability?

Would you freeze the intact lymph nodes or prepare a single-cell suspension first? Budget-friendly options (e.g., homemade cryopreservation media) would be especially appreciated.

Side question: We also need to detect the dextran-coated iron nanoparticles by flow. The only reagent I've found so far is an FITC-conjugated anti-dextran antibody. If anyone has experience detecting these nanoparticles or has alternative approaches, I'd love to hear your suggestions.

Thanks in advance!

Hello!

Does anyone know the actual flow velocity for a BD FACS ARIA II system (or similar sorting system). Does anyone know what the minimum you could reasonably achieve by lowing the sheath pressure is? This is for considering actual illumination time of the cell.

I can make an estimate of ~2 m/s by using the 10 micro litres per minute quoted for flow rate 1 and and the roughly 10 micron diameter core I could see with the microscope.

Thanks very much!

Hey folks,

I've got cells expressing a fluorescent protein. Am getting a bizarre expression pattern I've never seen before:

Sample showing weird expression.

We want to quantify how many cells are expressing the fluorescent protein. I've done this hundreds of times. But I've never seen a treatment which causes an expression pattern like this, where the negative(?) population shifts so deeply into the positive gate.

See negative control, for reference:

Negative control with no mCherry.

Meanwhile, our positive control exemplifies the expression I'd expect to see:

Positive control showing typical expression.

Wondering the flow community has any advice? I've repeated this experiment a few times and it's all similar (I've deliberately chosen the most stark, though). Our cytometry techie took a quick look and found no issue. We're getting to the stage where the PI's are thinking up future research based off this finding, but something about the way this looks just isn't sitting right with me...

Any opinions appreciated,

King_of_Mints

P.S. There is technically some GFP expression in the sample as well, but it is incredibly insignificant (<0.5%)!

HI. This is gated among single live cells of human PBMC (patients). 1st image PBMCs are thawed, stained and passed on a cytometer. 2nd, same staining after 24h culture (1M/ml if RPMI 10% FBS 1% P/S). Culture is done in 12 well plates. Cells are recovered and then the bottom is scraped to recover attached cells.

I have one population after thawing, but which is clearly composed of high CD33 and high CD14 cells. After culture, I have 2 distinct populations. The high ones and the one with 1D14 very low/negative. Both populations express CD11c and HLA-DR, are negative for CD3, CD56, CD19 , CD20.

When activated with IFNg, only the CD14high CD33 up regulate PD-L1, the CD14 low cells have keep the basal level (almost no expression).

I don't know much about myeloid cells. What is the population that becomes CD14 very low? Can it be an artefact?

Thank you for all the help:

https://preview.redd.it/zqvystx5cu9h1.jpg?width=681&format=pjpg&auto=webp&s=c64c0226646c7843bc6c5e3e8d28cfb548be67a5

I’m using a photoconvertible mouse model and the fluorescence is expressed on dendritic cells i.e. a pretty rare population. How can I get a good single stain for unmixing? There are no commercial beads.

Hi everyone, I'm the developer of Voxcyte. I came from a background in physiology before moving into data science and software engineering.

Voxcyte is an app that runs flow cytometry analysis natively on macOS. No browser, no cloud, no Java. Everything is local and your data never leaves your Mac.

The standout feature is a real-time interactive 3D scatter plot you can rotate and zoom, staying smooth across millions of events. Most tools either can't do 3D like this or lag on large files. It's built for Apple Silicon, so most actions are instant and even heavy operations on large files only take a few seconds.

Voxcyte is a complete analysis app including compensation, gating, 2D/3D/histogram plots, statistics, and publication-ready figure exports. It's also more affordable than some established tools, and you can try it with a 14-day free trial, no card.

I take feedback seriously and act on it fast, so if something doesn't fit your workflow or you run into an issue, just contact me and it will likely be in the next release.

 

Hey everyone,

I am encountering a weird problem with my FACS panel for human PBMCs and hope for some insights.

Last week, I stained the panel and it worked well. However, when I stained today, neither in my single-stained cells nor in my fully stained sample was there any signal for CD4 and CD8. All other markers (CCR7, CD95, CD45, CD45RO, CD28, etc.) worked, and I used the same antibody vials from my rack as last week. However I used a different healthy control.

In my protocol, I am staining the cells with some extracellular markers that I wasn't able to get working nicely post-fixation (CD95, CD28, CCR7), stimulate the cells for 15 mins with cytokines, then immediately fix with BD Phosflow Lyse/Fix Buffer, followed by methanol permeabilization and intracellular staining (looking for several pSTATs) and some other extracellular markers.

I am aware of problems arising from staining fixed epitopes, as well as effects of methanol fixation on fluorophores, which is why I thought I had arrived at a good compromise and found a good panel.

Have you encountered the issue that staining of some epitopes post-fixation is also dependent on the donor? Or do you have any other idea why this might be? It wouldn't be a problem to change the clone, as we have some more options on hand, but then I also don't know whether this new clone would be reliable. I was hesitant to stain more markers than necessary before the cells are fixed as I am concerned, this could cause some phosphorylation events by activating agonist signaling.

The antibodies I used were Anti-CD4 BUV496 (Clone SK3) and Anti-CD8a PE-Dazzle594 (Clone Hit8a). On the Thermo Fisher website, "Antibody clone performance following fixation/permeabilization," the CD4 clone is listed as one that should work post-fixation. The CD8 clone is not listed there, but it has worked post-fixation in the past for me, which is why I selected it. I also did a single stain for CD4 PE today with the clone RPA-T4, and this worked well.

I also should mention that I am quite new to flow cytometry.

I appreciate your input!

 

 

I am still a confused about compensation beads. I will be using FITC for CD31 and a LIVE/DEAD stain for some endothelial cells. Do I need compensation beads?

Hi everyone,

I am currently working on Aurora SpectoFlow and have 3 panels for the same tissue type. The first two panels are based on live samples while the third is on fixed samples. So far I have ran 11 samples with no issues, however I decided to re run my single control for 3 antibodies from beads to cells as I wasn't happy how it looked.

I was told to treat the beads same way I do cells but I don't think it produced good results so I switched to cells. I had all my panels exported just in case, but then I re ran my singles and now everything is looking weird. All events, singlets etc. When I re-ran my singles, alongside them I ran only one sample, and when it looked weird, I thought it was a one-off because the day before the sample looked fine in other panels.

Now, when I noticed the issue again, Aurora actually didn't have those 3 single controls saved, which I found weird. So I imported what I had and I think I might have imported the UNMIXED fsc file as it wouldn't let me re-unmix. But lets say I manage to import them back or worst case scenario I have to re run those controls..how does that affect the samples I have ran already. I understand that Aurora saves Raw and Unmixed data but I'm just very paranoid and wondering how it reflects down the line in FlowJo, if at all.

Thank you in advance!!

Hi Flow-People!

Have any of you worked with a modified protocol for the ThermoFisher EdU Click-iT kit to allow for staining in 96-well plates?

The current protocol requires staining in tubes to allow for 600ul of volume, which isn't feasible when working with 20+ samples.

Thanks!

I'm trying to help another department revive a long-idle Fortessa X-20, which sat untouched for >6 months and now doesn't work. It seems that sheath fluid makes it into the side panel, but not all the way to the flow cell. I can see that the flow cell is mostly full of air most of the time. I suspect salt crystals blocking sheath somewhere along the path, and am planning to replace all the narrow tubing and tiny T-connectors inside that I can reach. I've put some annotated pictures. Please hit me with suggestions. Eventually, I'll have to give up and tell them to call BD, but they're reluctant due to cost for an instrument that wasn't under contract and hasn't been touched in months. Also hoping someone can tell me whether sheath enters the flow cell through both the translucent teal and the dark blue tubing, and if not, what each of them delivers/removes from the flow cell.

https://preview.redd.it/h2zkm6nnxx8h1.jpg?width=1500&format=pjpg&auto=webp&s=a8fa816cfe3e7f6700150ce50cbb73af03d26b34

https://preview.redd.it/letzs7nnxx8h1.jpg?width=1500&format=pjpg&auto=webp&s=8c98cee29b3f1b39159df66c8817f39759ed8fe4

https://preview.redd.it/l9unx8nnxx8h1.jpg?width=1500&format=pjpg&auto=webp&s=30c9ce12067d7e4408ed1312373d9a04e367be5e

Hi everyone,

I am currently running a bacterial growth curve experiment (Faecalibacterium prausnitzii, a strict anaerobic Gram-positive bacterium) using SYTO 9 and Propidium Iodide (PI) for live/dead quantification on a Beckman Coulter CytoFLEX system.

I've hit a technical wall with template setting and gating, and I would appreciate any insights from the community.

Fig 1-a, b, c. Unstained and single-stained group
Fig 2. Sample acquisition (Red PI 70% EtOH gate is defined by the 2nd time single-stained acquisition)
Fig 3. Compensation adjustment of Fig 2 (FITC - 100% PE)

• Bact treatment: No permeabilization was performed on the bacteria. I only compared two different treatments for the PI dead-cell template: direct staining of intact cells versus 70% ethanol inactivation. The other are fresh bacteria at 0, 24, and 48 hours.
• Channel: FITC for SYTO9; PE for PI
• PMT Gains: FITC: ~180, PE: ~180 (Signal peak natively lands around 10^4 for stained samples).
• Threshold: Manual (FSC-A: 3500, FITC-A: 1000, AND logic) to clear medium debris.
• Dye Concentrations: 1.5 µM SYTO 9 + 10 µM PI (Wash-free protocol, as suggested by Beckman application notes for probiotics). Stained for 15 minutes.
• Compensation Matrix: Currently set to 0% for troubleshooting.

-With compensation completely turned off (0%), I ran three independent control groups, but their fluorescence profiles look identical

-When preparing the PI dead template (using 70% EtOH), the non-viable cluster drastically shifted rightward into the viable gate during a repeated run without any parameter changes. Has anyone experienced EtOH fixation causing such aggressive autofluorescence spectral shifts in Gram-positive anaerobes?

-In my 24-hour dual-stained samples, 99% of events remain in the viable region, but the population splits into two distinct dense sub-clusters (one at FITC 10^4 and an elongated one stretching to 10^5). FSC/SSC shows an elongated diagonal pattern. Is it safe to assume these are single cells vs. bacterial chains/doublets rather than distinct physiological states?

It's a big trouble for me, I can't figure out this machine... And anaerobic bacteria staining is also time-consuming. I would appreciate any advice. Thanks so much